M1/M2 Macrophages: the arginine fork in the road to health and disease

Mills, Charles Dudley

Lenz, Laurel L

Ley. Klaus

2015

Frontiers Media SA

Macrophages have unique and diverse functions necessary for survival. And, in humans (and other species), they are the most abundant leukocytes in tissues. The Innate functions of macrophages that are best known are their unusual ability to either "Kill" or "Repair". Since killing is a destructive process and repair is a constructive process, it was stupefying how one cell could exhibit these 2 polar – opposite functions. However, in the late 1980’s, it was shown that macrophages have a unique ability to enzymatically metabolize Arginine to Nitric Oxide (NO, a gaseous non – specific killer molecule) or to Ornithine (a precursor of polyamines and collagen for repair). The dual Arginine metabolic capacity of macrophages provided a functional explanation for their ability to kill or repair. Macrophages predominantly producing NO are called M1 and those producing Ornithine are called M2. M1 and M2 – dominant responses occur in lower vertebrates, and in T cell deficient vertebrates being directly driven by Damage and Pathogen Associated Molecular Patterns (DAMP and PAMP). Thus, M1 and M2 are Innate responses that protect the host without Adaptive Immunity. In turn, M1/M2 is supplanting previous models in which T cells were necessary to "activate" or "alternatively activate" macrophages (the Th1/Th2 paradigm). M1 and M2 macrophages were named such because of the additional key findings that these macrophages stimulate Th1 and Th2 – like responses, respectively. So, in addition to their unique ability to kill or repair, macrophages also govern Adaptive Immunity. All of the foregoing would be less important if M1 or M2 – dominant responses were not observed in disease. But, they are. The best example to date is the predominance of M2 macrophages in human tumors where they act like wound repair macrophages and actively promote growth. More generally, humans have become M2 – dominant because sanitation, antibiotics and vaccines have lessened M1 responses. And, M2 dominance seems the cause of ever - increasing allergies in developed countries. Obesity represents a new and different circumstance. Surfeit energy (e.g., lipoproteins) causes monocytes to become M1 dominant in the vessel walls causing plaques. Because M1 or M2 dominant responses are clearly causative in many modern diseases, there is great potential in developing the means to selectively stimulate (or inhibit) either M1 or M2 responses to kill or repair, or to stimulate Th1 or Th2 responses, depending on the circumstance. The contributions here are meant to describe diseases of M1 or M2 dominance, and promising new methodologies to modulate the fungible metabolic machinery of macrophages for better health.

Allergy and Immunology

Medicine (General)

English

9782889194995 (print)

16648714 (online)

M1/M2 MACROPHAGES: THE
ARGININE FORK IN THE ROAD
TO HEALTH AND DISEASE
Topic Editors
Charles Dudley Mills, Laurel L. Lenz
and Klaus Ley

IMMUNOLOGY

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ISSN 1664-8714
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ISBN 978-2-88919-308-0
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November
2014to| Health
Energyand
metabolism
March 2015    M1/M2 Macrophages: The Arginine Fork
in the Road
Disease  |  1

M1/M2 MACROPHAGES: THE
ARGININE FORK IN THE ROAD TO
HEALTH AND DISEASE
Topic Editors:
Charles Dudley Mills, BioMedical Consultants, USA
Laurel L. Lenz, University of Colorado School of Medicine, USA
Klaus Ley, La Jolla, Institute for Allergy & Immunology, USA

Macrophages have unique and diverse
functions necessary for survival. And,
in humans (and other species), they are
the most abundant leukocytes in tissues.
The Innate functions of macrophages
that are best known are their unusual
ability to either “Kill” or “Repair”. Since
killing is a destructive process and
repair is a constructive process, it was
stupefying how one cell could exhibit
these 2 polar – opposite functions.
However, in the late 1980’s, it was shown
that macrophages have a unique ability
The Fork in the Road to Health and Disease
to enzymatically metabolize Arginine
to Nitric Oxide (NO, a gaseous non –
specific killer molecule) or to Ornithine (a precursor of polyamines and collagen for repair).
The dual Arginine metabolic capacity of macrophages provided a functional explanation
for their ability to kill or repair. Macrophages predominantly producing NO are called M1
and those producing Ornithine are called M2. M1 and M2 – dominant responses occur
in lower vertebrates, and in T cell deficient vertebrates being directly driven by Damage
and Pathogen Associated Molecular Patterns (DAMP and PAMP). Thus, M1 and M2 are
Innate responses that protect the host without Adaptive Immunity. In turn, M1/M2 is
supplanting previous models in which T cells were necessary to “activate” or “alternatively
activate” macrophages (the Th1/Th2 paradigm). M1 and M2 macrophages were named such
because of the additional key findings that these macrophages stimulate Th1 and Th2 – like
responses, respectively. So, in addition to their unique ability to kill or repair, macrophages
also govern Adaptive Immunity. All of the foregoing would be less important if M1 or
M2 – dominant responses were not observed in disease. But, they are. The best example to

Frontiers in Immunology

March 2015    M1/M2 Macrophages: The Arginine Fork in the Road to Health and Disease    2

date is the predominance of M2 macrophages in human tumors where they act like wound
repair macrophages and actively promote growth. More generally, humans have become
M2 – dominant because sanitation, antibiotics and vaccines have lessened M1 responses.
And, M2 dominance seems the cause of ever - increasing allergies in developed countries.
Obesity represents a new and different circumstance. Surfeit energy (e.g., lipoproteins) causes
monocytes to become M1 dominant in the vessel walls causing plaques. Because M1 or M2
dominant responses are clearly causative in many modern diseases, there is great potential
in developing the means to selectively stimulate (or inhibit) either M1 or M2 responses to
kill or repair, or to stimulate Th1 or Th2 responses, depending on the circumstance. The
contributions here are meant to describe diseases of M1 or M2 dominance, and promising
new methodologies to modulate the fungible metabolic machinery of macrophages for
better health.

Frontiers in Immunology

March 2015    M1/M2 Macrophages: The Arginine Fork in the Road to Health and Disease    3

Table of Contents

Editorial
07


Macrophages at the fork in the road to health or disease
Charles D. Mills, Laurel L. Lenz and Klaus Ley

Introduction to Macrophage Biology and Functions
13

23

31

42

47

69

73


Metabolism via arginase or nitric oxide synthase: two competing arginine
pathways in macrophages
Meera Rath, Ingrid Müller, Pascale Kropf, Ellen I. Closs and Markus Munder
Evolution of innate immunity: clues from invertebrates via fish to mammals
Kurt Buchmann
Evolutionary roots of arginase expression and regulation
Jolanta Maria Dzik
Macrophage: SHIP of Immunity
Charles D. Mills, Anita C. Thomas, Laurel L. Lenz and Markus Munder
From monocytes to M1/M2 macrophages: phenotypical vs. functional
differentiation
Paola Italiani and Diana Boraschi
Spatial, temporal, and functional aspects of macrophages during “The Good,
the Bad, and the Ugly” phases of inflammation
Robert A. Harris
“Of mice and men”: arginine metabolism in macrophages
Anita C. Thomas and Joshua T. Mattila

Macrophage Influences in Different Diseases
A. Wounds and Cancer: Stark Examples of the Two-Edged Sword Nature of
Macrophage Responses
80

91

Rethinking regenerative medicine: a macrophage-centered approach
Bryan N. Brown, Brian M. Sicari and Stephen F. Badylak
Hepatic localization of macrophage phenotypes during fibrogenesis and
resolution of fibrosis in mice and humans

Leonie Beljaars, Marlies Schippers, Catharina Reker-Smit, Fernando O. Martinez,
Laura Helming, Klaas Poelstra and Barbro N. Melgert
102 Functional relationship between tumor-associated macrophages and
macrophage colony-stimulating factor as contributors to cancer progression

Damya Laoui, Eva Van Overmeire, Patrick De Baetselier, Jo A. Van Ginderachter and
Geert Raes

Frontiers in Immunology

March 2015    M1/M2 Macrophages: The Arginine Fork in the Road to Health and Disease    4

117 Depletion of tumor-associated macrophages slows the growth of chemically
induced mouse lung adenocarcinomas

Jason M. Fritz, Meredith A. Tennis, David J. Orlicky, Hao Yin, Cynthia Ju,
Elizabeth F. Redente, Kevin S. Choo, Taylor A. Staab, Ronald J. Bouchard,
Daniel T. Merrick, Alvin M. Malkinson and Lori D. Dwyer-Nield

B. Macrophages in Infections
128 Phenotypic diversity and emerging new tools to study macrophage activation
in bacterial infectious diseases

Mignane B. Ka, Aurélie Daumas, Julien Textoris and Jean-Louis Mege
135 The role of myeloid cell activation and arginine metabolism in the
pathogenesis of virus-induced diseases

Kristina S. Burrack and Thomas E. Morrison

C. Macrophage Responses in Atherosclerosis and Other Non-Pathogen-Induced
Inflammatory Conditions
147 Classical and alternative activation and metalloproteinase expression occurs
in foam cell macrophages in male and female ApoE null mice in the absence
of T
  and B lymphocytes

Elaine Mo Hayes, Aikaterini Tsaousi, Karina Di Gregoli, S. Rhiannon Jenkinson,
Andrew R. Bond, Jason L. Johnson, Laura Bevan, Anita C. Thomas and
Andrew C. Newby
163 Cholesterol efflux pathways regulate myelopoiesis: a potential link to altered
macrophage function in atherosclerosis

Andrew James Murphy, Dragana Dragoljevic and Alan Richard Tall
169 Dynamic aspects of macrophage polarization during atherosclerosis
progression and regression

Michael Peled and Edward A. Fisher
178 Functions of arginase isoforms in macrophage inflammatory responses:
impact on cardiovascular diseases and metabolic disorders

Zhihong Yang and Xiu-Fen Ming
188 Role of alveolar macrophages in chronic obstructive pulmonary disease

Ross Vlahos and Steven Bozinovski
195 Macrophage polarization in obesity and type 2 diabetes: weighing down our
understanding of macrophage function?

Michael James Kraakman, Andrew James Murphy, Karin Jandeleit-Dahm and
Hélène L. Kammoun
201 Are “resting” microglia more “M2”?

Jonathan D. Cherry, John A. Olschowka and M. Kerry O’Banion
205 M1/M2 macrophage polarity in normal and complicated pregnancy

Mary B. Brown, Maria von Chamier, Ayman B. Allam and Leticia Reyes
215 Ontogeny and polarization of macrophages in inflammation: blood monocytes
versus tissue macrophages

Adwitia Dey, Joselyn Allen and Pamela A. Hankey-Giblin

Frontiers in Immunology

March 2015    M1/M2 Macrophages: The Arginine Fork in the Road to Health and Disease    5

Regulation of Macrophage Differentiation and Functions
230 Molecular mechanisms that influence the macrophage M1–M2 polarization
balance

Nan Wang, Hongwei Liang and Ke Zen
239 SOCS proteins in macrophage polarization and function

Heather M. Wilson
244 Myeloid colony-stimulating factors as regulators of macrophage polarization

Thomas A. Hamilton, Chenyang Zhao, Paul G. Pavicic Jr. and Shyamasree Datta
250 Regulation of macrophage polarization by RON receptor tyrosine kinase
signaling

Amitabha Chaudhuri
255 Complement, C1q, and C1q-related molecules regulate macrophage
polarization

Suzanne S. Bohlson, Sean D. O’Conner, Holly Jo Hulsebus, Minh-Minh Ho and
Deborah A. Fraser
262 Purinergic and calcium signaling in macrophage function and plasticity

Bimal N. Desai and Norbert Leitinger
270 Nitric oxide synthase: non-canonical expression patterns

Joshua T. Mattila and Anita C. Thomas
275 Metabolic reprograming in macrophage polarization

Silvia Galván-Peña and Luke A. J. O’Neill

Frontiers in Immunology

March 2015    M1/M2 Macrophages: The Arginine Fork in the Road to Health and Disease    6

EDITORIAL
published: 16 February 2015
doi: 10.3389/fimmu.2015.00059

Macrophages at the fork in the road to health or disease
Charles D. Mills 1 *, Laurel L. Lenz 2 and Klaus Ley 3
1

BioMedical Consultants, Marine on St. Croix, MN, USA
Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA
3
La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA
*Correspondence: mills002@umn.edu
2

Edited and reviewed by:
Claudia Kemper, King’s College London, UK
Keywords: macrophage, innate immunity, M1, M2, wound, cancer, infection, atherosclerosis

Macrophages are the epicenter of all immune systems (1). The first
and the most abundant leukocyte observed (2), macrophage have
long been relegated to the role of “servants” of T or B cells/adaptive
immunity. This view is now known to be backward. Macrophages
necessarily initiate and direct virtually all immune responses from
simple multicellular animals to humans.
There is good news and bad news in the newly recognized
importance of macrophages/innate immunity. The well-known
“double-edged sword” nature of the immune system can largely be
attributed to macrophages’ unique ability to make polar-opposite
repair/heal (M2) or kill/inhibit (M1) type responses (3).
In health, M2-type macrophages maintain homeostasis by
helping repair and replace lost or effete cells. Ever-present in
tissues, macrophages are also the primary host defense against
pathogens (or altered self cells) because their unique physiology allows them to rapidly switch from their M2/heal mode to
an M1/inhibit mode: both powerful responses; both potentially
dangerous.
In disease, over expression of M2/heal macrophages contributes
to chronic infections, fibrosis, allergy, and cancer (3). Conversely,
M1/inhibit-dominant activity plays a major role in atherosclerosis,
autoimmunity, and other chronic inflammatory conditions.
Of fundamental importance is that both the routine M2/heal
and the induced M1/inhibit macrophage functions occur in all
animals whether they have T cells or not. Furthermore, M1 and
M2 macrophage responses play necessary roles in causing T cells
to make Th1 or Th2-type responses if pathogens or altered self are
present (4). Hence, the renaming of macrophage responses M1
and M2.
This new knowledge about the central role of macrophages
in immune systems brings great promise for increasing health/
decreasing disease. In this regard, the ability of macrophages to
exhibit the polar-opposite M2/heal and M1/kill functions result, in
part, from their unique ability to metabolize one amino acid – arginine – to either growth-promoting ornithine or growth-inhibiting
nitric oxide (NO) (5). Hence, the title of this Topic, “M1 and M2
Macrophages: The Arginine Fork in the Road to Health and Disease.” We hope that the articles assembled here help illuminate
the basic functions of macrophages referred to as SHIP [sample,
heal, inhibit, and present (antigen)]. Such knowledge is critical for developing the means to modulate the direct M2/heal or
M1/inhibit responses of macrophages, or their indirect abilities to
initiate and direct T and B cell responses. One can properly say
macrophages are the “chicken and the egg” of immunity (1).

www.frontiersin.org

ORIGIN OF M1/INHIBIT AND M2/HEAL MACROPHAGES AND
THE SCOPE OF THE TOPIC
As an introduction to M1 and M2 macrophages, a chronology of
results (and publications) that led to their discovery is summarized
below:
• Macrophages have the unique ability to produce a growthinhibiting molecule (NO) or a growth-promoting molecule
(ornithine), through the enzymatic conversion of l-arginine in
different ways (6–8).
• Macrophages in normal tissues, healing wounds, or in growing
tumors metabolize arginine primarily to ornithine via arginase
(later to be named M2-type). Macrophages can switch to producing NO via iNOS (to be named M1-type) that is necessary
to kill cancer or many pathogens. Arginine is the source of both
ornithine and NO (9–12).
• Macrophages were specifically renamed M1 and M2 to highlight
that innate immunity controls adaptive immunity. M1 (NO)
or M2 (ornithine)-type macrophage responses are T cell independent and they stimulate Th1-type and CTL responses, or
Th2-type responses, respectively (1, 3–5). Thus, M1/M2 represents a sea change in our understanding of how immune
responses occur.
These studies have stimulated thousands of publications that
have enhanced our knowledge of the importance of M1/inhibit
and M2/heal functions, and other cytokines and factors that
accompany these responses (1). Here, we have assembled papers
by contributors that focus on basic aspects of macrophage biology, their roles in various diseases, and how they are regulated.
Macrophages evolved long before other immune cell types and
are the foundation of all animal immunity (13). Therefore, we
begin with a series of “introductory” articles where readers can
find basic information about macrophage biology and functions.
These articles also trace the evolutionary origins of macrophages
to aid in understanding their central role in immune systems. Next,
articles detail the roles of macrophages in protection against (or
causation of) various diseases including wounds, cancer, infections, atherosclerosis, obesity, hypertension, and other conditions.
Finally, we look to the future with several articles detailing how
macrophage M1/inhibit and M2/heal functions might be modulated for therapeutic benefits. We hope that the articles enhance
your knowledge of this singularly multitalented and remarkable
leukocyte.

February 2015 | Volume 6 | Article 59 | 7

Mills et al.

INTRODUCTION TO MACROPHAGE BIOLOGY AND
FUNCTIONS
To better appreciate macrophages, it is useful to known about their
unique biochemistry, functions, and central role in all immune
systems. Drs. Rath and Munder provide a comprehensive biochemical introduction to macrophage arginine metabolism, and
how these cells can take “the fork in the road” to make either
an arginine-based M1/inhibit or M2/heal response (14). Because
“Nothing makes sense except in the light of evolution” (Theodosius Dobzhansky), Dr. Buchmann traces the evolutionary origins
of both innate and adaptive immunity, and shows when new
macrophage (and other) immune functions evolved, culminating
in humans (15). Dr. Dzik importantly reveals that the macrophage
M2/heal function (arginases) preceded the M1/inhibit function
in animal evolution (16). Drs. Mills, Thomas, Lenz, and Munder
describe the basic “SHIP” functions of macrophages [sample, heal,
inhibit, and present (antigen)], and why it is important to study
these functions to understand macrophage biology in vivo (17). In
a similar vein, Drs. Italiani and Boraschi elucidate why examining
macrophages by functions versus “phenotypes” can be critical for
understanding how they affect health (18). Dr. Harris’ piece colorfully describes the two-edged sword nature of macrophages as
“The Good the Bad and the Ugly” phases of inflammation. He also
illuminates that M1/inhibit or M2/heal-type macrophage activities vary enormously in different microenvironments of lungs or
other sites of inflammation (19). In turn, important local functions can be lost if one grinds up whole organs as is common. Drs.
Thomas and Mattila provide an in-depth look at macrophage arginine metabolism in different vertebrate species (20). Importantly,
they show that, contrary to some reports (21, 22), macrophages
in mice and humans are quite similar, as one might expect from
analyzing evolution (e.g., Drs. Buchmann and Dzik, mentioned
earlier).

MACROPHAGE INFLUENCES IN DIFFERENT DISEASES
WOUNDS AND CANCER: STARK EXAMPLES OF THE TWO-EDGED
SWORD NATURE OF MACROPHAGE RESPONSES

Wound healing requires M2/heal-type responses (9). If M1/inhibit
responses occur (e.g., infected wounds) wound healing is delayed
until the infection is cleared (5). M2/heal-type macrophages also
dominate inside tumors in experimental animals and humans (5,
11, 23). In marked contrast to their beneficial effect in wounds,
M2-type macrophages actively promote tumor growth [reviewed
in Ref. (3)], in part, by secreting growth factors (11, 24, 25).
Regarding the relative roles of M1 or M2-type macrophages
in wounds or implanted biomaterials, Drs. Brown, Sicari, and
Badylak demonstrate that there is 2–3 day dominance of M1-type
macrophages (26). These data support that the first biologic priority of hosts following injuries is to prevent infections (5). However,
if an injury is sterile, the priority switches to M2-type dominant
macrophages that are necessary for proper healing (17). Interestingly, they note that biomaterials with larger pore sizes have
less scarring/fibrosis. Thus, the physical properties of implanted
materials seem important in allowing functional regeneration
over typical imperfect wound healing found in adult humans
(27). Drs. Beljaars, Schippers, Smit, Martinez, Helming, Poelstra,
and Melgert compared M1- and M2-types of macrophages inside

Frontiers in Immunology | Molecular Innate Immunity

Summary of M1/M2 macrophages topic

chemically damaged livers in mice and cirrhotic livers in humans
(28). The liver is one of the few organs able to regenerate (though
not perfectly) (29). So, it was interesting that they found a predominance of M1 macrophages during resolution of damage, which
contrasts with M2 macrophages that dominate in wounds elsewhere, and which results in scarring/healing, as described. Also,
interesting was that the authors observed distinct M1 and M2type macrophages in close proximity to each other with little
overlap in markers. These findings do not support the hypothesis that macrophages only resemble a “color wheel” with multiple
overlapping characteristics (30).
In contrast to the beneficial effects of M2-type macrophages in
wounds, these same types of macrophages promote cancer growth
and metastases as mentioned [reviewed in Ref. (3)]. Why is there
cancer, and why does the immune system help it grow?
Species successfully evolve by acquiring traits that provide survival advantages, and losing undesirable traits. Environmental and
societal influences aside, the way animals (any species) change
heritable traits is through producing progeny: breeding. Cancer in humans occurs mainly after breeding age. So, there has
been little evolutionary pressure (or advantage) for humans to
acquire traits that prevent cancer, or that could stop it if it appears.
The same lack of evolutionary pressure applies to atherosclerosis,
or many other “adult” diseases that mainly occur during postbreeding (to be discussed later). Too, mate selection (important in
breeding success) is mostly unaffected by knowledge of whether
parents or grandparents died of cancer or other late-appearing
diseases (3).
Once it appears, cancer can be eliminated. How? Modulating macrophage functions. It is now known that the majority leukocytes in tumors are macrophages: sometimes >50% of
a tumor mass. However, as mentioned, these tumor-associated
macrophages (TAM) are primarily M2-type and actively promote tumor growth: Cancer is often referred to as “a wound
that doesn’t heal” (31). But, a wealth of recent evidence indicates that decreasing M2/heal-type and increasing M1/inhibittype macrophages can slow or reverse tumor growth (11, 32).
This is an exciting development because conventional immunologic thinking purports that tumors need to be recognized as
“foreign,” like a pathogen. But, most tumors are not “foreign.”
So, it falls on the shoulders of innate immunity, not adaptive
immunity (T and B cells), to stop cancer. Importantly, recent
antitumor effects being observed seem primarily (or solely) mediated by macrophages/innate immunity, not T cells (32). Even if
a human cancer is recognized as “foreign,” it is still critical to
switch M2- to M1-like macrophages. This is so because of the
new knowledge, discussed earlier, that M1-type macrophages are
necessary to stimulate T cells to make tumoricidal Th1-type cellular killer responses such as CTL and further amplify M1/inhibit
macrophages (1, 4). In a related connection, significant prolongation of survival in human cancer has recently been observed by
inhibiting immunoregulatory molecules, such as PD-1 and CTLA4
(33). The effects observed have been postulated to involve specific
anti tumor T cell activity. Such effects likely depend on modulating macrophage responses. Thus, increasing our knowledge of M1
and M2 polarization in cancer, and how to modulate it, is very
important.

February 2015 | Volume 6 | Article 59 | 8

Mills et al.

Drs. Laoui, Van Overmeire, Baetselier, Van Ginderachter, and
Raes review the evidence that M2-type macrophages predominate
in most human tumors with the notable exception of colorectal cancer (34). They also describe new evidence that colonystimulating factors are important players in determining the quantity and type of macrophages that populate tumors. MCSF is
normally present in tissues and plasma, and is associated with
M2-type macrophages. In contrast, GM-CSF is only present following injury or during infections, and is associated with M1-type
responses. Interestingly, the authors highlight findings suggesting inhibition of MCSF by various means in humans does not
only simply decrease M2-type TAM but also increases the M1/M2
ratio. Thus, altering macrophage differentiation signals can affect
macrophage polarization beneficially in clinical settings. Using a
different approach, Drs. Fritz, Tennis, Orlicky, Lin, Ju, Redente,
Choo, Staab, Bouchard, Merrick, Malkinson, and Dwyer-Nield
show that treatment of lung cancer in mice with a macrophagedepleting agent (clodronate-encapsulated liposomes) significantly
decreases tumor burden (35). They also show that this treatment stimulates lung TAM that have a mixed M1- and M2-type
phenotype suggesting that this depletion modality (like MCSF
inhibition) may also increase the M1/M2 TAM balance. Together,
these studies, like many others, are indicating that there are real and
important clinical benefits from immunologically manipulating
macrophage functions in cancer.
MACROPHAGES IN INFECTIONS

In the context of animal models of bacterial infection, M1-type
macrophages and NO production are often, but not exclusively
(36), associated with host protection (17). Conversely, M2-type
macrophages are typically associated with bacterial persistence.
An article by Drs. Ka, Daumas, Textoris, and Mege reviews
macrophage polarization in infectious diseases (37). They discuss some difficulties encountered when trying to extend these
concepts to bacterial infections in humans. In humans suffering from leprosy or Whipple’s, macrophage, M2-type polarization
can be readily observed. However, as mentioned earlier, analyzing whole organs can overlook microenvironmental differences
in inflammation (19). Also, studies with human patients often
utilize peripheral blood monocytes that lack the polarized M1or M2-type functions associated with tissue macrophages. They
also review that many pathogens, such as Leishmania, can survive
or spread by blocking or subverting the process of macrophage
development toward an M1/inhibit phenotype (27). An article
from Drs. Burrack and Morrison discusses how macrophage activation and arginine metabolism by M1- or M2-type macrophages
can have diverse effects on health and disease during viral infections (38). Macrophage NO production during viral infections,
as in other settings, can be induced independent of lymphocytes
(4). The production of NO can have direct anti-microbial effects
on certain bacteria, fungi, and viruses. Hence, the M1/inhibit
phenotype in these settings plays an immune protective role.
However, because NO kills non-specifically, it can also have
immunopathologic or immune suppressive effects during infections by influenza, herpes simplex virus-1, and cytomegalovirus.
Similarly, the two-edged sword nature of M2/heal macrophage
responses can cut both ways. For example, M2/heal responses

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Summary of M1/M2 macrophages topic

(via arginase and growth-promoting ornithine) usefully promote
tissue repair in some viral infection models, and via stimulating protective antibody responses. However, M2/heal responses
are also associated with viral persistence or immune pathology during many infections, such as coronavirus-induced sudden
acute respiratory syndrome (SARS), hepatitis B or C viruses,
Ross river virus, HIV, and influenza. These articles indicate
that it is important to understand the infectious disease type,
stage, and severity in order to properly modulate M1- or M2type responses to optimally eliminate pathogens and decrease
untoward pathology.
MACROPHAGE RESPONSES IN ATHEROSCLEROSIS AND OTHER
NON-PATHOGEN-INDUCED INFLAMMATORY CONDITIONS

In contrast to the primarily protective role of M1/inhibittype responses against infection agents, described above, these
killer/damaging activities are often associated with bad outcomes
in chronic inflammatory conditions.
Regarding atherosclerosis, Drs. Thomas and Mattila show that
both M1- and M2-types of macrophages are found during foam
cell formation, a hallmark of atherosclerosis (20). Drs. Hayes,
Tsaousi, Gregoli, Jenkinson, Bond, Johnson, Bevan, Thomas, and
Newby show that there is an altered expression of certain matrix
metalloproteinases in atherosclerosis (39). Interestingly, they also
show that M1 and M2 macrophage polarization in atherosclerotic
ApoE null mice occurs in the absence of T- and B-lymphocytes,
again highlighting the independence of innate immunity from
adaptive immunity discussed earlier. Drs. Murphy, Dragoljevic,
and Tall review the recent evidence that cholesterol efflux pathways
regulate myelopoiesis (40). Traditionally, cholesterol efflux was
considered as a safeguard against foam cell formation, but Tall’s
group has shown that knocking out cholesterol efflux molecules
like ABCA1 and ABCG1 cause profound changes in hematopoiesis
associated with more Ly6C+ inflammatory monocytes and more
neutrophils. This shift could lead to altered macrophage function. Drs. Peled and Fisher review dynamic aspects of macrophage
polarization during atherosclerosis progression and regression:
progression is associated with macrophage M1 polarization and
regression with M2 polarization (41). The article by Drs. Yang
and Ming looks at the less commonly studied arginase II enzyme.
They show that, unlike arginase I, that is typically inversely
related to macrophage NO production, arginase II seems under
different regulation. Also, Arg II expression in endothelial cells
tends to uncouple eNOS (or NOS1) causing loss of vascular
tone (42).
Regarding other non-pathogen associated inflammatory conditions, Drs. Vlahos and Bozinovski review the role of alveolar
macrophages in chronic obstructive pulmonary disease (COPD)
(43). COPD is a widespread chronic inflammatory condition
with immense medical and societal impact. Interestingly, in
COPD, there is an accumulation of airway macrophages that
show a transcriptome skewed toward wound healing M2 markers suggesting defective resolution of inflammation (as occurs
in wound healing). Drs. Kraakman, Murphy, Jandeleit-Dahm,
and Kammoun review macrophage polarization in obesity and
type 2 diabetes (44). M1 “pro-inflammatory” macrophages are
enhanced compared with M2 “anti-inflammatory” macrophages,

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Mills et al.

leading to chronic inflammation and the propagation of metabolic
dysfunctions. The brain and spinal cord are primarily populated by
macrophage-like microglial cells, which are derived from yolk sac
precursors under resting conditions. Drs. Cherry, Olschowka, and
O’Banion show that these microglia are normally M2-polarized,
although the microglia transcriptome is different from that of
M2 macrophages in other organs (45). This observation fits
well with an emerging concept that M1 and M2 polarization
varies between organs (46). In this connection, Drs. Brown,
von Chamier, Allam, and Reyes report on M1/M2 macrophage
polarity in normal and complicated pregnancies and find that
the balance and location of M1- and M2-type responses show
significant variation (47). In general, over expression of M1type macrophages is associated with untoward outcomes during
pregnancy.
Finally, this Topic focuses on macrophage polarization. Little
is currently known about how the origin of macrophages in tissues (e.g., yolk sac, Ly6C-high, Ly-6C-low monocytes) influence
M1/M2 polarization. Drs. Dey, Allen, and Hankey-Giblin begin
to explore how the ontogeny of monocytes and macrophage can
influence M1- and M2-type responses in different tissues (48).

REGULATION OF MACROPHAGE DIFFERENTIATION AND
FUNCTIONS
The importance of macrophage M1/inhibit and M2/heal imbalances in various disease or protective processes being clear,
immunologists and clinicians are interested in how they might
therapeutically intervene to shape macrophage differentiation or
modulate various macrophage functions to restore health. Drs.
Wang, Liang, and Zen provide an overview of the various molecular mechanisms known to impact macrophage M1 and M2 polarization (49). Several of these mechanisms are expanded upon in
subsequent articles. Cytokines such as interferon gamma (IFN-γ)
have profound impacts on development of M1 and M2 functionality in macrophages. Regulators of cytokine receptor signaling can
importantly impact macrophage functions. Dr. Wilson details in
her article evidence that various suppressor of cytokine signaling
(SOCS) protein family members shape M1/M2 macrophage functions in several disease settings (50). Colony-stimulating factors
(CSFs) and macrophage stimulating protein (MSP) also impact
macrophage responsiveness to polarization. Drs. Hamilton, Zhao,
Pavicic, and Datta introduce the concept that the myeloid colonystimulating factors MCSF and GM-CSF act not only to promote
the development and maintenance of various myeloid populations
but can also shape the responsiveness of macrophages to stimuli
that direct the M1 or M2 phenotype (51). The effects of MSP
and its receptor, a tyrosine kinase known as RON, on macrophage
polarization are clearly presented in an article by Dr. Chaudhuri
(52). Other extrinsic factors that regulate macrophage polarization include components of the complement cascade as well as
extracellular nucleotides. The contrasting effects of various complement components on M1 and M2 functions are described in
an article by Drs. Bohlson O’Conner, Hulsebus, Ho, and Fraser
(53). Drs. Desai and Leitinger go on to detail how purinergic
receptors for extracellular ATP and other nucleotides couple with
calcium signaling to modulate macrophage activities and resolution of inflammation (54). An improved understanding of how

Frontiers in Immunology | Molecular Innate Immunity

Summary of M1/M2 macrophages topic

these extrinsic factors and intrinsic signaling pathways regulate the
acquisition of M1 and M2-type functions will lead to improved
methods for fine-tuning of macrophage polarization to promote
health.
As mentioned above, a key difference between M1 and M2
macrophages is in their processing of l-arginine. NO can not only
kill susceptible microbes but also has a variety of signaling and
regulatory effects on macrophages and other cell types. NO is
generated from l-arginine through the activities of three NO synthase (NOS) enzymes. In contrast to canonical views, Drs. Mattila
and Thomas present the perspective that “constitutive” enzyme
activities (NOS1 and 3) can be induced, and “inducible” NOS2 is
constitutively expressed in several tissues (55). The different functions of M1 and M2 macrophages are also associated with changes
in the metabolic pathways they use to produce ATP. Drs. GalvánPeña and O’Neill discuss the differences in metabolism between
M1 and M2 macrophages and how these differences impact other
aspects of macrophage function (56). M1 macrophages mainly
rely on glycolysis for energy, while M2 macrophages primarily
use oxidative phosphorylation. Accumulation of succinate in M1
macrophages can stimulate HIF1α to sustain production of factors
such as IL-1β and thus can impact the ability of M1 macrophages
to prolong inflammation. Manipulation of NOS enzyme expression and activities as well as the products and consequences of the
different metabolic processes in M1 and M2 macrophages should
further our ability to shape the outcome of infections and other
diseases.

SUMMARY
The “Fork in the Road” that macrophages take in making
either M1/inhibit or M2/heal-type responses define “immunity”
throughout the animal kingdom. In all animals, M1/inhibittype responses are the primary host defense, and M2/heal-type
responses help repair and replace lost or effete tissue to maintain host homeostasis. In humans (and other higher animals),
macrophage M1/inhibit or M2/heal-type responses necessarily
direct T (and B) cells/adaptive immunity to make Th1 or Th2like responses. Thus, whether acting directly or indirectly, which
“fork” macrophages take is the central controlling element that
promotes health (as in pathogen control or wound repair) or
impedes health (as in atherosclerosis, autoimmunity, or cancer).
By illuminating the biochemical underpinnings, evolution, diseases, and regulation of macrophage functions, the papers in
this Topic advance our understanding of how to modulate this
most important of all leukocytes: the chicken and the egg of
immunity (1).

ACKNOWLEDGMENTS
We are grateful for the many good papers and discussions that
have occurred in creating this Topic and EBook.

REFERENCES
1. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity.
J Innate Immun (2014) 6:716–26. doi:10.1159/000364945
2. Metchnikoff E. Untersuchungen uber die intracellulare Verdauung bei Wirbellosen
Thieren. Arbeit Zoologischen Instituten, Universitat Wien (1883) 5:141–68.
3. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32:463–88. doi:10.1615/CritRevImmunol.v32.i6.10

February 2015 | Volume 6 | Article 59 | 10

Mills et al.

4. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages and
the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.4049/jimmunol.
164.12.6166
5. Mills CD. Macrophage arginine metabolism to ornithine/urea or nitric
oxide/citrulline: a life or death issue. Crit Rev Immunol (2001) 21:399–425.
doi:10.1615/CritRevImmunol.v21.i5.10
6. Currie GA, Gyure L, Cifuentes L. Microenvironmental arginine depletion by
macrophages in vivo. Br J Cancer (1979) 39:613–20. doi:10.1038/bjc.1979.112
7. Hibbs JB, Vavrin Z, Taintor RR. L-arginine is required for expression of the activated macrophage effector mechanism causing selective metabolic inhibition in
target cells. J Immunol (1987) 138:550–65.
8. Albina JE, Caldwell MD, Henry WL, Mills CD. Regulation of macrophage
functions by L-arginine. J Exp Med (1989) 169:1021–9. doi:10.1084/jem.169.
3.1021
9. Albina JE, Mills CD, Henry WL, Caldwell MD. Temporal expression of different pathways of 1-arginine metabolism in healing wounds. J Immunol (1990)
144:3877–80.
10. Nathan CF, Hibbs JB. Role of nitric oxide synthesis in macrophage antimicrobial activity. Curr Opin Immunol (1991) 3:65–70. doi:10.1016/0952-7915(91)
90079-G
11. Mills CD, Shearer J, Evans R, Caldwell MD. Macrophage arginine metabolism
and the inhibition or stimulation of cancer. J Immunol (1992) 149:2709–14.
12. Wu G, Morris SM. Arginine metabolism: nitric oxide and beyond. Biochem J
(1998) 336(Pt 1):1–17.
13. Dzik JM. The ancestry and cumulative evolution of immune reactions. Acta
Biochim Pol (2010) 57:443–66.
14. Rath M, Müller I, Kropf P, Closs EI, Munder M. Metabolism via arginase or
nitric oxide synthase: two competing arginine pathways in macrophages. Front
Immunol (2014) 5:532. doi:10.3389/fimmu.2014.00532
15. Buchmann K. Evolution of innate immunity: clues from invertebrates via fish
to mammals. Front Immunol (2014) 5:459. doi:10.3389/fimmu.2014.00459
16. Dzik JM. Evolutionary roots of arginase expression and regulation. Front
Immunol (2014) 5:544. doi:10.3389/fimmu.2014.00544
17. Mills CD, Thomas AC, Lenz LL, Munder M. Macrophage: SHIP of Immunity.
Front Immunol (2014) 5:620. doi:10.3389/fimmu.2014.00620
18. Italiani P, Boraschi D. From monocytes to M1/M2 macrophages: phenotypical
vs. functional differentiation. Front Immunol (2014) 5:514. doi:10.3389/fimmu.
2014.00514
19. Harris RA. Spatial, temporal, and functional aspects of macrophages during “the
good, the bad, and the ugly” phases of inflammation. Front Immunol (2014)
5:612. doi:10.3389/fimmu.2014.00612
20. Thomas AC, Mattila JT. “Of mice and men”: arginine metabolism in
macrophages. Front Immunol (2014) 5:479. doi:10.3389/fimmu.2014.00479
21. Schneemann M, Schoedon G. Species differences in macrophage NO production
are important. Nat Immunol (2002) 3:102–102. doi:10.1038/ni0202-102a
22. Martinez FO, Gordon S, Locati M, Mantovani A. Transcriptional profiling
of the human monocyte-to-macrophage differentiation and polarization: new
molecules and patterns of gene expression. J Immunol (2006) 177:7303–11.
doi:10.4049/jimmunol.177.10.7303
23. Sica A, Schioppa T, Mantovani A, Allavena P. Tumour-associated macrophages
are a distinct M2 polarised population promoting tumour progression: potential targets of anti-cancer therapy. Eur J Cancer (2006) 42:717–27. doi:10.1016/
j.ejca.2006.01.003
24. Baay M, Brouwer A, Pauwels P, Peeters M, Lardon F. Tumor cells and tumorassociated macrophages: secreted proteins as potential targets for therapy. Clin
Dev Immunol (2011) 2011:565187. doi:10.1155/2011/565187
25. Noy R, Pollard JW. Tumor-associated macrophages: from mechanisms to therapy. Immunity (2014) 41:49–61. doi:10.1016/j.immuni.2014.06.010
26. Brown BN, Sicari BM, Badylak SF. Rethinking regenerative medicine: a
macrophage-centered approach. Front Immunol (2014) 5:510. doi:10.3389/
fimmu.2014.00510
27. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol (2011) 11:723–37. doi:10.1038/nri3073
28. Beljaars L, Schippers M, Reker-Smit C, Martinez FO, Helming L, Poelstra
K, et al. Hepatic localization of macrophage phenotypes during fibrogenesis
and resolution of fibrosis in mice and humans. Front Immunol (2014) 5:430.
doi:10.3389/fimmu.2014.00430

www.frontiersin.org

Summary of M1/M2 macrophages topic

29. Chazaud B. Macrophages: supportive cells for tissue repair and regeneration.
Immunobiology (2014) 219:172–8. doi:10.1016/j.imbio.2013.09.001
30. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8:958–69. doi:10.1038/nri2448
31. Dvorak HF. Tumors: wounds that do not heal. Similarities between tumor
stroma generation and wound healing. N Engl J Med (1986) 315:1650–9.
doi:10.1056/NEJM198612253152606
32. Beatty GL, Chiorean EG, Fishman MP, Saboury B, Teitelbaum UR, Sun W,
et al. CD40 agonists alter tumor stroma and show efficacy against pancreatic
carcinoma in mice and humans. Science (2011) 331:1612–6. doi:10.1126/science.
1198443
33. Swanson MS, Sinha UK. Rationale for combined blockade of PD-1 and CTLA-4
in advanced head and neck squamous cell cancer – review of current data. Oral
Oncol (2015) 51:12–5. doi:10.1016/j.oraloncology.2014.10.010
34. Laoui D, Van Overmeire E, De Baetselier P, Van Ginderachter JA, Raes G. Functional relationship between tumor-associated macrophages and macrophage
colony-stimulating factor as contributors to cancer progression. Front Immunol
(2014) 5:489. doi:10.3389/fimmu.2014.00489
35. Fritz JM, Tennis MA, Orlicky DJ, Lin H, Ju C, Redente EF, et al. Depletion of
tumor-associated macrophages slows the growth of chemically induced mouse
lung adenocarcinomas. Front Immunol (2014) 5:587. doi:10.3389/fimmu.2014.
00587
36. Cole C, Thomas S, Filak H, Henson PM, Lenz LL. Nitric oxide increases susceptibility of toll-like receptor-activated macrophages to spreading Listeria monocytogenes. Immunity (2012) 36:807–20. doi:10.1016/j.immuni.2012.03.011
37. Ka MB, Daumas A, Textoris J, Mege J-L. Phenotypic diversity and emerging
new tools to study macrophage activation in bacterial infectious diseases. Front
Immunol (2014) 5:500. doi:10.3389/fimmu.2014.00500
38. Burrack KS, Morrison TE. The role of myeloid cell activation and arginine
metabolism in the pathogenesis of virus-induced diseases. Front Immunol
(2014) 5:428. doi:10.3389/fimmu.2014.00428
39. Hayes EM, Tsaousi A, Gregoli KD, Jenkinson SR, Bond AR, Johnson JL, et al.
Classical and alternative activation and metalloproteinase expression occurs in
foam cell macrophages in male and female ApoE null mice in the absence of
T and B lymphocytes. Front Immunol (2014) 5:537. doi:10.3389/fimmu.2014.
00537
40. Murphy AJ, Dragoljevic D, Tall AR. Cholesterol efflux pathways regulate
myelopoiesis: a potential link to altered macrophage function in atherosclerosis.
Front Immunol (2014) 5:490. doi:10.3389/fimmu.2014.00490
41. Peled M, Fisher EA. Dynamic aspects of macrophage polarization during atherosclerosis progression and regression. Front Immunol (2014) 5:579. doi:10.
3389/fimmu.2014.00579
42. Yang Z, Ming XF. Functions of arginase isoforms in macrophage inflammatory
responses: impact on cardiovascular diseases and metabolic disorders. Front
Immunol (2014) 5:533. doi:10.3389/fimmu.2014.00533
43. Vlahos R, Bozinovski S. Role of alveolar macrophages in chronic obstructive pulmonary disease. Front Immunol (2014) 5:435. doi:10.3389/fimmu.2014.00435
44. Kraakman MJ, Murphy AJ, Jandeleit-Dahm K, Kammoun HL. Macrophage
polarization in obesity and type 2 diabetes: weighing down our understanding of macrophage function? Front Immunol (2014) 5:470. doi:10.3389/fimmu.
2014.00470
45. Cherry JD, Olschowka JA, O’Banion MK. Are “resting” microglia more “M2”?
Front Immunol (2014) 5:594. doi:10.3389/fimmu.2014.00594
46. Okabe Y, Medzhitov R. Tissue-specific signals control reversible program
of localization and functional polarization of macrophages. Cell (2014)
157:832–44. doi:10.1016/j.cell.2014.04.016
47. Brown MB, von Chamier M, Allam AB, Reyes L. M1/M2 macrophage polarity in normal and complicated pregnancy. Front Immunol (2014) 5:606.
doi:10.3389/fimmu.2014.00606
48. Dey A, Allen J, Hankey-Giblin PA. Ontogeny and polarization of macrophages
in inflammation: blood monocytes versus tissue macrophages. Front Immunol
(2014) 5:683. doi:10.3389/fimmu.2014.00683
49. Wang N, Liang H, Zen K. Molecular mechanisms that influence the macrophage
M1-M2 polarization balance. Front Immunol (2014) 5:614. doi:10.3389/fimmu.
2014.00614
50. Wilson HM. SOCS proteins in macrophage polarization and function. Front
Immunol (2014) 5:357. doi:10.3389/fimmu.2014.00357

February 2015 | Volume 6 | Article 59 | 11

Mills et al.

51. Hamilton TA, Zhao C, Pavicic PG, Datta S. Myeloid colony-stimulating factors as regulators of macrophage polarization. Front Immunol (2014) 5:554.
doi:10.3389/fimmu.2014.00554
52. Chaudhuri A. Regulation of macrophage polarization by RON receptor
tyrosine kinase signaling. Front Immunol (2014) 5:546. doi:10.3389/fimmu.
2014.00546
53. Bohlson SS, O’Conner SD, Hulsebus HJ, Ho M-M, Fraser DA. Complement, C1q,
and C1q-related molecules regulate macrophage polarization. Front Immunol
(2014) 5:402. doi:10.3389/fimmu.2014.00402
54. Desai BN, Leitinger N. Purinergic and calcium signaling in macrophage
function and plasticity. Front Immunol (2014) 5:580. doi:10.3389/fimmu.2014.
00580
55. Mattila JT, Thomas AC. Nitric oxide synthase: non-canonical expression patterns. Front Immunol (2014) 5:478. doi:10.3389/fimmu.2014.00478
56. Galván-Peña S, O’Neill LAJ. Metabolic reprograming in macrophage polarization. Front Immunol (2014) 5:420. doi:10.3389/fimmu.2014.00420

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Summary of M1/M2 macrophages topic

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 20 January 2015; accepted: 30 January 2015; published online: 16 February
2015.
Citation: Mills CD, Lenz LL and Ley K (2015) Macrophages at the fork in the road to
health or disease. Front. Immunol. 6:59. doi: 10.3389/fimmu.2015.00059
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2015 Mills, Lenz and Ley. This is an open-access article distributed under
the terms of the Creative Commons Attribution License (CC BY). The use, distribution
or reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
accepted academic practice. No use, distribution or reproduction is permitted which
does not comply with these terms.

February 2015 | Volume 6 | Article 59 | 12

REVIEW ARTICLE
published: 27 October 2014
doi: 10.3389/fimmu.2014.00532

Metabolism via arginase or nitric oxide synthase: two
competing arginine pathways in macrophages
Meera Rath 1 , Ingrid Müller 2 , Pascale Kropf 2 , Ellen I. Closs 3 and Markus Munder 4,5 *
1

Department of Pharmacology, Institute of Medical Sciences, Faculty of Medical Sciences, Siksha ’O’ Anusandhan University, Bhubaneshwar, India
Section of Immunology, Department of Medicine, Imperial College London, London, UK
3
Department of Pharmacology, University Medical Center, Johannes Gutenberg University, Mainz, Germany
4
Third Department of Medicine (Hematology, Oncology, and Pneumology), University Medical Center, Johannes Gutenberg University, Mainz, Germany
5
Research Center for Immunotherapy, University Medical Center, Johannes Gutenberg University, Mainz, Germany
2

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Takayuki Yoshimoto, Tokyo Medical
University, Japan
Jose M. Fuentes, Universidad de
Extremadura, Spain
*Correspondence:
Markus Munder , Third Department of
Medicine (Hematology, Oncology, and
Pneumology), University Medical
Center, Johannes Gutenberg
University, Langenbeckstrasse 1,
Mainz 55131, Germany
e-mail: munder@uni-mainz.de

Macrophages play a major role in the immune system, both as antimicrobial effector cells
and as immunoregulatory cells, which induce, suppress or modulate adaptive immune
responses. These key aspects of macrophage biology are fundamentally driven by the
phenotype of macrophage arginine metabolism that is prevalent in an evolving or ongoing immune response. M1 macrophages express the enzyme nitric oxide synthase, which
metabolizes arginine to nitric oxide (NO) and citrulline. NO can be metabolized to further
downstream reactive nitrogen species, while citrulline might be reused for efficient NO
synthesis via the citrulline–NO cycle. M2 macrophages are characterized by expression of
the enzyme arginase, which hydrolyzes arginine to ornithine and urea. The arginase pathway limits arginine availability for NO synthesis and ornithine itself can further feed into the
important downstream pathways of polyamine and proline syntheses, which are important
for cellular proliferation and tissue repair. M1 versus M2 polarization leads to opposing outcomes of inflammatory reactions, but depending on the context, M1 and M2 macrophages
can be both pro- and anti-inflammatory. Notably, M1/M2 macrophage polarization can be
driven by microbial infection or innate danger signals without any influence of adaptive
immune cells, secondarily driving the T helper (Th)1/Th2 polarization of the evolving adaptive immune response. Since both arginine metabolic pathways cross-inhibit each other on
the level of the respective arginine break-down products and Th1 and Th2 lymphocytes can
drive or amplify macrophage M1/M2 dichotomy via cytokine activation, this forms the basis
of a self-sustaining M1/M2 polarization of the whole immune response. Understanding the
arginine metabolism of M1/M2 macrophage phenotypes is therefore central to find new
possibilities to manipulate immune responses in infection, autoimmune diseases, chronic
inflammatory conditions, and cancer.
Keywords: macrophage, M1 and M2, arginine, arginase, nitric oxide synthase, immunoregulation, amino acid
transporter

INTRODUCTION: ARGININE IN THE CENTER OF M1/M2
MACROPHAGE DICHOTOMY
Macrophages are highly versatile cells, which are (i) crucial
for infection control (“kill/fight mode”) and tissue homeostasis
(“default mode”, phagocytosing cellular debris) and (ii) involved
in disease pathophysiology in cancer, autoimmunity, metabolic,
and fibrotic disorders (1). Macrophages react to a wide variety
of external stimuli and are able to produce a multitude of effector molecules for intercellular communication, microbial defense,
and modulation of inflammatory reactions (1). They induce, suppress, or modulate both innate and adaptive immune responses.
Considering this enormous complexity it is reasonable to deduce
classification schemes to create order and sense in the experimental
results of macrophage research (2, 3). Potential macrophage diversity, both in terms of activation states, surface marker expression,
metabolic phenotype, and interspecies differences clearly requires
rigid standards for experimental set-up and reporting (4). While
undue simplification hampers the comparability between studies

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(4), a reductionist approach tries to avoid getting lost in the complexities of macrophage biology and has both enormous power
for the explanation of reality and can be the basis for experimentally testing of hypotheses. One should never forget that even
the most sophisticated modern computers are based on the “0–1”
dichotomy!
One of the most fruitful and reasonable classification of
macrophages relates to their two main functions, namely, to
kill/fight or to heal/fix. Within this classification view, macrophage
biology is driven by two phenotypes (M1 for killing/fighting
versus M2 for healing/fixing), which are also relevant in an
evolving or ongoing immune response (5). M1 or M2 dominant macrophages then direct T lymphocytes to produce Th1 or
Th2 responses, respectively, to further amplify M1 or M2 type
responses in positive feed-back loops stabilizing the predominant
immune phenotype in the respective setting of infection, tumor,
or inflammation. The M1/M2 macrophage classification can be
condensed into two opposing pathways for the metabolism of one

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Rath et al.

amino acid: the preference of macrophages to metabolize arginine via nitric oxide synthase (NOS) to NO and citrulline or via
arginase to ornithine and urea defines them as M1 (NOS) or M2
(arginase) macrophages (5). NOS or arginase are enzymes that
catalyze a “reaction,” but we will use “pathway” here to illuminate that they are part of multi-enzyme pathways producing other
physiologically important products.
In this introductory review, we will describe macrophage
arginine metabolism, its functional consequences and how the
macrophage arginine metabolic phenotype defines the two opposing M1 and M2 types of macrophages. While various molecules and features of macrophages are reciprocally or mutually
exclusively associated with the M1 versus M2 phenotype, the
dichotomous regulation of arginine metabolism is at the center
of the different functions that are associated with M1 and M2
macrophages.

ARGININE: ONE SMALL AMINO ACID FOR MACROPHAGE
METABOLISM, A GIANT CONTROLLER FOR MAMMALIAN
(PATHO-)PHYSIOLOGY
Mammalian arginine metabolism is complex both at the level
of the whole organism (6, 7) and at the level of the individual
cell types (8) and we would like to adopt the term “argenomics”
that was suggested by Sidney M. Morris Jr (7) for the regulation
of gene expression via arginine availability in an even broader
sense for the whole fascinating complexity of arginine-driven cellular regulation. Before focusing in on arginine metabolism of
macrophages and its determining role for the M1/M2 dichotomy,
let us first have a short overview on some historical facts relating
to arginine in mammalian physiology. The story started nearly
130 years ago, arginine was first isolated in 1886 and was identified as a component of animal proteins in 1895 (8). The role
of arginine in metabolic physiology was first demonstrated in
1932, when Krebs and Henseleit discovered the urea cycle. In 1981,
Windmueller and Spaeth reported that the small intestine is the
major source of citrulline for synthesis of arginine by the kidneys,
now called the intestinal–renal axis for arginine synthesis on an
organismal level (9). In 1987, it was shown that arginine is the
precursor for macrophage citrulline and nitrite synthesis (10) and
that arginine-derived NO is the elusive endothelium-derived relaxing factor (EDRF) (11, 12). Soon afterwards, NO was categorized
as physiologically active intermediate of the arginine to nitrite
(+nitrate) pathway in macrophages (13, 14) and endothelial cells
(15). The discovery of the fundamental role of arginine-derived
NO for human cardiovascular physiology already led to the award
of the Nobel Prize in 1998 to Robert F. Furchgott, Louis J. Ignarro,
and Ferid Murad. The importance of arginine has still risen since
then, it is now clear that immune cell arginine metabolism is fundamentally involved in cancer, inflammation, infections, fibrotic
diseases, pregnancy, and immune regulation in general (16–21). A
huge responsibility for a small molecule!

Macrophage arginine metabolism

characterized as a semi- or conditionally essential amino acid.
Average arginine ingestion with a Western style diet is around
4–5 g/day and the normal plasma arginine level is 100–200 µM.
Besides dietary intake, arginine is derived from cellular protein
break-down or endogenous de novo arginine production. Mammalian arginine biosynthesis (Figure 1) involves mainly the amino
acids glutamine, ornithine, and citrulline and the involved enzymatic steps are compartmentalized in different tissues and also on
the subcellular level so that not all reactions can take place in each
individual cell type or tissue. For a more detailed description of the
chemical pathways of arginine metabolism, the reader is referred
to the excellent review by Wu and Morris (8). (i) Glutamine can
be converted to ornithine via glutaminase (yielding glutamate),
pyrroline-5-carboxylate synthetase (P5CS), which is almost exclusively expressed in the intestinal mucosa, and ornithine aminotransferase (OAT). (ii) Ornithine transcarbamylase (OTC) and
carbamoyl phosphate synthetase (CPS) are involved in the formation of citrulline from ornithine. The enzymes are restricted
to the mitochondrial matrix of hepatocytes and epithelial cells
of small and (to a minor extent) large intestine. This reaction
is therefore a part of the hepatic urea cycle and also involved in
intestinal synthesis of citrulline, which is released into the circulation. The proximal tubules of the kidneys take up most of the
circulating citrulline, which is then converted within the kidney
to arginine and again released into the circulation. (iii) Argininosuccinate synthetase (ASS) and argininosuccinate lyase (ASL)
are cytosolic enzymes responsible for the biosynthesis of arginine
from citrulline (and aspartate as a co-substrate). While ASS and
ASL are expressed constitutively or inducibly in many different
cell types, their degree of expression, and the efficiency of their
catalytic pathways vary between different tissue types. On a whole
body level, the latter enzymatic steps form the basis of the so-called
intestinal–renal axis with intestinal production of citrulline (see
above) and renal synthesis of arginine (7).
M1 macrophages can also synthesize arginine in a cyclic fashion
(Figure 2): during NO synthesis, arginine is converted to NO and
citrulline via Nω -OH-arginine (22, 23). Murine macrophages have
long been known to (i) upregulate ASS and constitutively express
ASL when stimulated with the NOS-inducing agents lipopolysaccharide (LPS) and IFN-γ (24) and (ii) to partially rescue NO
synthesis via citrulline uptake and ASS-mediated recycling to arginine (25). This set of reactions via ASS and ASL forms the so-called
citrulline–NO cycle (26). The importance of this pathway for
the resynthesis of arginine to ensure sufficient substrate supply
for prolonged NO synthesis under arginine limitation has been
recently demonstrated in vivo in murine mycobacteria infection
(27). Despite the upregulation of ASS1, availability of arginine
remains a rate-limiting step for synthesis of NO and cellular uptake
of arginine also determines the amount of NO synthesized in case
of NOS and ornithine in case of arginase (28, 29).

CELLULAR UPTAKE OF ARGININE
MACROPHAGE ARGININE AVAILABILITY: SEVERAL ROADS
LEAD TO ONE AMINO ACID
On the level of the whole organism, arginine is a non-essential
amino acid for healthy adult humans, but it has to be supplemented
during growth or various disease states (6, 7) and has thus been

Frontiers in Immunology | Inflammation

The transmembranous arginine transport is one essential component of cellular arginine metabolism and important for the
cells to perform their tasks. Amino acids do not pass membranous lipid bilayers freely, but are transported via specialized
proteins. These amino acid transporters show different transport

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FIGURE 1 | Important pathways of mammalian arginine metabolism.
M1 and M2 macrophages are characterized by the metabolism of arginine via
NOS or arginase with important functional consequences. This dichotomy is
put into the context of other important pathways that can lead to the
synthesis or degradation of arginine in mammalian cells. For sake of clarity,
the diagram focuses on relevant enzymes (gray-shaded boxes), metabolites
(red boxes), and the position of NOS and arginase within the network is
highlighted. Various intermediate steps, by-products, or substrates are

FIGURE 2 | Arginine metabolism via NOS or arginase is at the center of
the M1/M2 polarization of macrophages. M1 and M2 macrophages are
characterized by the metabolism of arginine via NOS or arginase with
important functional consequences. Abbreviations: see Figure 1.

characteristics and specificities for the various amino acids, based
on their physicochemical properties. Arginine, like the other
cationic amino acids ornithine and lysine is preferentially taken

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Macrophage arginine metabolism

omitted and the reader is referred to more extensive chemical reviews (see
text). ADC, arginine decarboxylase; AGAT, arginine:glycine amidinotransferase;
ASL, argininosuccinate lyase, ASS, argininosuccinate synthase; CP, carbamoyl
phosphate; CPS, CP synthase; NOS, nitric oxide synthase; OAT, ornithine
aminotransferase; ODC, ornithine decarboxylase; OTC, ornithine
transcarbamylase; P5C, pyrroline-5-carboxylate; P5CDH, P5C dehydrogenase;
P5CR, P5C reductase; P5CS, P5C synthase; PRODH, proline dehydrogenase;
SRS, spermidine synthase; SMS, spermine synthase.

up via members of the solute carrier family 7 (SLC7) (30–32).
The subfamily of cationic amino acid transporters (CAT1–3, i.e.,
SLC7A1–3) recognizes exclusively cationic amino acids, while
members of the subfamily of heteromeric amino acid transporters,
y+ LAT1 (SLC7A7) and y+ LAT2 (SLC7A6), and b0,+ AT transport
also neutral amino acids. CAT and y+ LAT proteins are widely distributed in various tissue types. CAT proteins are the main transporters for arginine uptake into cells, where the amino acid is then
used for protein synthesis and all arginine-dependent metabolic
pathways (7). CAT-1 is constitutively expressed in most tissues,
with the exception of the liver, while CAT-2B is cytokine-inducible.
CAT-2A is predominantly expressed in liver while CAT-3 is widely
expressed during embryonal development but largely restricted
to central nervous system and thymus in adults (30). In contrast
to CAT, y+ LAT proteins exchange primarily extracellular neutral
against intracellular cationic amino acids and are therefore responsible for arginine export rather then import (30, 32). b0,+ AT is
expressed in epithelial cells of small intestine and kidney where it
is responsible for the (re)absorption of cationic amino acids and
cysteine (32). ATB0,+ , a member of the SLC6 family, also transports both cationic and neutral amino acids and is expressed in
the apical membrane of epithelial cells in various tissues (30).
The complex regulation of CAT expression has been studied quite extensively (30, 32, 33), whereas comparatively little is
known about arginine transporter expression in most cells of the
hematopoetic system in general or the immune system specifically. Induction of CAT-2 has been shown in murine macrophages
upon both Th1 and Th2 cytokine stimulation (28), in murine
dendritic cells (DCs) by retinoic acid (RA) (34) and in murine

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microglia upon stimulation with IFN-γ+/− LPS (35). In contrast,
CAT-1 is either expressed constitutively (28) or even downregulated upon activation (35). Coordinated induction of CAT-2
and arginase (partially dependent on CAT-2 expression) has been
demonstrated in macrophages during murine allergic lung inflammation, forming the basis of bleomycin-induced fibrosis (36) and
also in RA-activated DCs (34). On the other hand, sustained production of NO in murine macrophages is also based on CAT-2
expression and CAT-2-mediated arginine uptake (37). The induction of CAT-2 in both M1 and M2 type murine macrophages
has been shown to differ quantitatively between different mouse
strains with important pathophysiological consequences: a deletion in the SLC7A2 promoter of C57BL/6 mice leads to impaired
CAT-2 expression, reduced arginine uptake, and decreased susceptibility to Leishmania infection as compared to BALB/c mice (38).
In contrast to murine macrophages, arginine transport is based on
system y+ L in IFN-γ-activated human primary monocytes (39)
or LPS-stimulated alveolar macrophages (40), another example of
interspecies differences, which are so prominent in various aspects
of arginine metabolism in the immune system (17, 41, 42).
A crucial principle of immune cell signaling is the constitutive preexistence or the activation-induced assembly of multiprotein complexes that facilitate efficient transduction of stimuli.
In murine myeloid cells, enhanced arginine import via CAT-2 is
coupled to the induction of the arginine-catabolizing enzymes
arginase I (28) and NOS (28, 34, 35, 43). It remains to be analyzed if
(i) further similar higher-order structures or coordinated enzyme
induction, combining arginine transporters with specific argininemetabolizing proteins, can be found in macrophages and if (ii)
differences in M1 and M2 type cells do exist. This is a valid hypothesis since amino acid transporters are not only final elements
of distinct signal transduction pathways, which immune cells
need for nutrient supply, but are also rather intricately involved
in complex metabolic networks in which they influence further
downstream signaling nodes and metabolic pathways (44). Interestingly, the ornithine-derived polyamine spermine enhances the
expression of CAT-1 mRNA in human retinal pigment epithelial
cells (45). This observation leads to an interesting question regarding macrophages: do M2 type macrophages increase their arginine
transport capacity via endogenous arginase-mediated synthesis
of polyamines, which would then further amplify arginase-based
metabolism in a positive feed-back loop? An alternative scenario
might result from the extracellular synthesis of ornithine after
arginase has been secreted (46) or liberated unspecifically during cell death of myeloid cells (47). As ornithine is a cationic
amino acid and substrate of cationic amino acid transporters,
it (i) competitively inhibits arginine uptake and (ii) can be
exchanged for intracellular arginine via CAT transporter proteins
(31), potentially leading to a cellular depletion of arginine.

ARGININE CATABOLISM: A BIPOLAR WORLD IN
MACROPHAGES
In mammalian cells, arginine can be catabolized by four classes
of enzymes (Figure 1): NOS, arginase, arginine decarboxylase
(ADC), and arginine:glycine amidinotransferase (AGAT) (8).
Although the enzymes are of course regulated and expressed
in a cell-type-specific manner, the metabolism of arginine is

Frontiers in Immunology | Inflammation

Macrophage arginine metabolism

potentially complex since its downstream metabolites encompass
NO, urea, ornithine, citrulline, creatine, agmatine, glutamate, proline, and the family of polyamines (7, 8). In macrophages, arginine
is a precursor for mainly two important metabolic pathways: it is
metabolized either by inducible nitric oxide synthase (iNOS) to
NO and citrulline or it is hydrolyzed by arginase to ornithine
and urea (Figure 2). This fundamental dichotomy of macrophage
arginine metabolism has wide ranging implications for their function as well as for the type and outcome of immune responses in
which these innate immune cells are involved in. Before these consequences are discussed (see below), let us first look at the two
important distinct arginine enzymatic pathways in macrophages
in more detail.
NITRIC OXIDE SYNTHASE: ARGININE – NITRIC OXIDE PATHWAY

In 1980, Furchgott and co-workers discovered (i) the necessity of
an intact endothelium for relaxation of isolated blood vessels and
(ii) the presence of an endothelial cell-secreted unknown soluble relaxing factor (EDRF) (48). In 1987, this factor was found
to be identical with NO (11) and arginine was revealed as the
precursor for NO synthesis in cardiovascular physiology (15). In
−
parallel studies, NO−
2 and NO3 were measured as end products of
a novel oxidation pathway expressed in macrophages upon stimulation with LPS (49) or IFN-γ (50). Arginine was recognized as the
biological precursor molecule of nitrite/nitrate released from activated macrophages (10). Further studies demonstrated that NO
is an intermediate of macrophage arginine oxidation to the final
end products nitrite/nitrate (14) and that NO synthesis is required
for macrophage cytotoxic activity (22). In 1991, the enzyme converting arginine to NO was purified, cloned, and was named NOS
(now known as neuronal NOS, nNOS) (51). Shortly after this, two
additional NOS isozymes were cloned: iNOS from macrophages
(52, 53) and endothelial NOS (eNOS) (54).
The three NOS isozymes differ in structure, distribution, regulation, and synthetic capacity, but they catalyze the same reaction:
the incorporation of molecular oxygen and the release of NO
from the terminal guanidino nitrogen group of arginine and generation of citrulline as a byproduct (22). All NOS enzymes are
large homodimeric proteins with two functional domains: (i)
an N-terminal oxygenase and catalytic domain, which binds an
iron–protoporphyrin IX (heme) prosthetic group and the cofactor
tetrahydrobiopterin (BH4 ) and (ii) a C-terminal reductase domain
with binding sites for flavin mononucleotide (FMN) and flavin–
adenine dinucleotide (FAD). The catalytic reaction involves two
monooxygenation steps: (i) arginine is hydroxylated by O2 and
NADPH to form Nω -hydroxy-l-arginine, which is then (ii) oxidized to yield NO and citrulline. All three NOS isoforms can also
synthesize superoxide in the absence of arginine and BH4 . This
NOS-derived superoxide can react with NO to form peroxynitrite.
Both nNOS and eNOS are constitutively expressed enzymes and
calcium-dependent in their activity. In contrast, iNOS is regulated
via inducible transcription and synthesizes NO independent of
calcium since the essential subunit calmodulin is bound to iNOS
even at low intracellular calcium concentrations. Most prominently known as microbicidal and inflammatory effector pathway
in macrophages (22), iNOS activity has also been demonstrated in
a variety of other cell types, e.g., hepatocytes (55), pulmonary

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epithelium (56), and colon epithelium (57). A variety of proinflammatory cytokines (e.g., IL-1β, IFN-γ, or TNF-α), microbial
products (e.g., LPS), and hypoxia can induce macrophage iNOS
transcription, whereas other cytokines (e.g., IL-4, IL-10, TGF-β)
suppress iNOS gene transcription (58). Additive or synergistic
activities of combinations of multiple cytokines are most efficient
in inducing or suppressing iNOS gene expression. NO synthesis
can also be limited by arginine availability and/or on the level of
iNOS protein expression (59). NOS can further be inhibited by
endogenously produced asymmetric dimethylarginine (ADMA)
or pharmacologically by synthetic arginine analogs with substitutions at the terminal guanidino group, e.g., Nω -monomethyll-arginine (L-NMMA), Nω -nitro-l-arginine (L-NNA), or Nω nitro-l-arginine methyl ester (L-NAME). Once iNOS has been
translated, a prolonged production of potentially large amounts
of NO is detectable. NO can act via stimulation of soluble guanylate cyclase to generate cyclic GMP within the target cell. Besides
its physiological role as guanylate cyclase stimulator, NO is also
a radical with a very short half-life of approximately 3–5 s and
it reacts with a variety of molecules leading to (i) further reac−
tive nitrogen species (RNS)

like N2 O3 , peroxynitrite (ONOO ),
or nitronium ion NO+
in
the
presence
of
oxygen
radicals
(60)
2
and (ii) nitrosylated proteins with potentially altered or impaired
function.
ARGINASE: ARGININE – ORNITHINE PATHWAY

The enzyme arginase drives the second or alternative pathway of
arginine metabolism in macrophages, catalyzing the hydrolysis of
arginine to ornithine and urea. While arginase was known as an
enzyme of the hepatic urea cycle since the discovery of the latter
in 1932 by Krebs and Henseleit, it is also expressed in many nonhepatic cells. There are two isozymes of arginase (arginase I and
arginase II), which catalyze the same biochemical reaction but differ in cellular expression, cell-type-specific regulation, and subcellular localization (17, 61). Hepatic urea cycle arginase I is expressed
as a cytosolic enzyme, while human granulocyte arginase I is
found in the granular compartment (41) and arginase II is a mitochondrial enzyme (61). It was initially demonstrated that murine
macrophage arginase is inducible by PGE2 (62), Th2 cytokines, and
cAMP, both alone (62) and synergistically acting together (63). The
molecular details of this gene regulation were then clarified: the
transcription factors STAT-6 and CEBP/β assemble at an enhancer
element 3 kb upstream from the basal promoter of arginase I and
Th2 cytokine-mediated murine arginase I mRNA induction is controlled by this mechanism (64). Meanwhile, human macrophage
arginase I expression was demonstrated by synergistic induction
with cAMP increasing treatments (PDE4 inhibition) in combination with IL-4 or TGF-β (65). The molecular details of this
induction were also clarified recently in the murine RAW264.7
macrophage cell line, involving STAT-6 and CEBP/β binding to an
IL-4 response element in the arginase I promoter (66). Another
layer of complexity comes into play by the demonstration of
pathogen-induced toll-like receptor (TLR)-mediated induction of
arginase I in murine macrophages (67, 68).
Ornithine serves as a substrate for ornithine decarboxylase
(ODC), a rate-limiting enzyme in the synthesis of polyamines.
Polyamines are small, polycationic molecules, which regulate a

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Macrophage arginine metabolism

multitude of cellular processes like DNA replication, protein translation, cell growth, and differentiation (69, 70). Much less is
known about an involvement of polyamines in immune reactions:
the polyamine spermine, e.g., inhibits pro-inflammatory cytokine
synthesis of human, LPS-stimulated PBMC (71), arginine transport (72), and NO synthesis in rat (72) and murine macrophages
(73). Ornithine, via the enzyme OAT, is also a precursor amino acid
for the synthesis of proline, which itself is essential for the synthesis of collagen. Accordingly, arginase-derived ornithine might be
important in tissue (re-)modeling processes. This hypothesis was
supported by studies that demonstrated an increase in arginase
levels in fibrotic lung disease (74) or allergic asthma (75, 76).
What are the biological functions of arginase-expressing
macrophages? While there is ample evidence, at least in the murine
system, of the fundamental role of NO-producing macrophages
for infection control (22), a multitude of pathophysiological scenarios have also been described in which arginase-expressing
macrophages are key players (17). In the initial groundbreaking analysis on macrophage phenotypes during wound healing,
Mills and co-workers (77, 78) showed that arginase-expressing
and ornithine-producing macrophages were crucial for wound
healing as opposed to NO-producing macrophages, which dominated the initial phase of antibacterial inflammation. The same
two macrophage phenotypes were then also correlated with tumor
growth (arginase/ornithine) or tumor killing (NOS/NO) (79).
Based on these dramatic differences in function, the two different
macrophage populations were then named as M1 and M2, based
on their route of arginine metabolism (Figure 2) (5). In general,
M2 type macrophages act as anti-inflammatory cells (“healing”
mode) via diversion of arginine away from NOS or via the synthesis
of downstream products derived from the ornithine that is generated via arginase (see above). For illustration, we want to list just a
few, more recently published examples for the role of macrophage
or, in this context, also DC arginase in (i) infection-induced
inflammation, (ii) immune evasion in tumor and infection, (iii)
regulation of fibrosis, and (iv) direct control of parasite growth. (i)
The anti-inflammatory property of macrophage arginase during
infection was shown in murine schistosomiasis where excessive tissue injury is prevented by arginase-expressing macrophages due
to suppression of pro-inflammatory cytokines IL-12 and IL-23
and the maintenance of the Treg/Th17 balance (80). (ii) IL-6induced arginase 1 in DCs leads to downmodulation of MHC-II
and this is subsequently responsible for suppression of CD4+ T
cell-mediated antitumoral immunity (81). Immune evasion of
chronic Helicobacter pylori infection is mediated by arginase II
induction in gastric macrophages, due to inhibition of NOSmediated bacterial killing and suppression of pro-inflammatory
cytokine production (82). (iii) Macrophage arginase I restricts
Th2 cytokine driven inflammation and fibrosis in murine schistosomiasis (83) although macrophage arginase itself can be profibrogenic via direct production of proline as collagen precursor
in schistosomiasis (84). (iv) Macrophage arginase-mediated synthesis of polyamines enhances growth of intracellular Leishmania
parasites in murine macrophages (85) and arginase-expressing
granulocytes and the levels of arginase activities are markers
of disease severity in human visceral leishmaniasis and in HIV
infections (86–88).

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Rath et al.

One further important consequence of macrophage arginase
expression is reduction of extracellular arginine. This is most
likely more apparent in the immediate microenvironment of M2
macrophages due to the continuous flux of nutrients and the arginine synthetic capacity of the whole organism. Suppression of
T cell activation via arginine depletion has been studied quite
extensively in vitro (47, 89–91) and is known as one of the
prime immunosuppressive mechanisms of arginase-expressing
myeloid-derived suppressor cells (MDSC) (18). In contrast, its
role in macrophage-driven pathophysiology in vivo is still not
really clear in most relevant disease entities and needs to be analyzed in the future. We speculate that M2/arginase macrophages
might be more efficient in the induction of extracellular arginine
depletion since there is no intracellular reconstitutive mechanism
for arginine recycling for M2 macrophages as opposed to M1
macrophages, which can use the citrulline–NO rescue pathway.
Suppression of T cell activation, proliferation, and/or differentiation by macrophage arginase I was shown in vivo in murine
disease models dominated by M2 macrophages, like schistosomiasis (80, 83) or leishmaniasis (92). Interestingly, extracellular
arginine depletion has also been shown to inhibit ERK1/2 activation and subsequently pro-inflammatory cytokine production in
LPS-stimulated macrophages (93). It will be interesting to study a
potential influence of intracellular arginine depletion on potential
pro-inflammatory signaling pathways within macrophages and to
analyze if there is regulation of innate immune responses and
macrophage polarization already at such basic level. Clearly, arginine depletion does not inhibit immune responses broadly and
indiscriminately: important activation aspects of T cells (91, 94)
and granulocytes (95) are preserved in an arginine-depleted milieu
and other cellular responses, e.g., induction of arginine transport
protein CAT-1 (33), are even enhanced in eukaryotic cells under
arginine nutrient deprivation.
INTERACTIONS BETWEEN NOS AND ARGINASE PATHWAYS

Nitric oxide synthase and arginase can both antagonize or synergize in the generation of oxidative and nitrosative stress: inadequate supply of arginine (or the cofactor

tetrahydrobiopterin)
leads to the production of superoxide O−
2 instead of NO, increasing oxidative stress and also the production of peroxynitrite (22).
In general, though, mutually exclusive expression of iNOS and
arginase I in individual macrophages prevails (96) and there are
multiple cross-inhibitory interactions between the two arginine
metabolic pathways in macrophages: NOHA, the intermediate
product in NO synthesis, is a potent inhibitor of both arginase
isoforms (97). In non-macrophage cell types, it was also demonstrated that NO is an effective inhibitor of ODC via nitrosylation
of the enzyme with consecutively reduced polyamine synthesis
(98). Arginase can limit arginine availability for NO synthesis, as
demonstrated by pharmacological arginase inhibition in different
types of macrophages (99). The expression of iNOS is translationally controlled by the availability of arginine (100) and in murine
M2 macrophages, induced by the cytokine IL-13, iNOS translation,
and NO production are restricted by arginase-mediated arginine
depletion (59). Polyamines and/or polyamine aldehyde metabolites can inhibit NO synthesis in murine and rat macrophages
(101). Spermine suppresses the induction of both iNOS and

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Macrophage arginine metabolism

CAT-2B arginine transporter (72) and inhibition of ODCmediated polyamine synthesis in murine macrophages enhances
LPS-induced iNOS expression and NO synthesis (102). Spermine
also inhibits H. pylori-induced iNOS protein translation in the
RAW264.7 macrophage cell line and siRNA-mediated ODC inhibition enhances macrophage NO-mediated bacterial killing (103).
In summary, these biochemical crossregulatory interactions are
in line with the concept of two types of polarized macrophages –
M1/NOS versus M2/arginase – which are defined not only by the
intracellular fate of arginine, but – most importantly – also by its
associated functional consequences.

M1/NOS VERSUS M2/ARGINASE MACROPHAGES: NOVEL
ASPECTS
The earlier simplified scheme of “proinflammation = M1/Th1”
versus “anti-inflammation = M2/Th2” has meanwhile been clarified as one possible scenario of a more broader conceptual
framework: M1 versus M2 polarization clearly leads to opposing outcomes of inflammatory reactions, but depending on the
inflammatory or infectious context, M1 and M2 macrophages can
be central players of both pro- or anti-inflammatory reactions.
Notably, M1/M2 macrophage polarization can be driven by microbial infection or innate danger signals without any influence of
adaptive immune cells, secondarily driving the Th1/Th2 polarization of the evolving adaptive immune response (104). Microbial
stimulation of macrophages via TLRs leads to the activation of
certain transcription factors (e.g., NF-κB, AP-1), which upregulate pro-inflammatory cytokines like IFN-γ and TNF-α leading to
M1 macrophage polarization with high iNOS expression whereas
cytokines like IL-4 or IL-13 lead to STAT6 phosphorylation with
consecutive arginase expression and varying further aspects of M2
polarization (mannose receptor, Ym1, Fizz1).
Various novel aspects regarding the evolution of the M1/M2
macrophage polarization in light of the differential expression
of NOS versus arginase have been clarified recently and some
examples are summarized in the following sections:
NOVEL EXOGENOUS ENVIRONMENTAL FACTORS INFLUENCING THE
NOS/ARGINASE BALANCE

Tumor cells are known to metabolize glucose preferentially via
aerobic glycolysis, known as “Warburg phenomenon,” and this
leads to high concentrations of lactate in the tumor microenvironment. This tumor cell-derived lactic acid is a potent inducer
of arginase I expression in tumor-associated macrophages (TAM)
and these M2/arginase macrophages then foster tumor growth
(105). Since activated T cells also use aerobic glycolysis with consecutive production of lactic acid, it will be interesting to study
if the same mechanism of arginase I induction is also operative
in macrophage-T cell interactions. Gliadin, a major component
of cereal gluten, and therefore omnipresent in our daily food,
was shown to induce arginase I in human monocytes (106). A
parallel stimulation with IFN-γ leads to a reduction of cellular
arginine efflux via downregulation of the arginine export protein
y+ LAT-2 (SLC7A6), thereby increasing intracellular availability of
arginine for gliadin-induced arginase (106). Finally, growth factors have also entered the M1/M2 macrophage world: in a hamster
model of visceral leishmaniasis, macrophage arginase I expression

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Rath et al.

is driven by fibroblast growth factor-2 (FGF-2) and insulin-like
growth factor-1 (IGF-1), which both signal via STAT6 and are
amplified by co-stimulation with IL-4 (107).
INTRACELLULAR SIGNALING MODULES INVOLVED IN
M1(NOS)/M2(ARGINASE) POLARIZATION

The tyrosine phosphatase Shp2 restricts M2 macrophage polarization as demonstrated by the preferential polarization of Shp2−/−
macrophages into an M2 direction with enhanced arginase expression, associated with a better protection against schistosomiasis
(108). Also, PI3K/PTEN activity is involved in regulating arginase
expression in murine macrophages since deletion of PTEN leads
to M2 polarization via C/EBPβ and STAT3 (109). These results are
in line with earlier reports demonstrating that SHIP phosphatase
(which – like PTEN – also downregulates the PI3K pathway)
dampens M2 polarization and arginase I expression in different
types of tissue macrophages (110). Deacetylation of C/EBPβ is
required for its binding to a DNA enhancer element and its role
in IL-4-mediated arginase I induction in bone marrow-derived
murine macrophages (111).
AUTO- OR PARACRINE M1/M2 POLARIZATION

In murine macrophages, induction of arginase I by mycobacteria
is driven by an autocrine–paracrine signaling loop: TLR-MyD88
mediate induction of the cytokines IL-6, G-CSF, and IL-10, which
then induce arginase I, involving the transcription factors STAT3
and C/EBPβ (112). Respiratory syncytial virus (RSV) infection
induces production of IL-4 and IL-13 of macrophages themselves
and this leads to auto- and paracrine induction of arginase I
expression in macrophages (68). It has long been known that
murine DCs share the fundamental arginase/iNOS polarization
with macrophages (113). A novel aspect here is that DCs produce
RA, which then induces arginase I and the arginine transporter
CAT-2B in the DCs themselves (34). This RA-mediated autocrine–
paracrine induction of arginase I in DCs is induced by binding of
RA to an RA-responsive element in the 50 non-coding region of
the arginase I gene and is enhanced by known exogenous arginase
inducers like IL-4 or GM-CSF (34).
M1/M2 POLARIZATION IN VIVO

In human tuberculosis, the distribution of M1 (iNOS) and M2
(arginase) macrophages is spatially organized within granulomas: M1 macrophages can preferentially be found in the inner
region closer to viable mycobacteria, whereas a higher frequency
of M2 macrophages is detectable on the outer “healing” margins. This clearly forms an organized microenvironment in which
antibacterial (M1) responses are physically separated by M2based anti-inflammation and fibrosis from uninvolved tissue (96).
In a murine tuberculosis model, overexpression of IL-13 precipitates expansion of the M2 arginase-expressing macrophage
response to the pathogens recapitulating human pathology of
post-primary tuberculosis, while the endogenous inhibition of
arginase I expression via NOHA restrains arginase expression
and pathology (114). This is in line with an earlier study showing a disease-exacerbating role of macrophage arginase (67). In
contrast to the latter two reports, a recently published study
analyzed the role of macrophage arginase in a hypoxic model

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Macrophage arginine metabolism

of Mycobacterium tuberculosis granuloma formation, in which
iNOS-based synthesis of RNS is impaired and which likely reflects
in vivo reality. Here, it was shown that arginase I expression in
granuloma-associated macrophages restricts immune pathology
since macrophage-specific deletion of arginase I led to larger granulomas and bacterial burden load (115). These discrepant study
results clearly demonstrate the fundamental importance of the
microenvironment and the multitude of potential factors that act
on macrophages in vivo and which can simply not be mimicked
in vitro.

SUMMARY AND OUTLOOK
The concept of macrophage M1/M2 dichotomy, based on differential usage of arginine via NOS or arginase was born 25 years
ago based on analyses of healing wounds (5, 104). Since then,
we have witnessed a bewildering explosion of knowledge regarding macrophage surface markers, activation requirements, signal
transduction elements and gene regulation. Within this universe of
complexity the simple discriminator arginase versus NOS expression has not only remained valid to explain how a mammalian
organism deals with wounding (which is important enough in
itself) but also has demonstrated enormous power to better explain
and understand such diverse problems as cancer control versus cancer-induced immunosuppression, autoimmune pathology
versus preservation of tolerance to self, infection control versus
chronicity or death due to infection as well as tissue healing versus exaggerated fibrosis. Despite this scientific progress during
the last quarter-century, we have by far not reached the summit of the Everest, but rather a solid base camp I, in which to
plan and prepare the next steps. The arginase/iNOS dichotomy
of macrophage amino acid metabolism has counterparts in the
other major components of cellular metabolism: (a) genes of fatty
acid oxidation are preferentially expressed in M2 macrophages and
inhibition of fatty acid oxidation leads to an abrogation of M2
activation (116); (b) M1 macrophages preferentially use glycolysis
and glutamine anaplerosis while M2 macrophages preferentially
use oxidative metabolism (117). The emerging interconnections
between macrophage metabolism and M1/M2 polarization (118)
have recently been reviewed (119, 120) and we anticipate exciting progress in this field in the upcoming years. Another crucial
aspect of the M1/M2 dichotomy that eagerly awaits more progress
and clarification is the current discrepancy between murine and
human macrophage biology in terms of iNOS/arginase expression
and regulation (17, 42, 121). We definitely need more carefully
executed in vivo and ex vivo analyses of human macrophage activation phenotypes in diverse disease settings. This will then also
lay the foundation for targeted therapeutic intervention to harness the enormous power of the arginine metabolic phenotypes
of M1/M2 macrophage polarization.

ACKNOWLEDGMENTS
Supported by the Deutsche Forschungsgemeinschaft (MU 1547/41 to Markus Munder and Cl100/6-1 to Ellen I. Closs).

REFERENCES
1. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol (2011) 11:723–37. doi:10.1038/nri3073

October 2014 | Volume 5 | Article 532 | 19

Rath et al.

2. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism and functions. Immunity (2010) 32:593–604. doi:10.1016/j.immuni.
2010.05.007
3. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122:787–95. doi:10.1172/JCI59643
4. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental
guidelines. Immunity (2014) 41:14–20. doi:10.1016/j.immuni.2014.06.008
5. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
6. Boger RH, Bode-Boger SM. The clinical pharmacology of L-arginine. Annu Rev
Pharmacol Toxicol (2001) 41:79–99. doi:10.1146/annurev.pharmtox.41.1.79
7. Morris SM Jr. Arginine: beyond protein. Am J Clin Nutr (2006) 83:508S–12S.
8. Wu G, Morris SM Jr. Arginine metabolism: nitric oxide and beyond. Biochem
J (1998) 336:1–17.
9. Windmueller HG, Spaeth AE. Source and fate of circulating citrulline. Am
J Physiol (1981) 241:E473–80.
10. Hibbs JB Jr, Taintor RR, Vavrin Z. Macrophage cytotoxicity: role for L-arginine
deiminase and imino nitrogen oxidation to nitrite. Science (1987) 235:473–6.
doi:10.1126/science.2432665
11. Palmer RM, Ferrige AG, Moncada S. Nitric oxide release accounts for the
biological activity of endothelium-derived relaxing factor. Nature (1987)
327:524–6. doi:10.1038/327524a0
12. Ignarro LJ, Buga GM, Wood KS, Byrns RE, Chaudhuri G. Endothelium-derived
relaxing factor produced and released from artery and vein is nitric oxide. Proc
Natl Acad Sci U S A (1987) 84:9265–9. doi:10.1073/pnas.84.24.9265
13. Hibbs JB Jr, Taintor RR, Vavrin Z, Rachlin EM. Nitric oxide: a cytotoxic activated macrophage effector molecule. Biochem Biophys Res Commun (1988)
157:87–94. doi:10.1016/S0006-291X(88)80015-9
14. Marletta MA, Yoon PS, Iyengar R, Leaf CD, Wishnok JS. Macrophage oxidation
of L-arginine to nitrite and nitrate: nitric oxide is an intermediate. Biochemistry
(1988) 27:8706–11. doi:10.1021/bi00424a003
15. Palmer RM, Ashton DS, Moncada S. Vascular endothelial cells synthesize nitric
oxide from L-arginine. Nature (1988) 333:664–6. doi:10.1038/333664a0
16. Noy R, Pollard JW. Tumor-associated macrophages: from mechanisms to therapy. Immunity (2014) 41:49–61. doi:10.1016/j.immuni.2014.06.010
17. Munder M. Arginase: an emerging key player in the mammalian immune system. Br J Pharmacol (2009) 158:638–51. doi:10.1111/j.1476-5381.2009.00291.x
18. Raber P, Ochoa AC, Rodriguez PC. Metabolism of L-arginine by myeloidderived suppressor cells in cancer: mechanisms of T cell suppression and therapeutic perspectives. Immunol Invest (2012) 41:614–34. doi:10.3109/08820139.
2012.680634
19. Gabrilovich DI, Nagaraj S. Myeloid-derived suppressor cells as regulators of
the immune system. Nat Rev Immunol (2009) 9:162–74. doi:10.1038/nri2506
20. Kropf P, Baud D, Marshall SE, Munder M, Mosley A, Fuentes JM, et al. Arginase
activity mediates reversible T cell hyporesponsiveness in human pregnancy. Eur
J Immunol (2007) 37:935–45. doi:10.1002/eji.200636542
21. Muller I, Munder M, Kropf P, Hansch GM. Polymorphonuclear neutrophils
and T lymphocytes: strange bedfellows or brothers in arms? Trends Immunol
(2009) 30:522–30. doi:10.1016/j.it.2009.07.007
22. MacMicking J, Xie QW, Nathan C. Nitric oxide and macrophage function. Annu
Rev Immunol (1997) 15:323–50. doi:10.1146/annurev.immunol.15.1.323 Epub
1997/01/01. ,
23. Stuehr DJ. Mammalian nitric oxide synthases. Biochim Biophys Acta (1999)
1411:217–30. doi:10.1016/S0005-2728(99)00016-X
24. Nussler AK, Billiar TR, Liu ZZ, Morris SM Jr. Coinduction of nitric oxide
synthase and argininosuccinate synthetase in a murine macrophage cell line.
Implications for regulation of nitric oxide production. J Biol Chem (1994)
269:1257–61.
25. Baydoun AR, Bogle RG, Pearson JD, Mann GE. Discrimination between citrulline and arginine transport in activated murine macrophages: inefficient
synthesis of NO from recycling of citrulline to arginine. Br J Pharmacol (1994)
112:487–92. doi:10.1111/j.1476-5381.1994.tb13099.x
26. Mori M. Regulation of nitric oxide synthesis and apoptosis by arginase and
arginine recycling. J Nutr (2007) 137:1616S–20S.
27. Qualls JE, Subramanian C, Rafi W, Smith AM, Balouzian L, DeFreitas AA, et al.
Sustained generation of nitric oxide and control of mycobacterial infection
requires argininosuccinate synthase 1. Cell Host Microbe (2012) 12:313–23.
doi:10.1016/j.chom.2012.07.012

Frontiers in Immunology | Inflammation

Macrophage arginine metabolism

28. Yeramian A, Martin L, Serrat N, Arpa L, Soler C, Bertran J, et al. Arginine transport via cationic amino acid transporter 2 plays a critical regulatory role in classical or alternative activation of macrophages. J Immunol (2006) 176:5918–24.
doi:10.4049/jimmunol.176.10.5918
29. Rothenberg ME, Doepker MP, Lewkowich IP, Chiaramonte MG, Stringer KF,
Finkelman FD, et al. Cationic amino acid transporter 2 regulates inflammatory homeostasis in the lung. Proc Natl Acad Sci U S A (2006) 103:14895–900.
doi:10.1073/pnas.0605478103
30. Closs EI, Simon A, Vekony N, Rotmann A. Plasma membrane transporters for
arginine. J Nutr (2004) 134:2752S–9S.
31. Closs EI, Boissel JP, Habermeier A, Rotmann A. Structure and function of
cationic amino acid transporters (CATs). J Membr Biol (2006) 213:67–77.
doi:10.1007/s00232-006-0875-7
32. Fotiadis D, Kanai Y, Palacin M. The SLC3 and SLC7 families of amino acid transporters. Mol Aspects Med (2013) 34:139–58. doi:10.1016/j.mam.2012.10.007
33. Hatzoglou M, Fernandez J, Yaman I, Closs E. Regulation of cationic amino acid
transport: the story of the CAT-1 transporter. Annu Rev Nutr (2004) 24:377–99.
doi:10.1146/annurev.nutr.23.011702.073120
34. Chang J, Thangamani S, Kim MH, Ulrich B, Moris SM Jr, Kim CH. Retinoic
acid promotes the development of Arg1-expressing dendritic cells for the regulation of T-cell differentiation. Eur J Immunol (2013) 43:967–78. doi:10.1002/
eji.201242772
35. Kawahara K, Gotoh T, Oyadomari S, Kajizono M, Kuniyasu A, Ohsawa
K, et al. Co-induction of argininosuccinate synthetase, cationic amino acid
transporter-2, and nitric oxide synthase in activated murine microglial cells.
Brain Res Mol Brain Res (2001) 90:165–73. doi:10.1016/S0169-328X(01)
00100-0
36. Niese KA, Chiaramonte MG, Ellies LG, Rothenberg ME, Zimmermann N. The
cationic amino acid transporter 2 is induced in inflammatory lung models and
regulates lung fibrosis. Respir Res (2010) 11:87. doi:10.1186/1465-9921-11-87
37. Nicholson B, Manner CK, Kleeman J, MacLeod CL. Sustained nitric oxide production in macrophages requires the arginine transporter CAT2. J Biol Chem
(2001) 276:15881–5. doi:10.1074/jbc.M010030200
38. Sans-Fons MG, Yeramian A, Pereira-Lopes S, Santamaria-Babi LF, Modolell M,
Lloberas J, et al. Arginine transport is impaired in C57Bl/6 mouse macrophages
as a result of a deletion in the promoter of Slc7a2 (CAT2), and susceptibility to Leishmania infection is reduced. J Infect Dis (2013) 207:1684–93.
doi:10.1093/infdis/jit084
39. Rotoli BM, Bussolati O, Sala R, Barilli A, Talarico E, Gazzola GC, et al.
INFgamma stimulates arginine transport through system y+L in human monocytes. FEBS Lett (2004) 571:177–81. doi:10.1016/j.febslet.2004.06.086
40. Rotoli BM, Dall’asta V, Barilli A, D’Ippolito R, Tipa A, Olivieri D, et al.
Alveolar macrophages from normal subjects lack the NOS-related system
y+ for arginine transport. Am J Respir Cell Mol Biol (2007) 37:105–12.
doi:10.1165/rcmb.2006-0262OC
41. Munder M, Mollinedo F, Calafat J, Canchado J, Gil-Lamaignere C, Fuentes JM,
et al. Arginase I is constitutively expressed in human granulocytes and participates in fungicidal activity. Blood (2005) 105:2549–56. doi:10.1182/blood2004-07-2521
42. Raes G, Van den Bergh R, De Baetselier P, Ghassabeh GH, Scotton C, Locati M,
et al. Arginase-1 and Ym1 are markers for murine, but not human, alternatively
activated myeloid cells. J Immunol (2005) 174:6561. doi:10.4049/jimmunol.
174.11.6561
43. Closs EI, Scheld JS, Sharafi M, Forstermann U. Substrate supply for nitricoxide synthase in macrophages and endothelial cells: role of cationic amino
acid transporters. Mol Pharmacol (2000) 57:68–74.
44. Taylor PM. Amino acid transporters: eminences grises of nutrient signalling
mechanisms? Biochem Soc Trans (2009) 37:237–41. doi:10.1042/BST0370237
45. Kaneko S, Okuda-Ashitaka E, Ando A, Nishimura K, Igarashi K, Maeda M,
et al. Polyamines upregulate the mRNA expression of cationic amino acid
transporter-1 in human retinal pigment epithelial cells. Am J Physiol Cell Physiol
(2007) 293:C729–37. doi:10.1152/ajpcell.00018.2007
46. Rotondo R, Barisione G, Mastracci L, Grossi F, Orengo AM, Costa R, et al.
IL-8 induces exocytosis of arginase 1 by neutrophil polymorphonuclears
in nonsmall cell lung cancer. Int J Cancer (2009) 125:887–93. doi:10.1002/
ijc.24448
47. Munder M, Schneider H, Luckner C, Giese T, Langhans CD, Fuentes JM, et al.
Suppression of T-cell functions by human granulocyte arginase. Blood (2006)
108:1627–34. doi:10.1182/blood-2006-11-010389

October 2014 | Volume 5 | Article 532 | 20

Rath et al.

48. Furchgott RF. The role of endothelium in the responses of vascular smooth
muscle to drugs. Annu Rev Pharmacol Toxicol (1984) 24:175–97. doi:10.1146/
annurev.pa.24.040184.001135
49. Stuehr DJ, Marletta MA. Mammalian nitrate biosynthesis: mouse macrophages
produce nitrite and nitrate in response to Escherichia coli lipopolysaccharide.
Proc Natl Acad Sci U S A (1985) 82:7738–42. doi:10.1073/pnas.82.22.7738
50. Stuehr DJ, Marletta MA. Induction of nitrite/nitrate synthesis in murine
macrophages by BCG infection, lymphokines, or interferon-gamma. J Immunol
(1987) 139:518–25.
51. Bredt DS, Hwang PM, Glatt CE, Lowenstein C, Reed RR, Snyder SH. Cloned
and expressed nitric oxide synthase structurally resembles cytochrome P-450
reductase. Nature (1991) 351:714–8. doi:10.1038/351714a0
52. Lyons CR, Orloff GJ, Cunningham JM. Molecular cloning and functional
expression of an inducible nitric oxide synthase from a murine macrophage
cell line. J Biol Chem (1992) 267:6370–4.
53. Xie QW, Cho HJ, Calaycay J, Mumford RA, Swiderek KM, Lee TD, et al.
Cloning and characterization of inducible nitric oxide synthase from mouse
macrophages. Science (1992) 256:225–8. doi:10.1126/science.1373522
54. Michel T, Lamas S. Molecular cloning of constitutive endothelial nitric oxide
synthase: evidence for a family of related genes. J Cardiovasc Pharmacol (1992)
20:S45–9. doi:10.1097/00005344-199204002-00014
55. Geller DA, Lowenstein CJ, Shapiro RA, Nussler AK, Di Silvio M, Wang
SC, et al. Molecular cloning and expression of inducible nitric oxide synthase from human hepatocytes. Proc Natl Acad Sci U S A (1993) 90:3491–5.
doi:10.1073/pnas.90.2.522
56. Asano K, Chee CB, Gaston B, Lilly CM, Gerard C, Drazen JM, et al. Constitutive
and inducible nitric oxide synthase gene expression, regulation, and activity in
human lung epithelial cells. Proc Natl Acad Sci U S A (1994) 91:10089–93.
doi:10.1073/pnas.91.21.10089
57. Perner A, Andresen L, Normark M, Rask-Madsen J. Constitutive expression of
inducible nitric oxide synthase in the normal human colonic epithelium. Scand
J Gastroenterol (2002) 37:944–8. doi:10.1080/003655202760230919
58. Modolell M, Corraliza IM, Link F, Soler G, Eichmann K. Reciprocal regulation
of the nitric oxide synthase/arginase balance in mouse bone marrow-derived
macrophages by TH1 and TH2 cytokines. Eur J Immunol (1995) 25:1101–4.
doi:10.1002/eji.1830250436
59. El-Gayar S, Thuring-Nahler H, Pfeilschifter J, Rollinghoff M, Bogdan C.
Translational control of inducible nitric oxide synthase by IL-13 and arginine availability in inflammatory macrophages. J Immunol (2003) 171:4561–8.
doi:10.4049/jimmunol.171.9.4561
60. Ignarro LJ. Biosynthesis and metabolism of endothelium-derived nitric oxide.
Annu Rev Pharmacol Toxicol (1990) 30:535–60. doi:10.1146/annurev.pa.30.
040190.002535
61. Jenkinson CP, Grody WW, Cederbaum SD. Comparative properties of
arginases. Comp Biochem Physiol B Biochem Mol Biol (1996) 114:107–32.
doi:10.1016/0305-0491(95)02138-8
62. Corraliza IM, Soler G, Eichmann K, Modolell M. Arginase induction by
suppressors of nitric oxide synthesis (IL-4, IL-10 and PGE2) in murine
bone-marrow-derived macrophages. Biochem Biophys Res Commun (1995)
206:667–73. doi:10.1006/bbrc.1995.1094
63. Munder M, Eichmann K, Modolell M. Alternative metabolic states in murine
macrophages reflected by the nitric oxide synthase/arginase balance: competitive regulation by CD4+ T cells correlates with Th1/Th2 phenotype. J Immunol
(1998) 160:5347–54.
64. Pauleau AL, Rutschman R, Lang R, Pernis A, Watowich SS, Murray PJ.
Enhancer-mediated control of macrophage-specific arginase I expression.
J Immunol (2004) 172:7565–73. doi:10.4049/jimmunol.172.12.7565
65. Erdely A, Kepka-Lenhart D, Clark M, Zeidler-Erdely P, Poljakovic M, Calhoun
WJ, et al. Inhibition of phosphodiesterase 4 amplifies cytokine-dependent
induction of arginase in macrophages. Am J Physiol Lung Cell Mol Physiol
(2006) 290:L534–9. doi:10.1152/ajplung.00326.2005
66. Sheldon KE, Shandilya H, Kepka-Lenhart D, Poljakovic M, Ghosh A, Morris
SM Jr. Shaping the murine macrophage phenotype: IL-4 and cyclic AMP synergistically activate the arginase I promoter. J Immunol (2013) 191:2290–8.
doi:10.4049/jimmunol.1202102
67. El Kasmi KC, Qualls JE, Pesce JT, Smith AM, Thompson RW, Henao-Tamayo
M, et al. Toll-like receptor-induced arginase 1 in macrophages thwarts effective
immunity against intracellular pathogens. Nat Immunol (2008) 9:1399–406.
doi:10.1038/ni.1671

www.frontiersin.org

Macrophage arginine metabolism

68. Shirey KA, Pletneva LM, Puche AC, Keegan AD, Prince GA, Blanco JC, et al.
Control of RSV-induced lung injury by alternatively activated macrophages
is IL-4R alpha-, TLR4-, and IFN-beta-dependent. Mucosal Immunol (2010)
3:291–300. doi:10.1038/mi.2010.6
69. Tabor CW, Tabor H. Polyamines. Annu Rev Biochem (1984) 53:749–90.
doi:10.1146/annurev.bi.53.070184.003533
70. Pegg AE. Mammalian polyamine metabolism and function. IUBMB Life (2009)
61:880–94. doi:10.1002/iub.230
71. Zhang M, Caragine T, Wang H, Cohen PS, Botchkina G, Soda K, et al. Spermine
inhibits proinflammatory cytokine synthesis in human mononuclear cells: a
counterregulatory mechanism that restrains the immune response. J Exp Med
(1997) 185:1759–68. doi:10.1084/jem.185.10.1759
72. Mossner J, Hammermann R, Racke K. Concomitant down-regulation of Larginine transport and nitric oxide (NO) synthesis in rat alveolar macrophages
by the polyamine spermine. Pulm Pharmacol Ther (2001) 14:297–305. doi:10.
1006/pupt.2001.0297
73. Szabo C, Southan GJ, Thiemermann C, Vane JR. The mechanism of the
inhibitory effect of polyamines on the induction of nitric oxide synthase: role of
aldehyde metabolites. Br J Pharmacol (1994) 113:757–66. doi:10.1111/j.14765381.1994.tb17058.x
74. Kitowska K, Zakrzewicz D, Konigshoff M, Chrobak I, Grimminger F, Seeger
W, et al. Functional role and species-specific contribution of arginases in
pulmonary fibrosis. Am J Physiol Lung Cell Mol Physiol (2008) 294:L34–45.
doi:10.1152/ajplung.00007.2007
75. Zimmermann N, King NE, Laporte J, Yang M, Mishra A, Pope SM, et al.
Dissection of experimental asthma with DNA microarray analysis identifies arginase in asthma pathogenesis. J Clin Invest (2003) 111:1863–74.
doi:10.1172/JCI17912
76. Maarsingh H, Dekkers BG, Zuidhof AB, Bos IS, Menzen MH, Klein
T, et al. Increased arginase activity contributes to airway remodelling in
chronic allergic asthma. Eur Respir J (2011) 38:318–28. doi:10.1183/09031936.
00057710
77. Albina JE, Mills CD, Barbul A, Thirkill CE, Henry WL Jr, Mastrofrancesco B,
et al. Arginine metabolism in wounds. Am J Physiol (1988) 254:E459–67.
78. Albina JE, Mills CD, Henry WL Jr, Caldwell MD. Temporal expression of different pathways of 1-arginine metabolism in healing wounds. J Immunol (1990)
144:3877–80.
79. Mills CD, Shearer J, Evans R, Caldwell MD. Macrophage arginine metabolism
and the inhibition or stimulation of cancer. J Immunol (1992) 149:2709–14.
80. Herbert DR, Orekov T, Roloson A, Ilies M, Perkins C, O’Brien W, et al.
Arginase I suppresses IL-12/IL-23p40-driven intestinal inflammation during
acute schistosomiasis. J Immunol (2010) 184:6438–46. doi:10.4049/jimmunol.
0902009
81. Narita Y, Kitamura H, Wakita D, Sumida K, Masuko K, Terada S, et al. The key
role of IL-6-arginase cascade for inducing dendritic cell-dependent CD4(+)
T cell dysfunction in tumor-bearing mice. J Immunol (2013) 190:812–20.
doi:10.4049/jimmunol.1103797
82. Lewis ND, Asim M, Barry DP, de Sablet T, Singh K, Piazuelo MB, et al. Immune
evasion by Helicobacter pylori is mediated by induction of macrophage arginase
II. J Immunol (2011) 186:3632–41. doi:10.4049/jimmunol.1003431
83. Pesce JT, Ramalingam TR, Mentink-Kane MM, Wilson MS, El Kasmi KC,
Smith AM, et al. Arginase-1-expressing macrophages suppress Th2 cytokinedriven inflammation and fibrosis. PLoS Pathog (2009) 5:e1000371. doi:10.
1371/journal.ppat.1000371
84. Hesse M, Modolell M, La Flamme AC, Schito M, Fuentes JM, Cheever
AW, et al. Differential regulation of nitric oxide synthase-2 and arginase-1
by type 1/type 2 cytokines in vivo: granulomatous pathology is shaped by
the pattern of L-arginine metabolism. J Immunol (2001) 167(11):6533–44.
doi:10.4049/jimmunol.167.11.6533
85. Kropf P, Fuentes JM, Fahnrich E, Arpa L, Herath S, Weber V, et al. Arginase and
polyamine synthesis are key factors in the regulation of experimental leishmaniasis in vivo. FASEB J (2005) 19:1000–2. doi:10.1096/fj.04-3416fje
86. Abebe T, Takele Y, Weldegebreal T, Cloke T, Closs E, Corset C, et al. Arginase
activity – a marker of disease status in patients with visceral leishmaniasis
in ethiopia. PLoS Negl Trop Dis (2013) 7:e2134. doi:10.1371/journal.pntd.
0002134
87. Takele Y, Abebe T, Weldegebreal T, Hailu A, Hailu W, Hurissa Z, et al. Arginase
activity in the blood of patients with visceral leishmaniasis and HIV infection.
PLoS Negl Trop Dis (2013) 7:e1977. doi:10.1371/journal.pntd.0001977

October 2014 | Volume 5 | Article 532 | 21

Rath et al.

88. Cloke TE, Garvey L, Choi BS, Abebe T, Hailu A, Hancock M, et al. Increased
level of arginase activity correlates with disease severity in HIV-seropositive
patients. J Infect Dis (2010) 202:374–85. doi:10.1086/653736
89. Rodriguez PC, Hernandez CP, Morrow K, Sierra R, Zabaleta J, Wyczechowska
DD, et al. L-arginine deprivation regulates cyclin D3 mRNA stability in human
T cells by controlling HuR expression. J Immunol (2010) 185:5198–204.
doi:10.4049/jimmunol.1001224
90. Munder M, Choi BS, Rogers M, Kropf P. L-arginine deprivation impairs Leishmania major-specific T-cell responses. Eur J Immunol (2009) 39:2161–72.
doi:10.1002/eji.200839041
91. Feldmeyer N, Wabnitz G, Leicht S, Luckner-Minden C, Schiller M, Franz T, et al.
Arginine deficiency leads to impaired cofilin dephosphorylation in activated
human T lymphocytes. Int Immunol (2012) 24:303–13. doi:10.1093/intimm/
dxs004
92. Modolell M, Choi BS, Ryan RO, Hancock M, Titus RG, Abebe T, et al. Local suppression of T cell responses by arginase-induced L-arginine depletion in nonhealing leishmaniasis. PLoS Negl Trop Dis (2009) 3:e480. doi:10.1371/journal.
pntd.0000480
93. Mieulet V, Yan L, Choisy C, Sully K, Procter J, Kouroumalis A, et al. TPL-2mediated activation of MAPK downstream of TLR4 signaling is coupled to
arginine availability. Sci Signal (2010) 3:ra61. doi:10.1126/scisignal.2000934
94. Munder M, Engelhardt M, Knies D, Medenhoff S, Wabnitz G, Luckner-Minden
C, et al. Cytotoxicity of tumor antigen specific human T cells is unimpaired
by arginine depletion. PLoS One (2013) 8:e63521. doi:10.1371/journal.pone.
0063521
95. Kapp K, Prufer S, Michel CS, Habermeier A, Luckner-Minden C, Giese T, et al.
Granulocyte functions are independent of arginine availability. J Leukoc Biol
(2014). doi:10.1189/jlb.3AB0214-082R
96. Mattila JT, Ojo OO, Kepka-Lenhart D, Marino S, Kim JH, Eum SY, et al.
Microenvironments in Tuberculous granulomas are delineated by distinct
populations of macrophage subsets and expression of nitric oxide synthase
and arginase isoforms. J Immunol (2013) 191:773–84. doi:10.4049/jimmunol.
1300113
97. Hecker M, Nematollahi H, Hey C, Busse R, Racke K. Inhibition of arginase
by NG-hydroxy-L-arginine in alveolar macrophages: implications for the utilization of L-arginine for nitric oxide synthesis. FEBS Lett (1995) 359:251–4.
doi:10.1016/0014-5793(95)00039-C
98. Bauer PM, Fukuto JM, Buga GM, Pegg AE, Ignarro LJ. Nitric oxide inhibits
ornithine decarboxylase by S-nitrosylation. Biochem Biophys Res Commun
(1999) 262:355–8. doi:10.1006/bbrc.1999.1210
99. Hey C, Boucher JL, Vadon-Le Goff S, Ketterer G, Wessler I, Racke K. Inhibition of arginase in rat and rabbit alveolar macrophages by N omega-hydroxyD,L-indospicine, effects on L-arginine utilization by nitric oxide synthase. Br
J Pharmacol (1997) 121:395–400. doi:10.1038/sj.bjp.0701143
100. Lee J, Ryu H, Ferrante RJ, Morris SM Jr, Ratan RR. Translational control of
inducible nitric oxide synthase expression by arginine can explain the arginine paradox. Proc Natl Acad Sci U S A (2003) 100:4843–8. doi:10.1073/pnas.
0735876100
101. Southan GJ, Szabo C, Thiemermann C. Inhibition of the induction of nitric
oxide synthase by spermine is modulated by aldehyde dehydrogenase. Biochem
Biophys Res Commun (1994) 203:1638–44. doi:10.1006/bbrc.1994.2374
102. Baydoun AR, Morgan DM. Inhibition of ornithine decarboxylase potentiates
nitric oxide production in LPS-activated J774 cells. Br J Pharmacol (1998)
125:1511–6. doi:10.1038/sj.bjp.0702231
103. Bussiere FI, Chaturvedi R, Cheng Y, Gobert AP, Asim M, Blumberg DR, et al.
Spermine causes loss of innate immune response to Helicobacter pylori by
inhibition of inducible nitric-oxide synthase translation. J Biol Chem (2005)
280:2409–12. doi:10.1074/jbc.C400498200
104. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity. J Innate Immun (2014) 6:716–26. doi:10.1159/000364945
105. Colegio OR, Chu NQ, Szabo AL, Chu T, Rhebergen AM, Jairam V, et al. Functional polarization of tumour-associated macrophages by tumour-derived lactic acid. Nature (2014) 513:559–63. doi:10.1038/nature13490
106. Barilli A, Rotoli BM, Visigalli R, Dall’Asta V. Gliadin activates arginase pathway in RAW264.7 cells and in human monocytes. Biochim Biophys Acta (2014)
1842:1364–71. doi:10.1016/j.bbadis.2014.04.021

Frontiers in Immunology | Inflammation

Macrophage arginine metabolism

107. Osorio EY, Travi BL, da Cruz AM, Saldarriaga OA, Medina AA, Melby PC.
Growth factor and Th2 cytokine signaling pathways converge at STAT6 to promote arginase expression in progressive experimental visceral leishmaniasis.
PLoS Pathog (2014) 10:e1004165. doi:10.1371/journal.ppat.1004165
108. Tao B, Jin W, Xu J, Liang Z, Yao J, Zhang Y, et al. Myeloid-specific disruption
of tyrosine phosphatase shp2 promotes alternative activation of macrophages
and predisposes mice to pulmonary fibrosis. J Immunol (2014) 193:2801–11.
doi:10.4049/jimmunol.1303463
109. Sahin E, Haubenwallner S, Kuttke M, Kollmann I, Halfmann A, Dohnal AB,
et al. Macrophage PTEN regulates expression and secretion of arginase I modulating innate and adaptive immune responses. J Immunol (2014) 193:1717–27.
doi:10.4049/jimmunol.1302167
110. Rauh MJ, Ho V, Pereira C, Sham A, Sly LM, Lam V, et al. SHIP represses the generation of alternatively activated macrophages. Immunity (2005) 23:361–74.
doi:10.1016/j.immuni.2005.09.003
111. Serrat N, Pereira-Lopes S, Comalada M, Lloberas J, Celada A. Deacetylation
of C/EBPbeta is required for IL-4-induced arginase-1 expression in murine
macrophages. Eur J Immunol (2012) 42:3028–37. doi:10.1002/eji.201242413
112. Qualls JE, Neale G, Smith AM, Koo MS, DeFreitas AA, Zhang H, et al. Arginine
usage in mycobacteria-infected macrophages depends on autocrine-paracrine
cytokine signaling. Sci Signal (2010) 3:ra62. doi:10.1126/scisignal.2000955
113. Munder M, Eichmann K, Moran JM, Centeno F, Soler G, Modolell M. Th1/Th2regulated expression of arginase isoforms in murine macrophages and dendritic cells. J Immunol (1999) 163:3771–7.
114. Heitmann L, Abad Dar M, Schreiber T, Erdmann H, Behrends J, McKenzie AN,
et al. The IL-13/IL-4Ralpha axis is involved in tuberculosis-associated pathology. J Pathol (2014) 234:338–50. doi:10.1002/path.4399
115. Duque-Correa MA, Kuhl AA, Rodriguez PC, Zedler U, Schommer-Leitner S,
Rao M, et al. Macrophage arginase-1 controls bacterial growth and pathology in hypoxic tuberculosis granulomas. Proc Natl Acad Sci U S A (2014)
111:E4024–32. doi:10.1073/pnas.1408839111
116. Vats D, Mukundan L, Odegaard JI, Zhang L, Smith KL, Morel CR, et al. Oxidative metabolism and PGC-1beta attenuate macrophage-mediated inflammation. Cell Metab (2006) 4:13–24. doi:10.1016/j.cmet.2006.08.006
117. Ganeshan K, Chawla A. Metabolic regulation of immune responses. Annu Rev
Immunol (2014) 32:609–34. doi:10.1146/annurev-immunol-032713-120236
118. Lamour SD, Choi BS, Keun HC, Muller I, Saric J. Metabolic characterization
of Leishmania major infection in activated and nonactivated macrophages.
J Proteome Res (2012) 11:4211–22. doi:10.1021/pr3003358
119. Biswas SK, Mantovani A. Orchestration of metabolism by macrophages. Cell
Metab (2012) 15:432–7. doi:10.1016/j.cmet.2011.11.013
120. Ghesquiere B, Wong BW, Kuchnio A, Carmeliet P. Metabolism of stromal and immune cells in health and disease. Nature (2014) 511:167–76.
doi:10.1038/nature13312
121. Murray PJ, Wynn TA. Obstacles and opportunities for understanding
macrophage polarization. J Leukoc Biol (2011) 89:557–63. doi:10.1189/jlb.
0710409

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 24 September 2014; paper pending published: 07 October 2014; accepted: 09
October 2014; published online: 27 October 2014.
Citation: Rath M, Müller I, Kropf P, Closs EI and Munder M (2014) Metabolism via
arginase or nitric oxide synthase: two competing arginine pathways in macrophages.
Front. Immunol. 5:532. doi: 10.3389/fimmu.2014.00532
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Rath, Müller, Kropf, Closs and Munder. This is an open-access
article distributed under the terms of the Creative Commons Attribution License (CC
BY). The use, distribution or reproduction in other forums is permitted, provided the
original author(s) or licensor are credited and that the original publication in this
journal is cited, in accordance with accepted academic practice. No use, distribution or
reproduction is permitted which does not comply with these terms.

October 2014 | Volume 5 | Article 532 | 22

REVIEW ARTICLE
published: 23 September 2014
doi: 10.3389/fimmu.2014.00459

Evolution of innate immunity: clues from invertebrates via
fish to mammals
Kurt Buchmann*
Department of Veterinary Disease Biology, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Carlo Pucillo, University of Udine, Italy
Jonathan P. Rast, Sunnybrook
Research Institute, Canada
*Correspondence:
Kurt Buchmann, Department of
Veterinary Disease Biology, Faculty of
Health and Medical Sciences,
University of Copenhagen, Stigbøjlen
7, Frederiksberg C, Copenhagen
DK-1870, Denmark
e-mail: kub@sund.ku.dk

Host responses against invading pathogens are basic physiological reactions of all living
organisms. Since the appearance of the first eukaryotic cells, a series of defense mechanisms have evolved in order to secure cellular integrity, homeostasis, and survival of
the host. Invertebrates, ranging from protozoans to metazoans, possess cellular receptors, which bind to foreign elements and differentiate self from non-self. This ability is in
multicellular animals associated with presence of phagocytes, bearing different names
(amebocytes, hemocytes, coelomocytes) in various groups including animal sponges,
worms, cnidarians, mollusks, crustaceans, chelicerates, insects, and echinoderms (sea
stars and urchins). Basically, these cells have a macrophage-like appearance and function
and the repair and/or fight functions associated with these cells are prominent even at the
earliest evolutionary stage. The cells possess pathogen recognition receptors recognizing
pathogen-associated molecular patterns, which are well-conserved molecular structures
expressed by various pathogens (virus, bacteria, fungi, protozoans, helminths). Scavenger
receptors, Toll-like receptors, and Nod-like receptors (NLRs) are prominent representatives
within this group of host receptors. Following receptor–ligand binding, signal transduction initiates a complex cascade of cellular reactions, which lead to production of one or
more of a wide array of effector molecules. Cytokines take part in this orchestration of
responses even in lower invertebrates, which eventually may result in elimination or inactivation of the intruder. Important innate effector molecules are oxygen and nitrogen species,
antimicrobial peptides, lectins, fibrinogen-related peptides, leucine rich repeats (LRRs),
pentraxins, and complement-related proteins. Echinoderms represent the most developed
invertebrates and the bridge leading to the primitive chordates, cephalochordates, and
urochordates, in which many autologous genes and functions from their ancestors can
be found. They exhibit numerous variants of innate recognition and effector molecules,
which allow fast and innate responses toward diverse pathogens despite lack of adaptive
responses. The primitive vertebrates (agnathans also termed jawless fish) were the first
to supplement innate responses with adaptive elements. Thus hagfish and lampreys use
LRRs as variable lymphocyte receptors, whereas higher vertebrates [cartilaginous and bony
fishes (jawed fish), amphibians, reptiles, birds, and mammals] developed the major histocompatibility complex,T-cell receptors, and B-cell receptors (immunoglobulins) as additional
adaptive weaponry to assist innate responses. Extensive cytokine networks are recognized
in fish, but related signal molecules can be traced among invertebrates. The high specificity, antibody maturation, immunological memory, and secondary responses of adaptive
immunity were so successful that it allowed higher vertebrates to reduce the number of
variants of the innate molecules originating from both invertebrates and lower vertebrates.
Nonetheless, vertebrates combine the two arms in an intricate inter-dependent network.
Organisms at all developmental stages have, in order to survive, applied available genes and
functions of which some may have been lost or may have changed function through evolution. The molecular mechanisms involved in evolution of immune molecules, might apart
from simple base substitutions be as diverse as gene duplication, deletions, alternative
splicing, gene recombination, domain shuffling, retrotransposition, and gene conversion.
Further, variable regulation of gene expression may have played a role.
Keywords: evolution, immunity, innate immunity, adaptive immunity, invertebrates, vertebrates

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INTRODUCTION
Host responses against invading pathogens are basic physiological reactions of all living organisms. Even prokaryotes protect
themselves by use of restriction enzymes and clustered regularly
interspaced palindromic repeats (CRISPRs), being able to degrade
invading foreign pathogens (1). Since the appearance of the first
eukaryotic cells, a series of additional defense mechanisms have
evolved in order to secure cellular integrity, homeostasis, and
survival of the host. Unicellular amebae developed the ability to
phagocytose foreign material as a part of their food uptake mechanisms (2) and this basic phagocyte function is conserved in higher
invertebrates and vertebrates in which the immunological function is more evident. Discrimination between self and non-self
is also crucial for sexual functions securing genetic variation by
exchange of genes between members of the same species. Recognition of non-self in both unicellular and multicellular organisms
is based on cellular receptors allowing the host organism to bind,
engulf, and/or kill potential invaders and offenders (3). Among
the invertebrates, important groups such as protozoans (amebae,
flagellates, and ciliates), sponges (such as bath sponges), cnidarians (e.g., jellyfish), worms (e.g., platyhelminths, annelids, and
nematodes), mollusks (snails and bivalves), crustaceans (e.g., crabs
and prawns), chelicerates (spiders, mites), insects (e.g., flies), and
echinoderms (sea stars and sea urchins), are known to possess
cells with receptors, which bind to foreign elements and allow
differentiation of self and non-self (4). This ability is associated
with presence of phagocytes bearing different names in various groups (amebocytes, hemocytes, coelomocytes, granulocytes,
monocytes, macrophages), but basically they have a macrophagelike appearance and have, to a certain extent, comparable functions
(5–7). Chordate evolution was based on the usage of existing
genomes from ancestors and although deletions of significant
parts of these have occurred, it is possible to trace some main
lines from early and primitive organisms to highly developed
mammals. The most primitive chordates comprising acranians
(Amphioxus) (8–10) and tunicates (ascidians) (11) display a wide
array of innate immune functions. In the primitive vertebrates
comprising jawless fish (agnathans such as hagfish and lampreys),
these functions became combined with an extensive use of leucine
rich repeats (LRRs) as lymphocyte receptors (12, 13). With the
advent of cartilaginous and bony fish, the adaptive armament
[major histocompatibility complex (MHC), immunoglobulins, Tcell receptors, extensive cytokine networks] appeared, and these
new tools were further developed to a high level of sophistication
through amphibians, reptiles, and birds to mammals (14). This
allowed a reduction of the copy number of many innate immune
genes, but still the innate effector molecules have been taken into
a complex network combining the obvious talents of fast acting
ancient molecules with the highly developed specific recognition
with memory seen in adaptive immunity. The main outlines of
these aspects, which are presented below, highlight how innate
immune responses evolved from ancient precursors and still play
a vital and basic role even in higher vertebrates where adaptive
elements are so prominent.

THE TIME SCALE – IN BRIEF
Evolution of the animal immune system, in its broadest sense, can
be viewed over a time span of at least 1000 million years (Figure 1).
Frontiers in Immunology | Molecular Innate Immunity

Evolution of immunity

The age of the Earth has been estimated to more than 4.6 billion
years, but the first traces of life appeared later with the appearance
of primitive prokaryotes. The initial relatively inactive period is
called the Precambrian period (or Proteozoic era), and it exhibited a series of primitive single celled organisms, which could exist
in colonies, toward its end (one billion years ago). However, even
these primitive organisms may have developed defense mechanisms to preserve their integrity. The environmental conditions
prevailing then and at later stages during the Earth’s life may have
placed a strong selective pressure on the organisms. Major extinctions of existing organisms (seen several times during evolution)
may be due to harsh environmental and physiochemical changes,
which probably have played an active role in creation of mutations,
gene and chromosome deletions, duplications, and gene shuffling.
The Paleozoic era spanning the period 542–240 million years ago
(mya) was initiated by a new period called Cambrium 542 mya. At
this stage, more complex organisms such as cnidarians (including
jellyfish) were prevalent but an impressive diversification, called
the Cambrian explosion or radiation, was put in action, which
resulted in appearance of some major animal groups. Then over a
relatively short time span, the ancestors of both invertebrates and
vertebrates known today appeared. The diversification of all the
multicellular animals continued. During the following, millions of
years called the Ordovician, Silurian, and Devonian periods more
advanced invertebrates (echinoderms), chordates (ascidians and
acranians), and vertebrates (jawless and jawed vertebrates) came
into play. Thus, in this last period, jawed fishes (and thereby the
adaptive immune system) were seen for the first time around 450
mya and they were soon followed by amphibians. In the Carboniferous period (from around 350 mya), the reptiles appeared and
diversified in the Permian period (from about 300 mya). By the
end of this period, a major extinction affecting parts of all animal
groups occurred probably due to some major climate changes.
With the advent of the Mesozoic era initiated with the Triassic
period (250–200 mya), the first dinosaurs and mammals were seen.
In the Jurassic period 200–140 mya, dinosaurs radiated and birds
appeared as one lineage in this group. In the following Cretaceous
period (140–65 mya), the first primates developed, but again a
major extinction process occurred, which primarily known as the
end of the dinosaur time span. This event was followed by the
Cenozoic era including the Paleogene and Neogene periods where
further mammalian diversification took place and finally, in the
Quaternary period, humans arose around 60,000–120,000 years
ago. When dealing with innate immune mechanisms, it is thus
likely that some genes involved in the defense of the early invertebrate ancestors 5–600 mya are still playing a role in the innate
and even adaptive immune reactions of mammals. As will be suggested from the report below, invertebrate genes (immune-related
or not) may have been used as bricks directly or modified for later
and alternative use when appropriate.

DISCRIMINATION OF SELF FROM NON-SELF
Even the most primitive unicellular organism needs to discriminate self from non-self. This applies for a basic nutrition and
feeding process in which the ameba or flagellate select food items
and subsequently exert phagocytosis or pinocytosis. In addition,
genetic exchange and sexual reproduction is dependent on this
type of discrimination. It may have arisen several times during
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Buchmann

Evolution of immunity

FIGURE 1 | Evolution of the immune system is shown. Immune cells and molecules from early invertebrates to vertebrates are shown. Geological periods
and time periods (million years ago, mya) are shown with extant representatives of animal groups appearing at different time during evolution.

evolution but genetic evidence points to a conservation of several genes encoding molecules active in cell to cell interaction.
The primitive cnidarian Hydractinia has at least two innate histocompatibility loci alr1 and alr2 (15). Allorecognition and rejection
has been well studied for the colonial tunicate Botryllus schlosseri
applying a locus called FuHC (fusion/histocompatibility) associated with putative receptor proteins named fester and Uncle fester,
which are very polymorphic (16) and it was recently reported that
a polymorphic HSP40-like protein is encoded within the FuHC
locus (17). The MHC, a central element in adaptive responses,
is well established in fish but its origin in invertebrates is still
enigmatic. A common ancestral region traced in the early chordates (urochordates and cephalochordates) is referred to as the
proto-MHC. It is likely to be the first building block for the MHC,
which probably was established later in evolution by the process
of chromosome duplications (18).

EFFECTOR CELLS
The basic phagocytic ability of unicellular organisms (e.g., amebae) is also found in the most primitive multicellular animals
belonging to the group Porifera (sponges) and cnidarians (the
group including jellyfish and sea-anemones). These animals apply
phagocytic amebocytes for nutrition and recognition of foreign
elements in the environment. Similar cell types have been conserved through evolution as they are recognized in all groups
from invertebrates (annelids, arthropods, mollusks, echinoderms)
to vertebrates (4). Several terms have been assigned to these cells in
various groups and it must be expected that future investigations

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will sub-divide groups further. Sponges carry amebocytes in their
mesoglea, cnidarians possess interstitial cells with a phagocytic
function, hemocytes are found in the vascular system, and coelomocytes occur in coelomate animals. Thus, earthworms possess
several subtypes of coelomocytes including eleocytes, and granular
amebocytes (5) and in arthropods, comprising both crustaceans
and insects, several effector cell types have been characterized
(19). The evolutionary importance of corresponding phagocytes/macrophages is reflected in the range of subsets described
from invertebrates and primitive chordates. Various cell types
within this theme are found in advanced invertebrates (represented by echinoderms such as sea stars and sea urchins) and
in the cephalochordate Branchiostoma (Amphioxus) and in urochordates (tunicates, ascidians) where both granulocyte-like cells
and macrophages occur (20, 21). An even more diverse array of
cell types and subsets occur in jawless vertebrates (hagfish and
lampreys), cartilaginous fish (sharks and rays), and in bony fish.
Besides phagocytes, jawless fish possess different subsets of lymphocytes with special membrane receptors. These primitive vertebrates without jaws have evolved an alternative antigen recognition
system, which are composed of LRRs. These molecules provide
agnathans a basis for establishing various lymphocyte lines corresponding to B and T lymphocytes. However, in cartilaginous and
bony fish, the lymphocyte receptors are immunoglobulin (B-cell
receptors) or T-cell receptors whereas agnathans apply at least two
forms of variable lymphocyte receptor (VLR) based on LRR (13).
In bony fish, the cellular armament might include lymphocytes,
macrophages, monocytes, dendritic cells, neutrophils, granulocytes,

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eosinophils, basophils, mast cells, and NK-cells and an even higher
specialization is known in mammals (6, 7, 22). Leukocytes have
traditionally been divided into the myeloid and lymphoid line
based on their development from certain stem cells. However,
B-lymphocytes in rainbow trout have been shown to exert phagocytosis (23), which suggests that the border between these developmental cell lines is less rigid at least in fish. In this context, it
is interesting that the Ikaros multigene family, which take part
in vertebrate hemopoietic stem cell differentiation and production of B, T and NK cell lineages, seems to find an early version
in the most primitive vertebrates (the agnathan hagfish Myxine)
and the even earlier urochordates (the tunicate Oikopleura) (24).
The ancient origin of genes, which are central in cellular adaptive
immunity in higher vertebrates, is also reflected by the finding of a
Nuclear Factor of Activated T-cells (NFAT)-like gene in the primitive chordate Branchiostoma belcheri (Amphioxus group). In this
chordate, this gene seems to play a role in innate recognition of
lipopolysaccharide (LPS).

RECEPTORS
In order to respond to non-self and potential pathogens and
initiate phagocytosis or production of killing mechanisms, the
phagocytic cells must possess receptors, which can bind relevant
ligands. The primitive multicellular sponges possess LPS binding receptors, which can interact with structural polysaccharides
(beta-glucan) from fungi (25). This group has also been reported
to express intracellular receptors nucleotide-binding domain and
LRR (NLR) (26) (also termed the nucleotide-binding oligomerization domain receptors, Nod-like receptors), which bind bacterial or viral RNA, flagellin, and peptidoglycan leaving the host
cell with an ability to fight pathogens or pathogen-related molecules, which have managed to enter the cytosol (26). RIG-like
receptors (RLR) are able to bind viral RNA and establish innate
defense reactions and their ancestral form seems to occur shortly
before the first vertebrates evolved (27). These are all examples
of pathogen-recognition receptors (PRRs) recognizing pathogenassociated molecular patterns (PAMPs), which are well-conserved
molecular structures expressed by various pathogens (virus, bacteria, fungi, protozoans, helminths). PAMPs may among others be
LPS, peptidoglycans, flagellin, double-strand RNA (dsRNA), and
structural carbohydrates. The term damage-associated molecular
patterns (DAMPs) are being used to signify the danger reflected
by presence of cell constituents released to the extracellular milieu
following tissue injury. Toll-like receptors (TLRs) play a major
role within this group of host receptors. They are composed of
an extracellular domain bearing LRRs and a cytoplasmic domain
(interleukin-1 receptor like). Following receptor–ligand binding,
signal transduction initiates a complex cascade of reactions, which
leads to production of one or more of a wide array of effector molecules eventually resulting in elimination or inactivation
of the intruder. A large number of TLRs are known with individual affinities to various PAMPs (28). TLRs have been traced
to the most ancient multicellular invertebrates such as sponges,
cnidarians (29), oligochaetes (earthworms) (30), mollusks (snails
and mussels) (31), crustaceans (e.g., shrimps), and insects (32).
The echinoderms, representing the most developed invertebrates,
exhibit a complex and rich array of innate recognition molecules

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Evolution of immunity

where among TLRs are present in numerous copies (33). The most
primitive fish, the agnathans, have at least 7 identified TLRs, bony
fish at least 18, amphibians 14, birds 10, and mammals 13 (28). One
major receptor group comprises the scavenger receptors binding
bacteria and a range of antigens including lipoproteins, which are
polyanionic (34). They are ancient receptors occurring on most
cells in sponges, the most primitive multicellular animals. They
have a cysteine rich domain (SRCR), which can be traced through
insects, echinoderms, early chordates, and fish (35).

EFFECTOR MOLECULES
Invertebrates exhibit a rich variation of innate immune molecules
allowing recognition, pathogen binding, and pathogen killing (16).
Sponges apply oxidative killing processes based on production
of reactive O- (ROS) and N-(NOS) species. Gastropods (snails)
exemplified by Biomphalaria glabrata are able to produce ROS
when exposed to one or more carbohydrate ligands (36) and
NOS when infected by sporocysts of the digenean trematode
Schistosoma mansoni (37). It is not clear if these animals possess
preformed molecules (or enzymes), which are released immediately upon stimulation in order to exert their function instantly.
Agglutination, clotting, and coagulation are other effective methods used to inactivate, and combat intruders and mollusks apply
fibrinogen-related peptides (FREPS) as central players in the
process. Melanization is another innate response mechanism in
which pathogens are encapsulated and inactivated by reaction
products including cytotoxic quinones and reactive O- and Nspecies. Melanin itself may protect against light and ionization
and the prophenoloxidase system is an enzyme complex associated with these reactions (3). Many similar mechanisms have
been extensively studied in fish in which inducible NO synthase is
readily expressed following parasite infection (38). Other innate
factors produced by fish include antimicrobial peptides (AMP),
lysozyme, hemolysins, transferrins, lectins (MBL), SAA, SAP, CRP,
and complement factors (39). The complement system, which is
linking innate and adaptive responses in vertebrates, can be traced
even in primitive invertebrates such as cnidarians (40) but exhibit
the most complex cascade reactions in vertebrates. The function
and interactions between the individual complement factors in
lower chordates and invertebrates are unexplored and probably
differ from the cascade reactions known from higher vertebrates
(41). With the advent of cartilaginous and bony fishes, the adaptive immune system found its basic form including the ability to
produce various classes of functional immunoglobulins. Although
immunoglobulin-like sequences have been found in invertebrates,
the high specificity and re-arrangement of V, D, and J domains
associated with antibodies was first seen in these fish groups. The
recombination activating genes RAG1 and RAG2 (RAGs) play
a central role in this process and it is noteworthy that RAGlike sequence genes have been recognized in the early chordate
Amphioxus (10). This adds to the notion that some immunerelated genes in invertebrates and early chordates have had other
functions before the adaptive immune system evolved.

SIGNAL MOLECULES
Coordination of cellular processes must be an integrated function even in the most primitive multicellular animals in order to

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maintain shape, structure, and function. Orchestration of complex
reactions is carried out by various cytokines. Such molecules have
been described in primitive invertebrates and although many of
these may not be homologous to vertebrate cytokines, several studies have shown effects on the immune reactivity in invertebrates
following stimulation with recombinant vertebrate cytokines.
Thus, TNF-alpha, IFN-γ, and IL-8, have been demonstrated to
induce reactions in worms, mollusks, and insects suggesting that
these animals apply interleukin-like signal molecules (42–44).
Earthworm coelomocytes responded to recombinant human IL12 and IFN-γ by increasing phagocytosis (43) and Blue mussel
hemocytes responded to TNF-α stimulation by increased stress
reaction and decreased phagocytosis (42). Likewise, insect (fruitfly) cells were stimulated by recombinant human IL-8, which is
associated with increase of phagocytic cells and subsequent expression of insect cytokines upd-3 and dhf (44). However, based on
the fact that corresponding genes have not yet been described in
these invertebrates it must be framed that these results should be
observed with some caution.
However, some cytokines have been found encoded in the
genome of certain invertebrates. A central regulating molecule is
TGF-β, which may secure moderation of inflammation and initiate
and sustain repair functions. It belongs to a family with numerous members in mollusks, nematodes, insects, echinoderms, and
tunicates. Even the genome of cnidarians represented by the sea
anemone Aiptasia pallida contains genes encoding TGF-β, and
it was demonstrated experimentally that this cytokine depressed
immune reactions including nitric oxide production (45). Another
central cytokine is the macrophage migration inhibitory factor
(MIF), which was released following infection with the digenean
trematode parasite Schistosoma mansoni (46). MIF has also been
described from the Pacific white shrimp Litopenaeus vannamei in
which it functions as a prominent pro-inflammatory cytokine,
which is up-regulated following viral infections (47) and predominantly expressed in blood cells, heart, and hepatopancreas.
The Pacific oyster genome encodes an IL-17 like cytokine, which
is highly expressed following injection with pathogenic bacteria
(48). The cytokine allograft inflammatory factor-1 (AIF-1) has
been described from the same host (49). It was found to stimulate phagocytic activity of oyster granulocytes. Crustaceans such
as the freshwater crayfish produce a series of astakine cytokines
(50–52), which have impact on hematopoiesis. The Chinese mitten crab produces suppressors of cytokine signaling (SOCS2) in
various cells and organs following challenge with pathogenic bacteria (53). Fruitflies produce various cytokines including helical
cytokines (44). In more developed invertebrates (echinoderms)
(33) and primitive chordates (8), corresponding signal molecules
have been described. The LPS-induced TNF-α factor (LITAF) gene
was recently detected in Amphioxus (8) where it functions not only
as a transcription factor for expression of TNF-α but also may be
regulating innate responses in general. In lampreys, one of the most
primitive vertebrates, a tumor protein homolog has been found
to regulate cytokine secretion from various leukocytes (54). Our
knowledge within cytokine evolution has recently been expanded
particularly with regard to fish. Thus, IL-1, IL-2, IL4/13, IL-6, IL7, IL-8, IL-10, IL-12, IL-13, IL-15, IL-17, IL-21, IL-22, IL-25, and
IL-35 have been recognized in bony fish (55) and corresponding

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Evolution of immunity

arrays might be expected to occur in cartilaginous fish (sharks and
rays) (22). It should be framed that although sequence similarities
suggest that lines of development from primitive animals (invertebrates and chordates) to higher vertebrates exist, one should be
open for change of function of gene products during evolution.
Thus, regulation of cellular communication may apply different
cytokines at different stages even for corresponding functions.

EVOLUTION OF MACROPHAGE FUNCTION
Macrophage function in a higher vertebrate host organism may
be directed along different pathways characterized as M1 and M2
functions (56). These lines are specialized in “Fight” or “Repair”
systems, respectively, related at least partly to cellular use of Arginine. This amino acid can be converted to nitric oxide (NO) by
inducible nitric oxide synthase (iNOS) or to Ornithine and Urea
by Arginase. The former reaction (previously termed classical activation) makes the macrophage capable of fighting and killing
invading microbes by use of the reactive N-species whereas the
latter (alternative activation) can be characterized as the repairing
pathway. In higher vertebrates, M1 and M2 cells are associated with
expression of different cytokine profiles but it cannot be excluded
that these two phagocyte functions are regulated in a special way
in invertebrates. Evidence has been produced that this division of
macrophage function may occur in fish. In rainbow trout putative macrophages (MHCII positive cells) are found widespread
in various tissues even in the early yolk sac larva (57). Infections
with Ichthyophthirus multifiliis (a ciliated skin and gill parasite)
elicit expression of iNOS in rainbow trout (38, 39) and Myxobolus
cerebralis infection lead to iNOS or Arginase-2 expression dependent on the susceptibility of the rainbow trout strain used (58).
In addition, salmon louse Lepeophtheirus salmonis infections of
Atlantic salmon skin was associated with an upregulation of the
arginase gene (59). A related switch from a Th1 to a Th2-like reaction in rainbow trout skin infected with flagellates was recently
described (60). So although M1 and M2 differentiation has not
yet been detected in invertebrates, at least fish seems to have developed arginase, which makes M1 and M2 differentiation possible.
Thus, Arginase is found in only one form in micro-organisms and
invertebrates, a form which is not related to the ornithine–urea
cycle, whereas fish may possess the necessary enzymes (61).

CONNECTING INNATE AND ADAPTIVE RESPONSES
It was with the appearance of the vertebrates that a higher
degree of immunological sophistication (adaptive immunity) was
evolved. Vertebrates developed the MHC, T-cell receptors, and
immunoglobulins as an additional weapon and regulatory system.
The most primitive fish (agnathans such as hagfish and lampreys)
possess special lymphocyte receptors composed of leucine reach
repeats suggesting that this group followed a divergent line of
development. With the cartilaginous fish (sharks and rays) and
bony fish, immunoglobulins appeared. Some modern fish today
carry at least three classes of immunoglobulins [IgM, IgT (Z),
IgD], an array, which has been further developed in amphibians (IgM, IgX, IgY, IgD, IgF), reptiles (IgM, IgY, IgA, IgD), birds
(IgM, IgY, IgA, IgD), and mammals (IgM, IgG, IgA, IgD, IgE) (62).
Despite the lack of these specialized proteins (immunoglobulins)
in lower vertebrates and invertebrates, this does not mean that

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Buchmann

immunity is less well developed in primitive animals. In fact, a
rich array of innate immune genes and high variability of innate
effector molecules provide animals such as earthworms, snails,
mussels, shrimps, and insects with a capability to combat continuous attacks from microbes in their environment. Although central
parts of these innate immune mechanisms present in invertebrates
are conserved in higher vertebrates, it seems that the variability
and diversity is much higher among invertebrates whereas higher
vertebrates by fine-tuning the adaptive components (Igs, TCR,
MHC) reach the same goal of clearing pathogens from the host
organism. It may be hypothesized that the efficacy of the adaptive
weaponry has allowed vertebrates to reduce the often impressing
variety of innate effector molecules, which was available in earlier
lineages.

COEVOLUTION OF PARASITE AND HOST AS AND
ADDITIONAL DRIVER OF INNATE IMMUNITY VARIATION
It is evident that ancestors of existing pathogens have been able
to evade innate and adaptive host immune mechanisms. Thus,
immune reactions against viral, bacterial, and parasitic pathogens
are in most cases only partly effective with regard to elimination
of the intruding or established parasite in the vertebrate host (63,
64). It is a characteristic trait of both protozoan and metazoan
parasites that the pathogens are able to deal with extensive cellular
and humoral elements of the host immune system, a trait, which is
securing parasite survival for extended periods (65). Coevolution
of hosts and parasites has resulted in a tight interaction between
innate and adaptive immune elements in the host and a rich but,
to a certain extent, unexplored array of immune evasion mechanisms in the parasites. Also bacteria and virus apply an intricate
system of immune evading mechanisms during invasion in order
to survive host defenses (66, 67). Consequently, hosts may only
survive, reproduce, and contribute to evolution by exhibiting new
and more efficient immune molecules. This never ending arms
race may be speculated to be at least partly responsible for the
presence in modern times of an immense number of both hosts
and parasites (68). However, in order to understand the principles of parasite immune evasion in higher vertebrates, including
humans, it may be speculated that the basis for evasion will be
found primarily in primitive invertebrates (16). Secondarily, we
may trace it in the oldest and most original hosts possessing an
adaptive immune system (12).

CONCLUSION
Immune factors and recognition systems involved in differentiation of self from non-self may have been an integrated part of
animal physiology since multicellular animals developed more
than 600 mya. These innate mechanisms differ from the MHC
system arising with the vertebrate lineage. Receptors, ligands, and
signal molecules may initiate relevant actions by use of a series
of effector molecules, which lead to elimination of pathogens or
re-establishment of the injured tissue in the individual. These
basic elements have been found even in sponges and cnidarians, two ancient invertebrate groups. The immune molecules
and cellular products involved in these reactions are encoded by
genes, which have similarities with elements even in higher vertebrates. Mollusks, crustaceans, insects, and echinoderms make use

Frontiers in Immunology | Molecular Innate Immunity

Evolution of immunity

of cytokine like molecules resembling TGF, MIF, TNF, and interleukins. In addition, receptor molecules (TLRs), complement, and
immunoglobulin-like sequences are being used by these invertebrates for various purposes. However, it is likely that although
many immune genes and effector molecules can be found in
the early invertebrates, their mode of action may differ considerably from the corresponding reactions in vertebrates. It is
even likely that genes encoding factors with non-immunological
roles in invertebrates may be used for immunological purposes
in higher vertebrates, and vice versa. The dramatic environmental events on the geological time scale, with several periods of
climate changes and extinction of major animal groups, have
created a basis for selection of a multitude of new variants. Interactions with pathogens, which continuously are developing immune
evasion mechanisms in their encounter with the host immune
system, may further stimulate the never ending evolution of the
immune system. The phagocyte function, taken by macrophages
in vertebrates, has also been present in the earliest invertebrates.
Corresponding cells have reached increasingly sophisticated levels during invertebrate evolution, and in vertebrates they exhibit
high diversity. These cells have, in vertebrates, been equipped with
MHC II molecules, which make them indispensable partners for
B- and T-lymphocytes. They have obtained the ability to produce and communicate through an extensive cytokine network
and they seem to be able to take a fight or repair function on
their own reactions, which were seen also in the early invertebrates. In brief, immune reaction building blocks are ancient and
appeared at various stages during evolution. Some were lost, some
were moderated, and some even obtained another function during
evolution. When adaptive immunity evolved with the vertebrate
lineage, the old and still existing elements were further incorporated in the new hosts for optimization of immunity under the
new conditions.

REFERENCES
1. Dunin-Horkawicz S, Klaus KO, Lupas AN. Prokaryotic protein networks
mediating innate immunity and apoptosis. J Mol Biol (2014) 426:1568–82.
doi:10.1016/j.jmb.2013.11.030
2. Desjardins M, Houde M, Gagnon E. Phagocytosis: the convoluted way from
nutrition to adaptive immunity. Immunol Rev (2005) 2007:158–67. doi:10.1111/
j.0105-2896.2005.00319.x
3. Cooper EL. Evolution of immune systems from self/not self to danger to artificial immune systems (AIS). Phys Life Rev (2010) 7:55–78. doi:10.1016/j.plrev.
2009.12.001
4. Dzik JM. The ancestry and cumulative evolution of immune reactions. Acta
Biochim Pol (2010) 57:443–66.
5. Bilej M, De Baetselier P, Beschin A. Antimicrobial defense of the earthworm.
Folia Microbiol (2000) 45(4):283–300. doi:10.1007/BF02817549
6. Reite OB, Evensen O. Inflammatory cells of teleostean fish: a review focusing
on mast cells/eosinophilic granule cells and rodlet cells. Fish Shellfish Immunol
(2006) 20:192–208. doi:10.1016/j.fsi.2005.01.012
7. Wittamer V, Bertrand JY, Gutschow PW, Traver D. Characterization of the
mononuclear phagocyte system in zebrafish. Blood (2011) 117:7126–35. doi:
10.1182/blood-2010-11-321448
8. Jin P, Hu J, Qian JJ, Chen LM, Xu XF, Ma F. Identification and characterization of a putative lipopolysaccharide-induced TNF-alpha factor (LITAF) gene
from Amphioxus (Branchiostoma belcheri): an insight into the innate immunity of Amphioxus and the evolution of LITAF. Fish Shellfish Immunol (2012)
32:1223–8. doi:10.1016/j.fsi.2012.03.030
9. Song XJ, Hu J, Jin P, Chen LM, Ma F. Identification and evolution of an
NFAT gene involving Branchiostoma belcheri innate immunity. Genomics (2013)
102:355–62. doi:10.1016/j.ygeno.2013.04.019

September 2014 | Volume 5 | Article 459 | 28

Buchmann

10. Zhang YN, Xu K, Deng AQ, Fu X, Xu AL, Liu X. An Amphioxus RAG1-like DNA
fragment encodes a functional central domain of vertebrate core RAG1. Proc
Natl Acad Sci U S A (2014) 11:397–402. doi:10.1073/pnas.1318843111
11. Ryzhakov G, Blazek KM, Udalova IA. Evolution of vertebrate immunity:
sequence and functional analysis of the SEFIR domain family member Act1.
J Mol Evol (2011) 72:521–30. doi:10.1007/s00239-011-9450-7
12. Litman GW, Rast JP, Fugmann SD. The origins of vertebrate adaptive immunity.
Nat Rev (2010) 10:543–53. doi:10.1038/nri2807
13. Kishishita N, Nagawa F. Evolution of adaptive immunity: implications of a third
lymphocyte lineage in lampreys. Bioessays (2014) 36:244–50. doi:10.1002/bies.
201300145
14. Zimmerman LM, Vogel LA, Bowden RM. Understanding the vertebrate immune
system: insights from the reptilian perspective. J Exp Biol (2010) 213:661–71.
doi:10.1242/jeb.038315
15. Dishaw LJ, Litman GW. Invertebrate allorecognition: the origins of histocompatibility. Curr Biol (2009) 19(7):R286–8. doi:10.1016/j.cub.2009.02.035
16. Gosh J, Lun CM, Majeske AJ, Sacchi S, Schrankel CS, Smith LC. Invertebrate
immune diversity. Dev Comp Immunol (2011) 35:959–74. doi:10.1016/j.dci.
2010.12.009
17. Nydam ML, Hoang TA, Shanley KM, De Tomaso AW. Molecular evolution of a polymorphic HSP40-like protein encoded in the histocompatibility
locus of an invertebrate chordate. Dev Comp Immunol (2013) 41(2):128–36.
doi:10.1016/j.dci.2013.03.004
18. Kasahara M, Suzuki T, Du Pasquier L. On the origins of the adaptive immune
system: novel insights from invertebrates and cold-blooded vertebrates. Trends
Immunol (2004) 25(2):105–11. doi:10.1016/j.it.2003.11.005
19. Salazar-Jaramillo L, Paspati A, Van de Zande L, Vermeulen CJ, Schwander T,
Wertheim B. Evolution of a cellular immune response in Drosophila: a phenotypic and genomic comparative analysis. Genome Biol Evol (2014) 6:273–89.
doi:10.1093/gbe/evu012
20. Rhodes CP, Ratcliffe NA, Rowley AF. Presence of coelomocytes in the primitive
chordate Amphioxus (Branchiostoma lanceolatum). Science (1982) 217:263–5.
doi:10.1126/science.7089565
21. Crivatello E, Ribatti D. The mast cell: an evolutionary perspective. Biol Rev
(2010) 85(2):347–60. doi:10.1111/j.1469-185X.2009.00105.x
22. Venkatesh B, Lee AP, Ravi V, Maurya AK, Lian MM, Swann JB, et al. Elephant
shark genome provides unique insights into gnathostome evolution. Nature
(2014) 505:174–9. doi:10.1038/nature12826
23. Zhang Y-A, Salinas I, Li J, Parra D, Bjork S, Xu Z, et al. IgT, a primitive
immunoglobulin class specialized in mucosal immunity. Nat Immunol (2010)
11:827–36. doi:10.1038/ni.1913
24. Cupit PM, Hansen JD, McCarthy AS, White G, Chioda M, Spada F, et al. Ikaros
family members from the agnathan Myxine glutinosa and the urochordata Oikopleura dioica: emergence of an essential transcription factor for adaptive immunity. J Immunol (2003) 171:6006–13. doi:10.4049/jimmunol.171.11.6006
25. Perovic-Ottstadt S, Adell T, Proksch P, Wiens M, Korshev M, Gamulin V, et al.
A (1-3) beta recognition protein from the sponge Suberites domuncula. Mediated activation of fibrinogen related protein and epidermal growth factor gene
expression. Eur J Biochem (2004) 271:1924–37. doi:10.1111/j.1432-1033.2004.
04102.x
26. Yuen B, Bayes JM, Degnan SM. The characterization of sponge NLRs provides
insight into the origin and evolution of this innate immune gene family in
animals. Mol Biol Evol (2013) 31:106–20. doi:10.1093/molbev/mst174
27. Mukherjee K, Korithoski B, Kolaczkowski B. Ancient origins of vertebrate specific innate antiviral immunity. Mol Biol Evol (2014) 31:140–53. doi:10.1093/
molbev/mst184
28. Boudinot P, Zou J, Ota T, Buonocore F, Scapigliati G, Canapa G, et al. A tetrapodlike repertoire of innate Immune receptors and effectors for coelacanths. J Exp
Zool B Mol Dev Evol (2014) 9999B:1–23. doi:10.1002/jez.b.22559
29. Franzenburg S, Fraune S, Kunzel S, Baines JF, Domazet-Loso T, Bosch TCG. My
D88-deficient Hydra reveal an ancient function of TLR signaling in sensing in
sensing bacterial colonizers. Proc Natl Acad Sci U S A (2012) 109(47):19374–9.
doi:10.1073/pnas.1213110109
30. Skanta F, Roubalova R, Dvorak J, Prochazkova P, Bilej M. Molecular cloning and
expression of TLR in the Eisenia andrei earthworm. Dev Comp Immunol (2013)
41(4):694–702. doi:10.1016/j.dci.2013.08.009
31. Elvitigala DAS, Premachandra HKA, Whang I, Nam B-H, Lee J. Molecular
insights of the first gastropod TLR counterpart from disk abalone (Haliotis

www.frontiersin.org

Evolution of immunity

32.
33.
34.

35.

36.

37.

38.

39.

40.

41.
42.

43.

44.

45.

46.

47.

48.

49.

50.

discus discus) revealing its transcriptional modulation under pathogenic stress.
Fish Shellfish Immunol (2013) 35:334–42. doi:10.1016/j.fsi.2013.04.031
Hauton C. The scope of the crustacean immune system for disease control.
J Invertebr Pathol (2012) 110:251–60. doi:10.1016/j.jip.2012.03.005
Meng FY, Mai KS, Ma HM, Zhang WB. The evolution of echinoderm immunology. Prog Biochem Biophys (2009) 36(7):803–9.
Peiser L, Mukhopadhyay S, Gordon S. Scavenger receptors in
innate immunity. Curr Opin Immunol (2002) 14:123–8. doi:10.1016/S09527915(01)00307-7
Kaur H, Jaso-Friedmann L, Evans DL. Identification of a scavenger receptor homologue on non-specific cytotoxic cells and evidence for binding to
oligodeoxyguanosine. Fish Shellfish Immunol (2003) 15:169–81. doi:10.1016/
S1050-4648(02)00156-0
Hahn UK, Bender RC, Bayne CJ. Production of reactive oxygen species by hemocytes of Biomphalaria glabrata: carbohydrate-specific stimulation. Dev Comp
Immunol (2000) 24:531–41. doi:10.1016/S0145-305X(00)00017-3
Hahn UK, Bender RC, Bayne CJ. Involvement of nitric oxide in killing
Schistosoma mansoni sporocysts by hemocytes from resistant Biomphalaria
glabrata. J Parasitol (2001) 87(4):778–85. doi:10.1645/0022-3395(2001)
087[0778:IONOIK]2.0.CO;2
Sigh J, Lindenstrøm T, Buchmann K. The parasitic ciliate Ichthyophthirius
multifiliis induces expression of immune relevant genes in rainbow trout,
Oncorhynchus mykiss (Walbaum). J Fish Dis (2004) 27:409–17. doi:10.1111/j.
1365-2761.2004.00558.x
Chettri JK, Holten-Andersen L, Raida MK, Kania P, Buchmann K. PAMP
induced expression of immune relevant genes in head kidney leukocytes of
rainbow trout (Oncorhynchus mykiss). Dev Comp Immunol (2011) 35:476–82.
doi:10.1016/j.dci.2010.12.001
Fujito NT, Sugimoto S, Nonaka M. Evolution of thioester containing proteins revealed by cloning and characterization of their genes from a cnidarian sea anemone, Haliplanella lineata. Dev Comp Immunol (2010) 34:775–84.
doi:10.1016/j.dci.2010.02.011
Nonaka M, Yoshizaki F. Primitive complement system of invertebrates. Immunol
Rev (2004) 198:203–15. doi:10.1111/j.0105-2896.2004.00118.x
Betti M, Ciacci C, Lorusso LC, Canonico B, Falcioni T, Gallo G, et al. Effects
of tumour necrosis factor alpha (TNFLx) on Mytilus haemocytes: role of
stress-activated mitogen-activated protein kinases (MAPKs). Biol Cell (2006)
98:233–44. doi:10.1042/BC20050049
Fuller-Espie S. Vertebrate cytokines interleukin12 and gamma interferon, but not
interleukin 10, enhance phagocytosis in the annelid Eisenia hortensis. J Invertebr
Pathol (2010) 104:119–24. doi:10.1016/j.jip.2010.02.009
Malagoli D, Sacchi S, Ottaviandi E. Unpaired (upd)-3 expression and other
immune-related functions are stimulated by interleukin-8 in Drosophila
melanogaster SL2 cell line. Cytokine (2008) 44:269–74. doi:10.1016/j.cyto.2008.
08.011
Detournay O, Schnitzler CE, Poole A, Weis VM. Regulation of cnidariandinoflagellate mutualisms: evidence that activation of a host TGF beta innate
immune pathway promotes tolerance to the symbiont. Dev Comp Immunol
(2012) 38(4):525–37. doi:10.1016/j.dci.2012.08.008
Garcia AB, Pierce RJ, Gourbal B, Werkmeister E, Colinet D, Reichart JM, et al.
Involvement of the cytokine MIF in the snail host immune response to the parasite Schistosoma mansoni. PLoS Pathog (2010) 6:e1001115. doi:10.1371/journal.
ppat.1001115
Zeng DG, Lei AY, Chen XH. Cloning, characterization and expression of the
macrophage migration inhibitory factor gene from Pacific white shrimp Litopenaeus vannamei (Penaeidae). Genet Mol Res (2013) 12:5872–9. doi:10.4238/
2013.November.22.15
Roberts S, Gueguen Y, de Lorgeril J, Goetz F. Rapid accumulation of an interleukin 17 homolog transcript in Crassostrea gigas hemocytes following bacterial
exposure. Dev Comp Immunol (2008) 32(9):1099–104. doi:10.1016/j.dci.2008.
02.006
Zhang Y, Li J, Yu F, He XC, Yu ZN. Allograft inflammatory factor-1 stimulates hemocyte immune activation by enhancing phagocytosis and expression
of inflammatory cytokines in Crassostrea gigas. Fish Shellfish Immunol (2013)
34:1071–7. doi:10.1016/j.fsi.2013.01.014
Lin XH, Novotny M, Söderhäll K, Söderhäll I. Ancient cytokines, the role of
astakines as hematopoietic growth factors. J Biol Chem (2010) 285:28577–86.
doi:10.1074/jbc.M110.138560

September 2014 | Volume 5 | Article 459 | 29

Buchmann

51. Lin XH, Söderhäll I. Crustacean hematopoiesis and the astakine cytokines. Blood
(2011) 117:6417–24. doi:10.1182/blood-2010-11-320614
52. Watthanasororut A, Söderhäll K, Jiravanichpaisal P, Söderhäll I. An ancient
cytokine, astakine, mediates circadian regulation of invertebrate hematopoiesis.
Cell Mol Life Sci (2011) 68(2):315–23. doi:10.1007/s00018-010-0458-8
53. Zhang Y, Zhao JM, Zhang H, Gai YC, Wang LL, Li FM, et al. The involvement
of suppressors of cytokine signaling 2 (SOCS2) in immune defense responses
of Chinese mitten crab Eriocheir sinensis. Dev Comp Immunol (2010) 34:42–8.
doi:10.1016/j.dci.2009.08.001
54. Sun J, Wu Y, Wang H, Ma F, Liu XW, Li Q. Novel translationally controlled
tumor protein homologue in the buccal gland secretion of Lampetra japonica.
Biochemie (2008) 90(11–12):1760–8. doi:10.1016/j.biochi.2008.08.002
55. Wang T, Secombes CJ. The cytokine networks of adaptive immunity in fish. Fish
Shellfish Immunol (2013) 35:1703–18. doi:10.1016/j.fsi.2013.08.030
56. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(06):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
57. Heinecke R, Buchmann K. Adaptive and innate immune molecules in developing rainbow trout, Oncorhynchus mykiss eggs and larvae: expression of genes
and occurrence of effector molecules. Fish Shellfish Immunol (2014) 38:25–33.
doi:10.1016/j.fsi.2014.02.010
58. Severin VI, Soliman H, El-Matbouli M. Expression of immune-regulatory genes,
arginase-2 and inducible nitric oxide synthase (iNOS), in two rainbow trout
(Oncorhynchus mykiss) strains following exposure to Myxobolus cerebralis. Parasitol Res (2010) 106(2):325–34. doi:10.1007/s00436-009-1661-0
59. Skugor S, Glover KA, Nilsen F. Local and systemic gene expression responses of
Atlantic salmon (Salmo salar L.) to infection with the salmon louse (Lepeophtheirus salmonis). BMC Genomics (2008) 9:498. doi:10.1186/1471-2164-9-498
60. Chettri JK, Kuhn JA, Jaafar RM, Kania PW, Møller OS, Buchmann K. Epidermal response of rainbow trout to Ichthyobodo necator: immunohistochemical
and gene expression studies indicate a Th1/Th2-like switch. J Fish Dis (2014)
37:771–83. doi:10.1111/jfd.12169
61. Srivastava S, Ratha BK. Does fish represent an intermediate stage in the evolution of ureotelic cytosolic arginase I? Biochem Biophys Res Commun (2010)
391(1):1–5. doi:10.1016/j.bbrc.2009.11.018
62. Kaiser P. Advances in avian immunology – prospects for disease control: a review.
Avian Pathol (2010) 39(5):309–24. doi:10.1080/03079457.2010.508777

Frontiers in Immunology | Molecular Innate Immunity

Evolution of immunity

63. Schmid-Hempel P. Immune defence, parasite evasion strategies and their relevance for macroscopic phenomena such as virulence. Philos Trans R Soc Lond
B Biol Sci (2009) 364:85–98. doi:10.1098/rstb.2008.0157
64. Maizels RM. Toxocara canis: molecular basis of immune recognition and evasion. Vet Parasitol (2013) 193:365–74. doi:10.1016/j.vetpar.2012.12.032
65. Buchmann K. Fish immune responses against endoparasitic nematodes – experimental models. J Fish Dis (2012) 35:623–35. doi:10.1111/j.1365-2761.2012.
01385.x
66. Clark SE, Eichelberger KR, Weiser JN. Evasion of killing by human antibody
and complement through multiple variations in the surface oligosaccharide
of Haemophilus influenzae. Mol Microbiol (2013) 88:603–18. doi:10.1111/mmi.
12214
67. Higa N, Toma C, Nohara T, Nakasone N, Takaesu G, Suzuki T. Lose the battle
to win the war: bacterial strategies for evading host inflammasome activation.
Trends Microbiol (2013) 21:342–9. doi:10.1016/j.tim.2013.04.005
68. Buchmann K, Lindenstrøm T. Interactions between monogenean parasites and
their fish hosts. Int J Parasitol (2002) 32:309–19. doi:10.1016/S0020-7519(01)
00332-0

Conflict of Interest Statement: The author declares that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 01 July 2014; paper pending published: 06 August 2014; accepted: 08
September 2014; published online: 23 September 2014.
Citation: Buchmann K (2014) Evolution of innate immunity: clues from invertebrates
via fish to mammals. Front. Immunol. 5:459. doi: 10.3389/fimmu.2014.00459
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Buchmann. This is an open-access article distributed under the terms
of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are
credited and that the original publication in this journal is cited, in accordance with
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does not comply with these terms.

September 2014 | Volume 5 | Article 459 | 30

REVIEW ARTICLE
published: 07 November 2014
doi: 10.3389/fimmu.2014.00544

Evolutionary roots of arginase expression and regulation
Jolanta Maria Dzik *
Department of Biochemistry, Faculty of Agriculture and Biology, Warsaw University of Life Sciences – SGGW, Warszawa, Poland

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Andrew Tasman Hutchinson,
University of Technology Sydney,
Australia
Dalil Hannani, APCure SAS, France
*Correspondence:
Jolanta Maria Dzik , Department of
Biochemistry, Faculty of Agriculture
and Biology, Warsaw University of
Life Sciences – SGGW, 159
Nowoursynowska, Warszawa 02-776,
Poland
e-mail: jolanta_dzik@sggw.pl

Two main types of macrophage functions are known: classical (M1), producing nitric oxide,
NO, and M2, in which arginase activity is primarily expressed. Ornithine, the product of
arginase, is a substrate for synthesis of polyamines and collagen, important for growth and
ontogeny of animals. M2 macrophages, expressing high level of mitochondrial arginase,
have been implicated in promoting cell division and deposition of collagen during ontogeny
and wound repair. Arginase expression is the default mode of tissue macrophages, but
can also be amplified by signals, such as IL-4/13 or transforming growth factor-β (TGF-β)
that accelerates wound healing and tissue repair. In worms, the induction of collagen gene
is coupled with induction of immune response genes, both depending on the same TGFβ-like pathway. This suggests that the main function of M2 “heal” type macrophages is
originally connected with the TGF-β superfamily of proteins, which are involved in regulation of tissue and organ differentiation in embryogenesis. Excretory–secretory products
of metazoan parasites are able to induce M2-type of macrophage responses promoting
wound healing without participation of Th2 cytokines IL-4/IL-13. The expression of arginase
in lower animals can be induced by the presence of parasite antigens and TGF-β signals
leading to collagen synthesis. This also means that the main proteins, which, in primitive
metazoans, are involved in regulation of tissue and organ differentiation in embryogenesis are produced by innate immunity. The signaling function of NO is known already from
the sponge stage of animal evolution. The cytotoxic role of NO molecule appeared later,
as documented in immunity of marine mollusks and some insects. This implies that the
M2-wound healing promoting function predates the defensive role of NO, a characteristic
of M1 macrophages. Understanding when and how the M1 and M2 activities came to be
in animals is useful for understanding how macrophage immunity, and immune responses
operate.
Keywords: arginase, evolution, hemocytes, M1/M2 macrophages, nitric oxide synthase, parasites, TGF-β, wound
healing

Vertebrate macrophages play an innate defense role against various
pathogens. They perform phagocytosis, bacterial killing, defend
against protozoan and metazoan parasites and take part in wound
healing. To fulfill such protective functions,“resting” macrophages
must be activated. Two main types of macrophage functions have
been identified: classical (M1), producing nitric oxide (NO), and
M2-type, in which arginase (producing the healing molecule,
ornithine) is expressed (1, 2). These responses from macrophages
demand different cascades of biochemical reactions, which are regulated by specific sets of cytokines. M1 type can be stimulated by
pathogen associated molecular patterns (PAMP) or amplified by
T cell cytokines, such as IFN-γ. In contrast, M2 activity is the resident tissue type, and can be amplified by molecules such as IL-4,
IL-13, and transforming growth factor-β (TGF-β). Local signals
polarize macrophages to an appropriate response. These immune
functions are indispensible for both life of an individual and lasting of a species. It is apparent that macrophages, as well as other
cells of innate immune response acting in vertebrates, have their
deep evolutionary roots in cells serving similar function in ancestral invertebrates. Various names have been used to define such
cells in different invertebrate groups, i.e., hemocyte, coelomocyte,
amebocyte, or plasmatocyte, collectively named immunocytes (3).

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However, regardless of the terminology, they perform the same
immune functions, and are of similar morphology.
Macrophages, the professional phagocytic cells in humans,
derive from circulating monocytes or by self renewal in the tissues,
and acquire new morphological and physiological characteristics
according to the organs and microenvironments, in which they settle. However, this unitarian origin is uncertain for circulating and
tissue phagocytes (immunocytes) in invertebrates (3). Immunocytes and macrophages share ability to be activated by non-self
material and to react through the release a variety of biologically
active molecules such as cytokines, NO, reactive oxygen species,
hydrolytic enzymes, and neuroendocrine mediators. In vertebrate
immunity, various organs and immune cells are involved, while
all the molecules determining invertebrate immune response are
harbored in the immunocytes. From the perspective of this review,
the multifunctional role of invertebrate cells seems instructive in
respect to its inheritance by vertebrates.
In search for selection pressure for macrophage differentiation into M1 and M2 phenotypes, it is tempting to look back
to recognize which of the functions of M2 macrophages is
aligned straight with invertebrate immunocytes and is found possibly in the most primitive invertebrates. Accumulated evidences

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strongly suggest that a primary function of M2 macrophages is
tissue repair and wound healing (4, 5). This process demands
polyamines and collagen synthesis what strongly depend on
arginase activity (6).

ANIMAL ARGINASES
Arginase (amidinohydrolase, EC 3.5.3.1) is an ubiquitous enzyme
found in bacteria, yeasts, plants, invertebrates, and vertebrates.
Some bacteria possess a related enzyme, agmatinase (Escherichia
coli or Methanobacterium, also Methanococcus), which belong to
Archaebacteria. Agmatinase produce putrescine and urea, arginase
ornithine and urea. Plant arginases are closer to the agmatinase
clade than to the animal one (7, 8). Agmatinases origin predates
that of arginases and the latter would have appeared in the Bacteria by recruitment of a wide specificity agmatinase and then its
transfer to an eukaryotic cell (according to Sekowska et al. perhaps
through mitochondria) (8).
Most microorganisms and invertebrates studied to date have
only one type of arginase (9). Arginases from plants and ammoniotelic animals are localized in mitochondria (10). In ureotelic
animals, arginase is involved in ammonia detoxication in the
ornithine-urea cycle and is localized in cytosol. The cytosolic
and mitochondrial arginases are isoenzymes named A-I and AII, respectively (11). They are encoded by two separate genes.
The arginase gene duplication is relatively recent, and occurred
after separation of vertebrates and invertebrates (9). It has been
suggested that the mitochondrial A-II is a surviving form of the
ancestral arginase, because the cytosolic A-I is restricted to a subset
of more recently evolved species (11).
The pattern of occurrence of the arginase isoenzymes implies
that the primordial function of the enzyme is regulation of cellular arginine and ornithine metabolism, unrelated to the urea
cycle. Ornithine, the product of arginase-catalyzed reaction, is a
substrate for synthesis of proline and polyamines. It is important
to know that ornithine is formed from glutamate via the pathway leading to arginine synthesis as well. This pathway occurs
in bacteria, plants and animals (10). Both in plants and animals,
polyamines (putrescine, spermidine, and spermine) are involved
in a variety of growth and developmental processes, and they
bind directly to DNA and RNA (12). It has been also shown
that polyamines play a pivotal role in wound healing. Proline (as
hydroxyproline) is indispensable for the collagen biosynthesis in
animals and for the synthesis of cell wall proteins in plants. Participation of ornithine in such vital processes suggests that arginase
could be regulated by factors influencing development and growth
of organisms. TGF-β belongs to a superfamily of ancient proteins, known in all bilaterians, members of which play important
signaling roles in embryogenesis (13).

TGF-β SUPERFAMILY OF PROTEINS
Transforming growth factor-β was originally discovered as a
secreted factor that induced malignant transformation in vitro.
It is a prototype member of a superfamily of secreted, homodimeric polypeptides. These factors affect a variety of biological
processes in both transformed and normal cells, including regulation of embryogenesis, adult cell differentiation, inflammation,
and wound repair (14, 15).

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The TGF-β superfamily may be divided into subfamilies
according to sequence homology. One subfamily consists of the
closely related TGF-β1, -2, and -3. TGF-β2 and -β3 take part in
development signaling, while TGF-β1 signals act in inflammatory responses and tissue necrosis (15). TGF-β1 cDNAs from
different animal species (together with chicken and Xenopus)
show an extremely high degree of conservation (14). Fish TGFβ homologs cluster with their mammalian TGF-β1, -β2, and -β3
counterparts (15).
Bone morphogenetic proteins were initially characterized as
factors that induce bone and cartilage formation (16, 17). Bone
morphogenetic proteins (BMPs) are critical in development,
hematopoiesis, as well as cellular chemotaxis, and cellular differentiation (18, 19). In the BMPs subfamily, BMPs-2 and -4 bear closest
homology to the decapentaplegic complex protein, a Drosophila
protein mediating dorsal/ventral axis specification (14). BMPs-5,
-6, -7, and -8 most closely resemble Drosophila protein 60A, which
is required for the growth of imaginal tissues and for patterning of
the adult wing (20). Genes encoding members of the bone morphogenetic factor (BMP) protein family have been identified in a
sea anemone and an echinoderm (21).
Genes of TGF-β superfamily members cluster in two major
clades: TGF-β sensu stricto/TGF-β related (e.g., Activins, Leftys,
and GDF8s) ligands and BMP related (e.g., BMPs and Nodals)
(22). TGF-β sensu stricto ligands have been identified only
in deuterostomes (Echinodermata, Hemichordata, and Chordata) and are not present in genomic screens of Caenorhabditis,
Drosophila, or Nematostella (23).
Receptors of TGF-β pathway are serine threonine kinases categorized as type I and type II (24, 25). Vertebrates have seven
distinct type I receptors, each of which can mix and match with
one of five type II receptors to mediate signals for the TGF-β family
ligands (26). Ligand binding to the constitutively phosphorylated
type II receptors stimulates recruitment of type I receptors and
formation of a heterodimeric receptor complex. In the complex,
type I receptors are transphosphorylated by type II receptors (13).
A signal from type I receptor to the nucleus is channeled into one
of two intracellular pathways via Smad family of proteins. Three
of the receptors phosphorylate the R-Smads (receptor-regulated
Smads); Smad2 and Smad3 and thereby transduce TGF-β-like
signals, whereas the other four receptors activate the R-Smads;
Smad1, Smad5, and Smad8 to mediate signals characteristic of
those initiated by BMPs (26–28). These R-Smads form multisubunit complexes with a common partner Smads (Co-Smads;
Smad4) before entering the nucleus to affect a response (29). Both
R-Smads and Co-Smads are found in all metazoans (30). I-Smads;
Smad6/7 play inhibitory function, stimulating receptor degradation, or competing with R-Smads in formation of complexes with
Smad4 (29, 31). The regulatory activity of I-Smads evolved after
divergence of the poriferan lineage (32).
Binding of Smads to DNA is not especially specific; they play
a role of comodulators, which act together with transcription factors (pan-metazoan Fos/Jun and Myc), transcription coactivators
(pan-metazoan CBP and CBF-β), and transcription corepressors
(Ski/Sno) to recruit basal transcription machinery (32). The formation of Smad complexes gives a wide range of cooperative interactions, thus enables TGF-β signaling to evoke multiple responses

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ranging from embryonic development to wound repair. ARSmads (activin/TGF-β-specific R-Smads) transactivate various
target genes through interaction with various DNA-binding partners, including plasminogen activator inhibitor-1 (PAI-1), type I
collagen, junB, Smad7, and Mix.2. For inhibition of cell growth by
TGF-β, AR-Smads induce the transcription of cyclin-dependent
kinase (CDK) inhibitors p21 and p15. In addition, Smad3 binds
directly to the promoter region of c-myc and represses the transcription of c-myc. In contrast, only a few target genes for BMPs
have been identified, including Id (inhibitor of differentiation
or inhibitor of DNA-binding) 1/2/3, Smad6, Vent-2, and Tlx-2.
Id proteins, however, play important roles in multiple biological
activities of BMPs (27). Id proteins act as negative regulators of cell
differentiation and positive regulators of cell proliferation (33).
Ligands, receptors of the TGF-β pathway, and Smads are ancient
proteins. They emerged already in the metazoan stem lineage. ISmads, multiple ligand traps, and SARA have been added to the
signaling pathway after the divergence of sponges (32).
Transforming growth factor-β and BMP signaling pathway is
evolutionary conserved, as it was shown for worms, flies, and
vertebrates [Ref. (26)].

TGF-β FUNCTION IN EMBRYOGENESIS AND WOUND
HEALING OF INVERTEBRATES
Caenorhabditis elegans is a free-living nematode, a member of the
lineage, which appeared more than 470 million years ago (34).
There are at least three distinct TGF-β-like pathways in this worm
(35). One of them controls the body size and morphology of the
male tail, but five genes of this pathway (dbl-1, sma-2/-3/-4/-6 )
contribute to resistance against Pseudomonas aeruginosa infection
(36). This pathway controls induction of some genes induced after
Serratia marcescens infection, including lys-8 (lysozyme) (37).
Moreover, dpp, a dbl-1 homolog in Drosophila, is up-regulated
upon immune challenge (38, 39). This pathway shows a clear
homology to the mammalian TGF-β pathway (40), which plays
an important role in immune responses (41). Some targets of this
pathway in Caenorhabditis are known: mab-21, involved in male
ray pattern formation, and lon-1 and lon-3, involved in regulation
of the body size. The latter two encode a cysteine-rich secretory
protein and collagen, respectively. The lon-1 and lon-3 are mainly
expressed in the hypodermis, as they are essential for the body size
regulation (42, 43).
This finding implies that the ancestral pathway of TGF-β signaling in embryogenesis is bound with immune reactions in the
Protostomia, suggesting that TGF-β pathway in immunity has been
conserved generally across their evolution. Induction of collagen
gene in Caenorhabditis during bacterial infection links developmental processes with the tissue repair induced by pathogens.
Collagen is needed for extracellular matrix deposition during both
embryogenesis and wound repair. Hemocytes must migrate to
wounded area for the synthesis of collagen fibrilles. In lesioned
leeches (Annelida), immunocytes are the first cells that are also
involved in closing the wound by using pseudopodia to bridge
the epithelial edges. Subsequently, additional immunocytes complete the obstruction together with granulocytes and NK-like
cells (3). Throughout embryogenesis, hemocytes carry out important developmental functions within the embryo, such as the

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Arginase evolution

engulfment and removal apoptic cells and the laying down of many
extracellular matrix molecules, including collagen IV and laminin,
which compose the basement membrane surrounding internal
organs [Ref. (44)]. Drosophila hemocytes are similar to leukocytes
in respect of activation and migration toward wounds. A requirement for phosphoinositide 3-kinase (PI3K) for the polarization
and active hemotaxis of hemocytes toward an epithelial wound
shows a striking analogy with the mechanism of cell chemotaxis used by Dictyostelium discoideum, mammalian neutrophils
(45), as well as fish macrophages (46). Migration of Drosophila
hemocytes toward wounds depends on CDC42 and Rho signal
transmission (47) and PI3K signaling (45). The migratory pattern
of hemocytes in Drosophila embryos is independent of PI3K signaling, but depends on chemotactic signals from the PDGF/VEGF
ligands (45).
During wound healing in the snails Lymnaea stagnalis (48) and
Limax maximus (49), hemocytes exhibit fibroblast activity while
secreting the extracellular matrix. Similar transformation of a cell
type was observed in fibroblasts transformed into myofibroblasts
at injured sites when acting in wound contraction (50). It points
to the role of hemocytes in invertebrate tissue repair, similarly
to the mammalian wound models, in which the macrophages are
essential cells participating in wound healing (51). PDGF-AB (hetrodimer) and TGF-β1 stimulate chemotaxis of different cell types,
especially hemocytes (49). The increased number of hemocytes
contribute to earlier wound closure at the injured site. The removal
of damaged tissue residues is also accelerated by stimulation of
the phagocytic activity of recruited hemocytes. TGF-β1 regulates
expression of genes of collagen type I and III, and fibronectin
(52, 53). This may mean that the mechanism of wound healing is
conserved from invertebrates to mammals what’s more, arginase
gene expression has been documented in hemocytes (54).
Tissue injury in humans triggers migration of macrophages,
platelets, fibroblasts, myofibroblasts, and eosinophils releasing
TGF-β. TGF-β stimulates fibroblasts and other reparative cells
to proliferate and synthesize extracellular matrix components
(elastin fibers, collagen fibrils, protein–polysaccharides, and glycoproteins). This leads to a provisional repair, followed by fibrosis
and ultimately scarring. Fibrosis of many organs (liver, heart, kidney, pancreas, and skin) is mediated by TGF-β [Ref. (55)]. In experimental murine Schistosoma mansoni infection, gene expression
of type I and type III interstitial collagens, basement membrane
collagen, and TGF-β1 show increased levels of expression after
primary infection (56). Transcription of type I procollagen chains
proα1 and proα2 is TGF-β-regulated through two different pathways during tissue fibrosis. Expression of proα1 depends on the
TGF-β activator protein and expression of proα2 depends on Smad
signaling of TGF-β pathway. In addition, there are other cellular
factors and DNA-binding elements required for the transcription
of these type I procollagen genes. New synthesized procollagen
molecules are processed by enzymes outside the cell.
In the evolution of nematodes, they changed their original freeliving habitus to commensal one and finally to parasitize tissues of
animals (57). The evolution of plathelminths was different. They
changed ectoparasitic mode of life of monogenean trematodes to
endoparasitic one of digeneans and tapeworms. Invertebrates that
are parasitized by these worms, heal damages to the tissue due

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to activation of polyamine and collagen synthesis. Thus arginase
induction required for wound healing in animals without acquired
immunity could be based on the TGF-β signaling pathway.
Aside being a substrate for proline synthesis pathway, ornithine
is a substrate for polyamine production in all eukaryotic cells. As
polyamines are required for high rates of protein synthesis and
cell proliferation (58), they play a pivotal role in repair processes.
Stimulation of putrescine synthesis was observed during regeneration of earthworms and planarians (59). Regenerating tissues
produce spermine, and injured or dying cells release spermine
into the extracellular milieu, so that tissue levels of this compound
increase significantly at inflammatory sites of infection or injury
(60). In snails resistant to the Schistosoma mansoni infection (61),
increased gene expression of ornithine decarboxylase in hemocytes
points to the enhancement of arginase activity, which results in
ornithine production (62, 63). Ornithine decarboxylase produces
putrescine used for the synthesis of other polyamines involved
in DNA protection during cell proliferation. Polyamines assist in
wound healing following miracidial penetration.
Mollusks can be infected with viruses, bacteria, fungi, protists,
digenean trematodes, polychetes, and copepods (64). The infective
stage of the protozoan (haplosporidian) Haplosporidium nelsoni
invades the bivalve tissues through gills and palps spreading then
through the body. Infection of the bivalve Crassostrea virginica
with this protist leads to an increase in the number of circulating
hemocytes and their infiltration into tissues (65). It is suggested
that these cells are involved in limiting parasite damage by plugging
lesions, removing debris, and repairing damaged tissue.
In hemocytes of the snail Biophalaria glabrata infected with the
digenean trematode Schistosoma mansoni, for which a definitive
host is human, the expression of TGF-β receptor gene was slightly
lowered in comparison with those of resistant strains. Early gene
expression was measured only 2 h post exposure to miracidia (61).
Transforming growth factor-β signaling is essential for extracellular matrix development in cold-blooded animals (66). As a
result of infection of salmonid fish with the ectoparasitic caligid
crustacean Lepeophtheirus salmonis insufficient expression of several regulatory proteins, among them TGF-β, brought up delayed
expression of collagen 2a and delayed wound healing. Arginase
gene expression was markedly increased in intact skin of infected
fish (67). Support for the hypothesis that arginase expression
is related to collagen expression comes from observations that
arginase trancripts are down-regulated in concert with collagen
a in resistant oysters five days after challenge with the gramnegative bacterium Roseovarius crassostreae (68). This extracellular
pathogen colonizes the oyster’s inner shell surface and causes
lesions in the epithelial mantle.
Efficient wound healing in invertebrates based on induction
of genes for arginase and collagen biosynthesis (68) mediated by
TGFβ (49, 65), but without cytokines of Th2 cluster being involved,
may mean that also in vertebrates such mechanism of healing is
possible. An innate response to injury may occur in absence of
any adaptive response and can be triggered solely by tissue injury
(69). Although IL-4/IL-13 mediated responses may be important
in tissue repair, they do not appear to be essential, as the incision
is effectively healed in the mice that lack IL-4 or IL-4 receptor.
Nonetheless, the importance of type 2 cytokines in damage tissue

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remodeling and fibrosis is well documented (70). Possibly IL-4
and/or IL-13 mediate a more rapid form of tissue repair that it
is necessary just to maintain tissue integrity. According to Allen
and Wynn (71), Th2 immunity in vertebrates evolved as a means
to rapid tissue damage repair caused by metazoan invaders rather
than just to control parasite numbers.

M2-TYPE OF MACROPHAGE RESPONSE WITHOUT HELP OF
Th2 CYTOKINES
Transforming growth factor-β, IL-4, and IL-13 are key cytokines
skewing macrophages to the M2-type response that is typical for
allergy and metazoan parasite infection. Arginase induction is
the hallmark of this response. This raises the question whether
M2-type of macrophage response could be induced solely by
multicellular parasites without help of Th2 cytokines.
A strong wound healing response would occur in helminth
infection, as tissue migratory or tissue invasive parasites often
lead to physical trauma. A Th2-type protective immune response
develops following infection with many tissue-dwelling intestinal
nematode parasites (Heligmosomoides polygyrus, Trichuris muris,
or Trichinella spiralis) or trematodes and is characterized by elevations in IL-4 and IL-13 and increased numbers of CD4+ T cells,
granulocytes, and macrophages. These cells accumulate at the site
of infection and may mediate resistance to worms (72, 73). This
is a kind of defense strategy of the host, but it eventually favors a
survival of parasite.
Excretory-secretory (ES) products or parasite enzymes activate and regulate host-immune response at the macrophage level
through inhibition of pro-inflammatory cytokines production
and induction of macrophages toward the M2-type of activation.
Trichinella spiralis is the parasitic nematode of higher vertebrates, which causes pathological changes in various tissues of the
host. Binding of TGF-β with specific antibodies abrogated effect of
infection on arginase activity in guinea pig alveolar macrophages
(74). ES products from Trichinella spiralis raise the expression of
interleukin-10, TGF-β, and arginase-1 in J774 A.1 macrophages
in the absence of Th2 cytokines (75). In addition, ES products
significantly inhibit translocation of NF-κB into the nucleus and
the phosphorylation of both ERK1/2- and p38MAP-kinases in
J774A.1 macrophages stimulated with lipopolysaccharide (LPS)
(an antigen of Gram-negative bacteria). Treatment of peritoneal
macrophages with a recombinant of 53-kDa protein derived from
T. spiralis brought about expression of mannose receptor, a novel
mammalian lectin (Ym1), arginase-1, and IL-10, hallmarks of M2
phenotype. This effect was independent of IL-4Rα, but dependent
on STAT6 (76).
Infections with the trematodes Fasciola hepatica or Schistosoma
mansoni cause destruction of the host liver tissues, damage to bile
ducts, atrophy of the portal vessels, and secondary pathological
conditions. Secreted peroxiredoxins may induce alternative activation of macrophages. They stimulate Ym1 expression in vitro,
which shows their action independent of IL-4/IL-13 signaling (77).
As expected, administration of recombinat peroxiredoxins from
these trematodes to the wild type and IL-4− /− and IL-13− /− mice
induces alternatively activated macrophages. Also eggs of S. mansoni laid in the smallest blood vessels cause tissue reaction in the
form of inflammation, necrosis, connective tissue encapsulation,

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and eventual scar formation during their migration through the
tissue to the colon. The eggs trapped in the liver induce fibrosis and
are associated with production of proline (78). The immunomodulatory pentasaccharide LNFPIII, which contains the Lewis X
trisaccharide, is a component of schistosome soluble egg antigen. It up-regulates expression and activity of arginase-1, as well
as expression of Ym1 in macrophages (79) but does not induce
expression of FIZZ-1, MGL-1, or MMR. Upregulation of arginase
I and Ym1 is independent of IL-4 and IL-13. Binding of LNFPIII
to C-type lectins on the surface of macrophages leads to alternative nuclear factor (NF)-κB activation (80) and may induce
arginase-1 and Ym1 directly without IL-4 and IL-13. An injection of LNFPIII initiates alternative activation, do not mimicking
complete infection because it does not cause FIZZ-1 expression,
besides upregulation of Ym1. Interestingly, Loke et al. (69) have
found that surgical trauma leads to elevation of markers of alternative activation without presence of T cells. However, the innate
expression of Ym1, FIZZ-1, and arginase-1 requires either IL-4
or IL-13. Expression of arginase-1 occurred early in response to
surgery. It increases with growing up to third day post surgery and
then returns to baseline by 1 week, but is sustained only in the
parasite-implanted animals.
Protozoan parasite Toxoplasma type I and type III strains may
induce the M2 phenotype, while the type II strain induces M1
phenotype (81). The alternative activation of macrophages is
dependent in large part on the Toxoplasma polymorphic protein
kinase ROP16, while the classical activation of macrophages by
the type II strain is due to unique ability of its GRA15 protein
to activate NF-κB pathway and elicit pro-inflammatory cytokines.
Both enzymes seem act in a way specific to the host. According
to authors, parasite effectors from different Toxoplasma strains
evolved to work optimally in hosts predisposed to certain types
of immune responses, such as those along the Th1/Th2/Th17 or
M1/M2 axes. Ending up to the wrong host might lead to severe
disease and failure to establish chronic infection.

MACROPHAGE POLARIZATION BY TGF-β SUPERFAMILY
PROTEINS
In the adult Drosophila immune response, Dpp (decapentaplegic),
a BMP-type signal, is rapidly activated by wounds and represses
the production of antimicrobial peptides. The activin/TGF-β-like
signal dawdle (daw), in contrast, is activated by Gram-positive
bacterial infection but repressed by Gram-negative infection or
wounding; its role is to limit infection-induced melanization.
Genes dpp and daw are expressed in hemocytes but also in other
tissues. The hemocyte population in the adult fly is comprised of
subsets of cell that can be defined through distinct gene expression
profile. According to Clark et al. (82), it is likely that expression of
dpp and daw by a subset or subsets of phagocytes indicates distinct
immunomodulatory functions by these cells. Both dpp and daw
inhibit immune responses. This makes the fly similar to mammals, in which both activin and TGF-β-like and BMP-like signals
are largely anti-inflammatory (83, 84), in contrast with the nematode Caenorhabditis, where the TGF-β superfamily member dbl-1
analog of dpp promotes a variety of antimicrobial responses (85).
Both anti- and pro-inflammatory response due to activation of TGF-β superfamily receptors by their ligands, TGF-β,

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Arginase evolution

bone morphogenetic protein-7 (BMP-7), BMP-6 have been found
in macrophages of rodents. Surprisingly few studies have evaluated the effect of TGF-β signaling on macrophages. Mouse
macrophages lacking TβRII (transforming growth factor-β receptor II) are defective in expression of genes that characterize the
M2-type of activation, suggesting that TGF-β signaling is needed
for the alternative activation of macrophages. Lack of T βRII −/−
is associated with basal expression of arginase-1 (protein and
mRNA) significantly decreased in comparison with the wild type
both in naïve peritoneal macrophages and bone marrow-derived
macrophages, BMDM, (86). Moreover, when T βRII −/− BMD
macrophages are polarized toward an M2 phenotype with IL-4,
induction of Arg-1 is very low. Expression of Arg-1 is increased
in WT macrophages stimulated with TGF-β1. As transcription of
other M2 markers including ym1 ceases in T βRII −/− BMDMs,
apparently signals through TβRII modulate the M2 transcription
program. TGF-β contributes to M2 polarization of macrophages
with IL-4 through co-signaling to Akt, which is one of the TGF-β1
non-Smad-associated signal transduction pathways in other cell
types (87).
Bone morphogenetic protein-7 activates receptor BMPR2
in monocytes, which results in phosphorylation of RSMAD1/5/8/and activation of down-stream mediators in the
Smad pathway. It plays a role in polarization also in M2
macrophages, as manifested by increased expression of antiinflammatory cytokines (88). In bone marrow-derived M2
macrophages, increased polarization results from activation of
PI3K pathway (89). Activation of the PI3K pathway controls production of transcription factors. They regulate key inflammatory
cytokines resulting in increased expression of anti-inflammatory
markers (90). Arginase-1 and IL-10 level is significantly increased
following treatment of monocytes with BMP-7 (88). In addition
to the canonical Smad-dependent pathway for TGF-β signaling,
a Smad-independent pathway, namely the mitogen-activated protein kinase (MAPK) pathway (p38MAPK and JNK) may act (91).
Activation of the NF-κB pathway via the X-linked inhibitor of
apoptosis (XIAP) transduces BMP signaling (92). Different to
BMP-7, BMP-6 may induce pro-inflammatory inducible NOS
(iNOS) and TNF-α in peritoneal macrophages (93). The general
phenotype of macrophages in response to BMP-6 is similar to
that of macrophages exposed to LPS (94). BMP-6 in macrophages
appears to counteract TGF-β. It is likely that the BMP-6 induction of expression of inducible NO synthase occurs through IL-1β
via Smad and NF-kappaB signaling pathways (95). IL-1β, in turn,
up-regulates iNOS expression via the NF-κB pathway. However,
the possibility that BMP-6 may directly activate NF-κB signaling
could not be excluded because TGF-β activates kinase 1 (TAK1),
which is a component of the BMP signaling pathway in Xenopus and mouse embryonic development (96, 97). TAK1 and its
regulators (TAB1 and TAB2) form complexes and activate the
IKK complex (98). The latter possibility suggest that pathway
used in embryonic signaling could be used in innate immunity response to induce NO production. Interestingly, NO and
ornithine (the product of arginase described earlier), both originate from the same amino acid, arginine, via different enzymatic
reactions, which were called figuratively “The arginine fork in the
road” (99).

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EVOLUTION OF NITRIC OXIDE SYNTHASE
In macrophages, NO is a crucial mediator of cytotoxicity. It has
been shown to have microbicidal, antiviral, antiparasitic, and
antitumor effects. NO production is usually mediated by nitric
oxide synthase (NOS) (EC 1.14.13.39). To date, three isoforms
of NOS have been characterized: iNOS, neuronal NOS (nNOS),
and endothelial NOS (eNOS). In macrophages, iNOS is transcriptionally induced in response to LPS, TNF-α, interferon-γ (IFN-γ),
and interleukin-1β (IL-1β) (100). The signaling pathway for iNOS
expression in macrophages involves NF-κB and signal transducer
and activator of transcription (STAT).
Three NOS isoforms originally described in mammalian tissues (100, 101) are encoded by distinct genes: for eNOS, nNOS,
and iNOS. All they share much of their sequence with cytochrome
P450 reductase in their C-terminal reductase domains and have a
common oxygenase domain.
The interdomain linker between the oxygenase and reductase domains contains a calmodulin-binding sequence. In eNOS
and nNOS, physiological concentrations of Ca+2 in cells regulate
the binding of calmodulin to the linker, thereby initiating electron transfer from the flavins to the heme moieties. In contrast,
calmodulin remains tightly bound to the iNOS (Ca+2 -insensitive
isoform). Expression of iNOS is strongly activated in the presence
of LPSs or in response to potentially damaging stimuli, resulting
in a high and long-term NO yield. iNOS is primarily involved in
defense reactions and cytotoxicity.
Nitric oxide synthase ancestry goes back to early bacteria from
before a couple billion years ago. In all prokaryotic enzymes only
the oxygenase domain is found (102, 103). In NOS evolution,
multiple events of gene loss and gain in various lineages occurred.
Nitric oxide synthase occurs probably in almost all invertebrates
ranging from jellyfish (104) and hydra (105) to fly (106) and parasitic worms (107), as well as mollusks and arthropods (108). NOS
enzymes from insects (109–111) and mollusks [(112), Ref. (113)]
have greater overall sequence similarity to a neuronal-like NOS
than to iNOS or eNOS in vertebrates. However, cnidarian (Discosoma) and slime mold (Physarum) NOSs (113, 114) lack the
distinct structural element that is present as an insertion in the
reductase domains of constitutive NOSs but is absent in iNOSs
of vertebrates. This insert is thought to be an autoinhibitory loop,
which impedes binding of Ca2+ to calmodulin and enzymatic activation. This insert reduces potentially toxic NO yields following
the activation of iNOS. Since the Discosoma NOS is structurally
similar both to the only known non-animal conventional NOS
and to vertebrate iNOS isoforms, the inducible type of the enzyme
may be ancestral for animal NOSs. In contrast to vertebrate species,
which have three NOS genes, only one type of NOS isoform has
been found in the genomes from insects and tunicates. Mollusks,
sea urchins, and cephalochordates have at least two NOS genes
but no NOS genes have been identified in Caenorhabditis elegans. These findings imply that more than one NOS co-existed
in the common ancestor of all animals and it was lost in some
animal lineages in the course of evolution. On the other hand, in
some groups, such as mollusks (with at least two different types of
NOS) and chordates (2–3 NOS genes), duplication events for NOS
genes may have occurred more than once. Moreover, duplications
happened independently in the evolution of inducible type NOS,

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Arginase evolution

since some fishes have more than one iNOS-like gene (113). The
diversification of vertebrate NOSs occurred in parallel in many
lineages, which cluster into three distinct groups corresponding
to the mammalian iNOS, eNOS, and nNOS, iNOS probably being
most basal. eNOS apparently originated as the last, within the
mammalian clade.
The primary and evolutionary conservative role of NO is
NO-cGMP signaling, acting in many different invertebrates from
sponges, insects, and mollusks to cephalopods [Ref. (115)]. A
defense function of NO was observed in the crustacean (116),
and mollusk hemocytes [Ref. (117)]. In general, defense functions in invertebrates are accomplished by superoxide produced
by phagocyte NADPH oxidase (which appeared before the divergence of the Choanoflagellata and metazoans), antimicrobial peptides, lysozymes, hemolymph clotting, and melanization [Ref.
(115)]. This suggests that the function of arginase as the key
enzyme producing ornithine in metazoans, indispensible for tissue repair, is more ancient than the cytotoxic activity of free
radical NO, the product of NOS. One may conclude that the
wound healing function of M2 macrophages is more deeply
rooted in history of life than the cytotoxic activity of M1
macrophages.

REGULATION OF NITRIC OXIDE PRODUCTION IN INSECTS
AND MICE
Most of research on the NOS refers to three animal species: fruit
flies Drosophila melanogaster, mice Mus musculus, and humans
Homo sapiens. Presumably, the insects are the least advanced its
evolution. Mice, as rodents, are relatively primitive mammals,
closest relatives of the order Primates, to which humans belong.
From evolutionary point of view, the rodent macrophages, commonly used as a model for immunological investigations, better
suit to studies on innate immunity reactions in vertebrates than
human ones.
The model for studying arthropod immunity is the antimicrobial defense in Drosophila. IMD pathway performs a signaling
function by inducing host defenses in response to Gram-negative
bacteria. Activation the IMD signaling pathway leads to the activation of NF-κB homolog Relish and production of antimicrobial
peptides (118, 119). The cells in Drosophila gut detect the pathogen
and activate hemocytes via an NO-dependent signal. The hemocytes act in turn to activate immune-inducible gene expression in
the fat body (the insect liver analog) by an as-yet-unknown signal
(118). In Anopheles stephensi, expression of immune responsive
genes, including NOS, is up-regulated in response to the presence
of Plasmodium parasites in the midgut (120).
It has been found that A. stephensi NOS possesses a putative LPS- and cytokine-responsive transcription factor binding
site (121). Invertebrates have cytokine-like proteins similar to
the interleukins and tumor necrosis factors of vertebrates (122).
Transcription factor binding sites in the 50 -flanking sequence
demonstrate a bipartite distribution of LPS- and inflammatory
cytokine-responsive elements that are strikingly similar to that
described for murine iNOS gene promoters (123, 124). Studies
of Drosophila NOS regulation have shown (125) that insect NOS
activity is solely dependent on Ca2+ and calmodulin, like the constitutive vertebrate NOSs. Although the activity of Drosophila NOS

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Dzik

is very low compared to other NOSs, low amount of NO produced
may be sufficient for functioning as a signaling molecule (126).
As already commented, NO is rather a signaling than cytotoxic
molecule in invertebrates. In vertebrates, as exemplified by rodent
macrophages, high amounts of NO are produced as a result of
activation of iNOS with LPS and IFNγ. This is a part of defense
armor, which otherwise acts as a double edge sword. However,
stimulation of macrophages only with LPS results in a low NO
production and less than 15% of cells is iNOS positive. IFNγ
enhanced LPS-induced secretion of NO by recruiting increasingly
greater numbers of macrophages into the production of iNOS
(127). The gene of iNOS is synergistically activated by LPS and
IFNγ (123, 124). The iNOS promotor contains two important
regions termed RI and RII. The effects of LPS stimulation are
mediated by elements in both RI and RII, whereas IFNγ functions
through RII only (123, 124). In addition, IFNγ alone is not able
to activate through RII, acting solely to augment the effect of LPS
on RI (123). A variety of LPS response elements, including NF-κB,
occurs and the NF-κB site in LPS-mediated transcriptional activation of iNOS is required (128). Perhaps the increase of frequency
of LPS responsive cells in effect of IFNγ action, and consequent
NO production is the key factor to enhance NO formation. It leads
to M1 type of response.
TGF-β1 seems to be the most potent regulator of iNOS. In
natural killer cells, neutrophils and macrophages, TGF-β1 diminishes iNOS activity, influencing gene expression, mRNA stability
and translation, and NOS protein stability (129). For suppression of LPS-stimulated iNOS in bone marrow-derived mouse
macrophages, both Smad2 and Smad3 are required. Down regulation of iNOS mRNA undergoes by suppressing the IRF3- IFNβSTAT1 pathway (130). Mutual feedback regulation between iNOS
and TGF-β1 is also possible, as latent TGF-β1 can be activated by
exogenous NO (131).TGF-β appears to be important endogenous

Arginase evolution

mediator that keeps resident/wound healing macrophages in M2
dominant mode. A decrease in TGF-β production in macrophages
brought about “activation” of these cells. Similarly, removing of
TGF-β from cell culture (coming from serum added), caused much
more NO produced, and less synergy between LPS and IFN-γ in
stimulation of NO production (1).

CONCLUDING REMARKS
Sophisticated ways of signaling observed in contemporary vertebrates show how complex are results of molecular evolution.
Although specialization of macrophage responses is based on
two ancient mechanisms: cytotoxic activity of iNOS and anabolic function of arginase (Figure 1). It is suggested here that
not defense against infection but rather the TGF-β-signaling was
at the origin of the M1/M2 macrophage specialized functions.
Such signaling is known to operate already in primitive invertebrates, both in their embryonic development and wound healing.
A prototypic inducer of M1 response bacterial LPS, alone activates production of only a small amount of NO by iNOS-type
enzyme and generates signal propagation through cGMP cyclase
in invertebrates. It remains unknown, to what degree invertebrate analogs of IFNγ would be able to enhance LPS-induced
NO production, as no experimental data about enhancement of
NO production by IFNγ-like cytokine in Drosophila are available. Presumably, the ability of M1 macrophages to produce large
amounts of NO in response to microbial infection is a vertebrate evolutionary invention, known to be present already at
the fish grade (132). At this stage the arginase function in M2
macrophages, inherited after invertebrate ancestors, was to deliver
ornithine for processes of extracellular matrix synthesis, of importance in organogenesis and wound healing. The latter serves also
as a protection against metazoan parasites. Thus, the main function of M2 macrophages is originally connected with the TGF-β

FIGURE 1 | Proposed order of appearance of arginase,TGF-β superfamily of proteins, and nitric oxide synthase (NOS), superimposed on the metazoan
phylogenetic tree, implies a pattern of polarization of vertebrate macrophage to M1 and M2-types of activation.

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Dzik

superfamily of proteins, which in primitive metazoans are involved
in regulation of tissue and organ differentiation in embryogenesis.
Looking back in evolution also indicates that both NOS/NO and
arginases/ornithine are primitive innate responses in macrophages
that long preceded the development of T and B cells (adaptive
immunity).

REFERENCES
1. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.4049/
jimmunol.164.12.6166
2. Mosser DM. The many faces of macrophage activation. J Leukoc Biol (2003)
73:209–12. doi:10.1189/jlb.0602325
3. Ottaviani E. Immunocyte: the invertebrate counterpart of the vertebrate
macrophage. Inv Surv J (2011) 8:1–4.
4. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8:958–69. doi:10.1038/nri2448
5. Mylonas KJ, Nair MG, Prieto-Lafuente L, Paape D, Allen JE. Alternatively
activated macrophages elicited by helminth infection can be reprogrammed
to enable microbial killing. J Immunol (2009) 182:3084–94. doi:10.4049/
jimmunol.0803463
6. Hesse M, Modolell M, La Flamme AC, Schito M, Fuentes JM, Cheever
AW, et al. Differential regulation of nitric oxide synthase-2 and arginase1 by type 1/type 2 cytokines in vivo: granulomatous pathology is shaped
by the pattern of L-arginine metabolism. J Immunol (2001) 167:6533–44.
doi:10.4049/jimmunol.167.11.6533
7. Chen H, McCaig BC, Melotto M, He SY, Howe GA. Regulation of plant arginase
by wounding, jasmonate, and the phytotoxin coronatine. J Biol Chem (2004)
279:45998–6007. doi:10.1074/jbc.M407151200
8. Sekowska A, Danchin A, Risler JL. Phylogeny of related functions: the case of
polyamine biosynthetic enzymes. Microbiology (2000) 146:1815–28.
9. Samson ML. Drosophila arginase is produced from a nonvital gene that contains the elav locus within its third intron. J Biol Chem (2000) 275:31107–14.
doi:10.1074/jbc.M001346200
10. Morris SM. Regulation of enzymes of the urea cycle and arginine metabolism.
Annu Rev Nutr (2002) 22:87–105. doi:10.1146/annurev.nutr.22.110801.140547
11. Jenkinson CP, Grody WW, Cederbaum SD. Comparative properties of
arginases. Comp Biochem Physiol (1996) 114B:107–32. doi:10.1016/03050491(95)02138-8
12. Romero-Calderon R, Krantz DE. Transport of polyamines in Drosophila S2
cells: kinetics, pharmacology and dependence on the plasma membrane proton
gradient. Biochem J (2006) 393:583–9. doi:10.1042/BJ20050981
13. Massagué J. TGF-β signal transduction. Annu Rev Biochem (1998) 67:753–91.
doi:10.1146/annurev.biochem.67.1.753
14. Gitelman SE, Derynck R. Transforming growth factor-β. In: Nicola NA, editor.
Guidebook to Cytokines and their Receptors. Oxford: A Sambrook and Tooze
Publication at Oxford University Press (1994). p. 223–6.
15. Magor BG, Magor KE. Evolution of effectors and receptors of innate immunity.
Dev Comp Immunol (2001) 25:651–82. doi:10.1016/S0145-305X(01)00029-5
16. Wang EA, Rosen V, Cordes P, Hewick RM, Kriz MJ, Luxenberg DP, et al. Purification and characterization of other distinct bone-inducing factors. Proc Natl
Acad Sci U S A (1988) 85:9484–8. doi:10.1073/pnas.85.24.9484
17. Wozney JM, Rosen V, Celeste AJ, Mitsock LM, Whitters MJ, Kriz RW, et al.
Novel regulators of bone formation: molecular clones and activities. Science
(1988) 242:1528–34. doi:10.1126/science.3201241
18. Hogan BL. Bone morphogenetic proteins: multifunctional regulators of vertebrate development. Genes Dev (1996) 10:1580–94. doi:10.1101/gad.10.13.1580
19. Zhao GQ. Consequences of knocking out BMP signaling in the mouse. Genesis
(2003) 35:43–56. doi:10.1002/gene.10167
20. Khalsa O, Yoon JW, Torres-Schumann S, Wharton KA. TGF-β/BMP superfamily members, Gbb-60A and Dpp, cooperate to provide pattern information
and establish cell identity in the Drosophila wing. Development (1998) 125:
2723–34.
21. Lelong C, Mathieu M, Favrel P. Identification of new bone morphogenetic
protein-related members in invertebrates. Biochimie (2001) 83:423–6. doi:10.
1016/S0300-9084(01)01260-3

Frontiers in Immunology | Molecular Innate Immunity

Arginase evolution

22. Matus DQ, Magie CR, Pang K, Martindale MQ, Thomsen GH. The Hedgehog
gene family of the cnidarian, Nematostella vectensis, and implications for understanding metazoan Hedgehog pathway evolution. Dev Biol (2008) 313:501–18.
doi:10.1016/j.ydbio.2007.09.032
23. Matus DQ, Thomsen GH, Martin dale MQ. Dorso/ventral genes are
asymmetrically expressed and involved in germ-layer demarcation during
cnidarian gastrulation. Curr Biol (2006) 16:499–505. doi:10.1016/j.cub.2006.
01.052
24. Wrana JL, Attisano L, Wieser R, Ventura F, Massagué J. Mechanism of activation
of the TGF-beta receptor. Nature (1994) 370:341–7. doi:10.1038/370341a0
25. Heldin CH, Miyazono K, ten Dijke P. TGF-β signalling from cell membrane
to nucleus through SMAD proteins. Nature (1997) 390:465–71. doi:10.1038/
37284
26. Attisano L, Wrana JL. Signal transduction by the TGF-β superfamily. Science
(2002) 296:1646–7. doi:10.1126/science.1071809
27. Miyazawa K, Shinozaki M, Hara T, Furuya T, Miyazono K. Two major Smad
pathways in TGF-β superfamily signalling. Genes Cells (2002) 7:1191–204.
doi:10.1046/j.1365-2443.2002.00599.x
28. Qin BY, Chacko BM, Lam SS, de Caestecker MP, Correia JJ, Lin K. Structural
basis of Smad1 activation by receptor kinase phosphorylation. Mol Cell (2001)
8:303–12. doi:10.1016/S1097-2765(01)00417-8
29. Massagué J, Seoane J, Wotton D. Smad transcription factors. Genes Dev (2005)
19:2783–810. doi:10.1101/gad.1350705
30. Srivastava M, Simakov O, Chapman J, Fahey B, Gauthier ME, Mitros T, et al. The
Amphimedon queenslandica genome and the evolution of animal complexity.
Nature (2010) 466:720–6. doi:10.1038/nature09201
31. Hanyu A, Ishidou Y, Ebisawa T, Shimanuki T, Imamura T, Miyazono K. The
N domain of Smad7 is essential for specific inhibition of transforming growth
factor-β signaling. J Cell Biol (2001) 155:1017–27. doi:10.1083/jcb.200106023
32. Richards GS, Degnan BM. The dawn of developmental signaling in the metazoa. Cold Spring Harb Symp Quant Biol (2009) 74:81–90. doi:10.1101/sqb.2009.
74.028
33. Yokota Y, Mori S. Role of Id family proteins in growth control. J Cell Physiol
(2002) 190:21–8. doi:10.1002/jcp.10042
34. Balinski A, Sun Y, Dzik J. Traces of marine nematodes from 470 million
years old early Ordovician rocks in China. Nematology (2013) 15:567–74.
doi:10.1163/15685411-00002702
35. Schulenburg H, Kurz CL, Ewbank JJ. Evolution of the innate immune system:
the worm perspective. Immunol Rev (2004) 198:36–58. doi:10.1111/j.01052896.2004.0125.x
36. Tan MW. Genetic and genomic dissection of host–pathogen interactions
using a P. aeruginosa-C. elegans pathogenesis model. Pediatr Pulmonol (2001)
32:96–7.
37. Mochii M, Yoshida S, Morita K, Kohara Y, Ueno N. Identification of transforming growth factor-β regulated genes in Caenorhabditis elegans by differential
hybridization of arrayed cDNAs. Proc Natl Acad Sci U S A (1999) 96:15020–5.
doi:10.1073/pnas.96.26.15020
38. Irving P, Troxler L, Heuer TS, Belvin M, Kopczynski C, Reichhart JM, et al. A
genome-wide analysis of immune responses in Drosophila. Proc Natl Acad Sci
U S A (2001) 98:15119–24. doi:10.1073/pnas.261573998
39. De Gregorio E, Spellman PT, Rubin GM, Lemaitre B. Genome-wide analysis of
the Drosophila immune response by using oligonucleotide microarrays. Proc
Natl Acad Sci U S A (2001) 98:12590–5. doi:10.1073/pnas.221458698
40. Newfeld SJ, Wisotzkey RG, Kumar S. Molecular evolution of a developmental
pathway: phylogenetic analyses of transforming growth factor-b family ligands,
receptors and Smad signal transducers. Genetics (1999) 152:783–95.
41. Letterio JJ, Roberts AB. Regulation of immune responses by TGF-β. Annu Rev
Immunol (1998) 16:137–61. doi:10.1146/annurev.immunol.16.1.137
42. Morita K, Flemming AJ, Sugihara Y, Mochii M, Suzuki Y, Yoshida S, et al. A
Caenorhabditis elegans TGF-β, DBL-1, controls the expression of LON-1, a
PR-related protein, that regulates polyploidization and body length. EMBO J
(2002) 21:1063–73. doi:10.1093/emboj/21.5.1063
43. Maduzia LL, Gumienny TL, Zimmerman CM, Wang H, Shetgiri P, Krishna
S, et al. lon-1 regulates Caenorhabditis elegans body size downstream of the
dbl-1TGF-β signaling pathway. Dev Biol (2002) 246:418–28. doi:10.1006/dbio.
2002.0662
44. Fessler JH, Fessler LI. Drosophila extracellular matrix. Annu Rev Cell Biol (1989)
5:309–39. doi:10.1146/annurev.cellbio.5.1.309

November 2014 | Volume 5 | Article 544 | 38

Dzik

45. Wood W, Faria C, Jacinto A. Distinct mechanisms regulate hemocyte chemotaxis during development and wound healing in Drosophila melanogaster. J Cell
Biol (2006) 173:405–16. doi:10.1083/jcb.200508161
46. Grabher C, Cliffe A, Miura K, Hayflick J, Pepperkok R, Rørth P, et al. Birth
and life of tissue macrophages and their migration in embryogenesis and
inflammation in medaka. J Leukoc Biol (2007) 81:263–71. doi:10.1189/jlb.
0806526
47. Stramer B, Wood W, Galko MJ, Redd MJ, Jacinto A, Parkhurst SM, et al. Live
imaging of wound inflammation in Drosophila embryos reveals key roles for
small GTPases during in vivo cell migration. J Cell Biol (2005) 168:567–73.
doi:10.1083/jcb.200405120
48. Sminia T, Pietersma K, Scheerboom JE. Histological and ultrastructural observations on wound healing in the freshwater pulmonate Lymnaea stagnalis.
Z Zellforsch Mikrosk Anat (1973) 141:561–73. doi:10.1007/BF00307125
49. Franchini A, Ottaviani E. Repair of molluscan tissue injury: role of PDGF and
TGF-beta1. Tissue Cell (2000) 32:312–21.
50. Rudolph R, Vande Berg J, Ehrlich PH. Wound contraction and scar contracture. In: Cohen IK, Diegelmann RF, Lindblad WJ, editors. Wound Healing;
Biochemical and Clinical Aspects. (Vol. 6), Philadelphia, PA: Saunders (1992).
p. 96–114.
51. Leibovich SJ, Ross R. The role of the macrophage in wound repair. Am J Pathol
(1975) 78:71–100.
52. Pierce GF, Vande Berg J, Rudolph R, Tarpley J, Mustoe TA. Platelet-derived
growth factor-BB and transforming growth factor β-1 selectively modulate
glycosaminoglycans, collagen, and myofibroblasts in excisional wounds. Am
J Pathol (1991) 138:629–46.
53. Quaglino D, Nanney LB, Ditesheim JA, Davidson JM. Transforming growth
factor-β stimulates wound healing and modulates extracellular matrix gene
expression in pig skin. Incisional wound model. J Invest Dermatol (1991)
97:34–42.
54. Pongsomboon S, Tang S, Boonda S, Aoki T, Hirono I, Yasuike M, Tassanakajon A. Differentially expressed genes in Penaeus monodon hemocytes following infection with yellow head virus. BMB Rep (2008) 41:670–7. doi:10.5483/
BMBRep.2008.41.9.670
55. Cutroneo KR. TGF-β-induced fibrosis and SMAD signaling: oligo decoys as
natural therapeutics for inhibition of tissue fibrosis and scarring. Wound Repair
Regen (2007) 15(Suppl 1):S54–60. doi:10.1111/j.1524-475X.2007.00226.x
56. Jacobs W, Kumar-Singh S, Bogers J, Van de Vijver K, Deelder A, Van Marck E.
Transforming growth factor-β, basement membrane components and heparan
sulphate proteoglycans in experimental hepatic schistosomiasis mansoni. Cell
Tissue Res (1998) 292:101–6. doi:10.1007/s004410051039
57. Blaxter ML. Molecular analysis of nematode evolution. In: Kennedy MW,
Harnett W, editors. Parasitic Nematodes: Molecular Biology, Biochemistry and
Immunology. St. Albans: CAB International (2001). p. 1–24.
58. Wu G. Synthesis of citrulline and arginine from proline in enterocytes of postnatal pigs. Am J Physiol Gastrointest Liver Physiol (1997) 272:G1382–90.
59. Hamana K, Hamana H, Shinozawa T. Alterations in polyamine levels of nematode, earthworm, leech and planarian during regeneration, temperature and
osmotic stresses. Comp Biochem Physiol B Biochem Mol Biol (1995) 111:91–7.
doi:10.1016/0305-0491(94)00222-G
60. Zhang M, Wang H, Tracey KJ. Regulation of macrophage activation and inflammation by spermine: a new chapter in an old story. Crit Care Med (2000) 28(4
Suppl):N60–6. doi:10.1097/00003246-200004001-00007
61. Lockyer AE, Emery AM, Kane RA, Walker AJ, Mayer CD, Mitta G, et al. Early
differential gene expression in haemocytes from resistant and susceptible Biomphalaria glabrata strains in response to Schistosoma mansoni. PLoS One (2012)
7(12):e51102. doi:10.1371/journal.pone.0051102
62. Albina JE, Caldwell MD, Henry WL, Mills CD. Regulation of macrophage functions by L-arginine. J Exp Med (1989) 169:1021–9. doi:10.1084/jem.169.3.1021
63. Witte MB, Barbul A. Arginine physiology and its implication for wound healing.
Wound Repair Regen (2003) 11:419–23. doi:10.1046/j.1524-475X.2003.11605.x
64. Kinne O. Diseases of Marine Animals. Hamburg: Biologische Anstalt Helgoland
(1983).
65. Ford SE, Kanaley SA, Littlewood DT. Cellular responses of oysters infected with
Haplosporidium nelsoni: changes in circulating and tissue-infiltrating hemocytes. J Invertebr Pathol (1993) 61:49–57. doi:10.1006/jipa.1993.1009
66. Roberts AB, Heine UI, Flanders KC, Sporn MB. Transforming growth-factorβ – major role in regulation of extracellular-matrix. Ann N Y Acad Sci (1990)
580:225–32. doi:10.1111/j.1749-6632.1990.tb17931.x

www.frontiersin.org

Arginase evolution

67. Skugor S, Glover KA, Nilsen F, Krasnov A. Local and systemic gene expression
responses of Atlantic salmon (Salmo salar) to infection with the salmon louse.
BMC Genomics (2008) 9:498. doi:10.1186/1471-2164-9-498
68. McDowell IC, Nikapitiya C, Aguiar D, Lane CE, Istrail S, et al. Transcriptome
of american oysters, Crassostrea virginica, in response to bacterial challenge:
insights into potential mechanisms of disease resistance. PLoS One (2014)
9(8):e105097. doi:10.1371/journal.pone.0105097
69. Loke P, Gallagher I, Nair MG, Zang X, Brombacher F, Mohrs M, et al. Alternative activation is an innate response to injury that requires CD4+ T cells
to be sustained during chronic infection. J Immunol (2007) 179:3926–36.
doi:10.4049/jimmunol.179.6.3926
70. Wynn TA. Fibrotic disease and the TH1/TH2 paradigm. Nat Rev Immunol
(2004) 4:583–94. doi:10.1038/nri1412
71. Allen JE, Wynn T. Evolution of Th2 immunity: a rapid repair response to tissue destructive pathogens. PLoS Pathog (2011) 7(5):e1002003. doi:10.1371/
journal.ppat.1002003
72. Patel N, Kreider T, Urban JF Jr, Gause WC. Characterisation of effector
mechanisms at the host: parasite interface during the immune response to
tissue-dwelling intestinal nematode parasites. Int J Parasitol (2009) 39:13–21.
doi:10.1016/j.ijpara.2008.08.003
73. Pearce EJ, Vasconcelos JP, Brunet LR, Sabin EA. IL-4 in schistosomiasis. Exp
Parasitol (1996) 84:295–9. doi:10.1006/expr.1996.0116
74. Dzik JM, Gołos B, Jagielska E, Zielinski Z, Wałajtys-Rode E. A non-classical
type of alveolar macrophage response to Trichinella spiralis infection. Parasite
Immunol (2004) 26:197–205. doi:10.1111/j.0141-9838.2004.00700.x
75. Bai X, Wu X, Wang X, Guan Z, Gao F, Yu J. et al. Regulation of cytokine
expression in murine macrophages stimulated by excretory/secretory products from Trichinella spiralis in vitro. Mol Cell Biochem (2012) 360:79–88.
doi:10.1007/s11010-011-1046-4
76. Du L, Wei H, Li L, Shan H, Yu Y, Wang Y, et al. Regulation of recombinant Trichinella spiralis 53-kDa protein (rTsP53) on alternatively activated
macrophages via STAT6 but not IL-4Rα in vitro. Cell Immunol (2014) 288:1–7.
doi:10.1016/j.cellimm.2014.01.010
77. Donnelly S, Stack CM, O’Neill SM, Sayed AA, Williams DL, Dalton JP.
Helminth 2-Cys peroxiredoxin drives Th2 responses through a mechanism
involving alternatively activated macrophages. FASEB J (2008) 22:4022–32.
doi:10.1096/fj.08-106278
78. Tanabe M, Kaneko N, Takeuchi T. Schistosoma mansoni: higher free proline levels in the livers of infected mice. Exp Parasitol (1991) 72:134–44.
doi:10.1016/0014-4894(91)90131-F
79. Atochina O, Da’dara AA, Walker M, Harn DA. The immunomodulatory glycan LNFPIII initiates alternative activation of murine macrophages in vivo.
Immunology (2008) 125:111–21. doi:10.1111/j.1365-2567.2008.02826.x
80. Thomas PG, Carter MR, Da’dara AA, DeSimone TM, Harn DA. A helminth
glycan induces APC maturation via alternative NF-κB activation independent
of I κBα degradation. J Immunol (2005) 175:2082–90. doi:10.4049/jimmunol.
175.4.2082
81. Jensen KD, Wang Y, Wojno ED, Shastri AJ, Hu K, Cornel L, et al. Toxoplasma polymorphic effectors determine macrophage polarization and intestinal inflammation. Cell Host Microbe (2011) 9:472–83. doi:10.1016/j.chom.
2011.04.015
82. Clark RI, Woodcock KJ, Geissmann F, Trouillet C, Dionne MS. Multiple TGFβ superfamily signals modulate the adult Drosophila immune response. Curr
Biol (2011) 21:1672–7. doi:10.1016/j.cub.2011.08.048
83. Bootcov MR, Bauskin AR, Valenzuela SM, Moore AG, Bansal M, He XY.
MIC-1, a novel macrophage inhibitory cytokine, is a divergent member
of the TGF-β superfamily. Proc Natl Acad Sci U S A (1997) 94:11514–9.
doi:10.1073/pnas.94.21.11514
84. Shull MM, Ormsby I, Kier AB, Pawlowski S, Diebold RJ, Yin M, et al. Targeted
disruption of the mouse transforming growth factor-β1 gene results in multifocal inflammatory disease. Nature (1992) 359:693–9. doi:10.1038/359693a0
85. Mallo GV, Kurz CL, Couillault C, Pujol N, Granjeaud S, Kohara Y, et al.
Inducible antibacterial defense system in C. elegans. Curr Biol (2002)
12:1209–14. doi:10.1016/S0960-9822(02)00928-4
86. Gong D, Shi W, Yi SJ, Chen H, Groffen J, Heisterkamp N. TGF-β signaling plays
a critical role in promoting alternative macrophage activation. BMC Immunol
(2012) 13:31. doi:10.1186/1471-2172-13-31
87. Mu Y, Gudey SK, Landstrom M. Non-Smad signaling pathways. Cell Tissue Res
(2012) 347:11–20. doi:10.1007/s00441-011-1201-y

November 2014 | Volume 5 | Article 544 | 39

Dzik

88. Rocher C, Singla DK. SMAD-PI3K-Akt-mTOR pathway mediates BMP7 polarization of monocytes into M2 macrophages. PLoS One (2013)
8(12):e84009. doi:10.1371/journal.pone.0084009
89. Rauh MJ, Ho V, Pereira C, Sham A, Sly LM, Lam V, et al. SHIP represses the generation of alternatively activated macrophages. Immunity (2005) 23:361–74.
doi:10.1016/j.immuni.2005.09.003
90. Weichhart T, Säemann MD. The PI3K/Akt/mTOR pathway in innate immune
cells: emerging therapeutic applications. Ann Rheum Dis (2008) 67(Suppl
3):iii70–4. doi:10.1136/ard.2008.098459
91. Miyazono K, Maeda S, Imamura T. BMP receptor signaling: transcriptional
targets, regulation of signals, and signaling cross-talk. Cytokine Growth Factor
Rev (2005) 16:251–63. doi:10.1016/j.cytogfr.2005.01.009
92. Lu M, Lin SC, Huang Y, Kang YJ, Rich R, Lo YC, et al. XIAP induces NF-κB activation via the BIR1/TAB1 interaction and BIR1 dimerization. Mol Cell (2007)
26:689–702. doi:10.1016/j.molcel.2007.05.006
93. Hong JH, Lee GT, Lee JH, Kwon SJ, Park SH, Kim SJ, et al. Effect of
bone morphogenetic protein-6 on macrophages. Immunology (2009) 128(1
Suppl):e442–50. doi:10.1111/j.1365-2567.2008.02998.x
94. Fujihara M, Muroi M, Tanamoto K, Suzuki T, Azuma H, Ikeda H. Molecular mechanisms of macrophage activation and deactivation by lipopolysaccharide: roles of the receptor complex. Pharmacol Ther (2003) 100:171–94.
doi:10.1016/j.pharmthera.2003.08.003
95. Kwon SJ, Lee GT, Lee JH, Kim WJ, Kim IY. Bone morphogenetic protein6 induces the expression of inducible nitric oxide synthase in macrophages.
Immunology (2009) 128(1 Suppl):e758–65. doi:10.1111/j.1365-2567.2009.
03079.x
96. Shibuya H, Iwata H, Masuyama N, Gotoh Y, Yamaguchi K, Irie K, et al. Role
of TAK1 and TAB1 in BMP signaling in early Xenopus development. EMBO J
(1998) 17:1019–28. doi:10.1093/emboj/17.4.1019
97. Jadrich JL, O’Connor MB, Coucouvanis E. Expression of TAK1, a mediator of
TGF-β and BMP signaling, during mouse embryonic development. Gene Expr
Patterns (2003) 3:131–4. doi:10.1016/S1567-133X(03)00012-7
98. Wang C, Deng L, Hong M, Akkaraju GR, Inoue J, Chen ZJ. TAK1 is a
ubiquitin-dependent kinase of MKK and IKK. Nature (2001) 412:346–51.
doi:10.1038/35083684
99. Mills CD. M1 and M2 Macrophages: Oracles of Health and Disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
100. Kleinert H, Schwarz PM, Forstermann U. Regulation of the expression of
inducible nitric oxide synthase. Biol Chem (2003) 384:1343–64. doi:10.1515/
BC.2003.152
101. Knowles RG, Moncada S. Nitric oxide synthases in mammals. Biochem J (1994)
298:249–58.
102. Zemojtel T, Wade RC, Dandekar T. In search of the prototype of
nitric oxide synthase. FEBS Lett (2003) 554:1–5. doi:10.1016/S0014-5793(03)
01081-0
103. Adak S, Aulak KS, Stuehr DJ. Direct evidence for nitric oxide production by
a nitric-oxide synthase-like protein from Bacillus subtilis. J Biol Chem (2002)
277:16167–71. doi:10.1074/jbc.M201136200
104. Moroz LL, Meech RW, Sweedler JV, Mackie GO. Nitric oxide regulates swimming in the jellyfish Aglantha digitale. J Comp Neurol (2004) 471:26–36.
doi:10.1002/cne.20023
105. Cristino L, Guglielmotti V, Cotugno A, Musio C, Santillo S. Nitric
oxide signaling pathways at neural level in invertebrates: functional implications in cnidarians. Brain Res (2008) 1225:17–25. doi:10.1016/j.brainres.2008.
04.056
106. Trimmer BA, Aprille J, Modica-Napolitano J. Nitric oxide signalling: insect
brains and photocytes. Biochem Soc Symp (2004) 71:65–83.
107. Kohn AB, Lea JM, Moroz LL, Greenberg RM. Schistosoma mansoni:
use of a fluorescent indicator to detect nitric oxide and related species in
living parasites. Exp Parasitol (2006) 113:130–3. doi:10.1016/j.exppara.2005.
12.013
108. Elofsson R, Carlberg M, Moroz L, Nezlin L, Sakharov D. Is nitric oxide
(NO) produced by invertebrate neurones? Neuroreport (1993) 4:279–82.
doi:10.1097/00001756-199303000-00013
109. Yuda M, Hirai M, Miura K, Matsumura H, Ando K, Chinzei Y. cDNA cloning,
expression and characterization of nitric-oxide synthase from the salivary
glands of the blood-sucking insect Rhodnius prolixus. Eur J Biochem (1996)
242:807–12. doi:10.1111/j.1432-1033.1996.0807r.x

Frontiers in Immunology | Molecular Innate Immunity

Arginase evolution

110. Nighorn A, Gibson NJ, Rivers DM, Hildebrand JG, Morton DB. The nitric
oxide-cGMP pathway may mediate communication between sensory afferents
and projection neurons in the antennal lobe of Manduca sexta. J Neurosci
(1998) 18:7244–55.
111. Regulski M, Tully T. Molecular and biochemical characterization of dNOS: a
Drosophila Ca2+ /calmodulin-dependent nitric oxide synthase. Proc Natl Acad
Sci U S A (1995) 92:9072–6. doi:10.1073/pnas.92.20.9072
112. Korneev SA, Piper MR, Picot J, Phillips R, Korneeva EI, O’Shea M. Molecular characterization of NOS in a mollusc: expression in a giant modulatory
neuron. J Neurobiol (1998) 35:65–76. doi:10.1002/(SICI)1097-4695(199804)
35:1<65::AID-NEU6>3.0.CO;2-9
113. Moroz LL, Kohn AB. Parallel evolution of nitric oxide signaling: diversity of
synthesis & memory pathways. Front Biosci (2014) 16:2008–51. doi:10.2741/
3837
114. Golderer G, Werner ER, Leitner S, Grobner P, Werner-Felmayer G. Nitric
oxide synthase is induced in sporulation of Physarum polycephalum. Genes
Dev (2001) 15:1299–309. doi:10.1101/gad.890501
115. Dzik JM. The ancestry and cumulative evolution of immune reactions. Acta
Biochim Pol (2010) 57:443–66.
116. Yeh FC, Wu SH, Lai CY, Lee CY. Demonstration of nitric oxide synthase activity in crustacean hemocytes and anti-microbial activity of hemocyte-derived
nitric oxide. Comp Biochem Physiol B Biochem Mol Biol (2006) 144:11–7.
doi:10.1016/j.cbpb.2006.01.007
117. Palumbo A. Nitric oxide in marine invertebrates: a comparative perspective.
Comp Biochem Physiol A Mol Integr Physiol (2005) 142:241–8. doi:10.1016/j.
cbpb.2005.05.043
118. Nappi AJ, Vass E, Frey F, Carton Y. Nitric oxide involvement in Drosophila
immunity. Nitric Oxide (2000) 4:423–30. doi:10.1006/niox.2000.0294
119. Foley E, O’Farrell PH. Nitric oxide contributes to induction of innate
immune responses to gram-negative bacteria in Drosophila. Genes Dev (2003)
17:115–25. doi:10.1101/gad.1018503
120. Dimopoulos G, Seeley D, Wolf A, Kafatos FC. Malaria infection of the mosquito Anopheles gambiae activates immune-responsive genes during critical transition stages of the parasite life cycle. EMBO J (1998) 17:6115–23.
doi:10.1093/emboj/17.21.6115
121. Luckhart S, Rosenberg R. Gene structure and polymorphism of an invertebrate nitric oxide synthase gene. Gene (1999) 232:25–34. doi:10.1016/S03781119(99)00121-3
122. Beschin A, Bilej M, Torreele E, De Baetselier P. On the existence of cytokines
in invertebrates. Cell Mol Life Sci (2001) 58(5–6):801–14. doi:10.1007/
PL00000901
123. Lowenstein CJ, Alley EW, Raval P, Snowman AM, Snyder SH, Russell SW, et al.
Macrophage nitric oxide synthase gene: two upstream regions mediate induction by interferon gamma and lipopolysaccharide. Proc Natl Acad Sci U S A
(1993) 90:9730–4. doi:10.1073/pnas.90.20.9730
124. Xie QW, Whisnant R, Nathan C. Promoter of the mouse gene encoding
calcium-independent nitric oxide synthase confers inducibility by interferon
gamma and bacterial lipopolysaccharide. J Exp Med (1993) 177:1779–84.
doi:10.1084/jem.177.6.1779
125. Sengupta R, Sahoo R, Mukherjee S, Regulski M, Tully T, Stuehr DJ, et al.
Characterization of Drosophila nitric oxide synthase: a biochemical study.
Biochem Biophys Res Commun (2003) 306:590–7. doi:10.1016/S0006-291X(03)
01003-9
126. Ghosh DK, Wu C, Pitters ER, Mayer B, Stuehr DJ. Characterization of the
inducible nitric oxide synthase oxygenase domain identifies a 49-amino acid
segment required for subunit dimerization and tetrahydrobiopterin interaction. Biochemistry (1997) 36:10609–19. doi:10.1021/bi9702290
127. Pace JL, Lowenstein CJ, Phillips TA, Chen LC, Morrison DC, Hunt JS, et al.
Population dynamics of inducible nitric oxide synthase production by LPSand LPS/IFN-gamma-stimulated mouse macrophages. J Endotoxin Res (1994)
1:227–33.
128. Xie QW, Kashiwabara Y, Nathan C. Role of transcription factor NF-κ B/Rel in
induction of nitric oxide synthase. J Biol Chem (1994) 269:4705–8.
129. Vodovotz Y. Control of nitric oxide production by transforming growth factorbeta1: mechanistic insights and potential relevance to human disease. Nitric
Oxide (1997) 1:3–17. doi:10.1006/niox.1996.0105
130. Sugiyama Y, Kakoi K, Kimura A, Takada I, Kashiwagi I, Wakabayashi Y. Smad2
and Smad3 are redundantly essential for the suppression of iNOS synthesis

November 2014 | Volume 5 | Article 544 | 40

Dzik

in macrophages by regulating IRF3 and STAT1 pathways. Int Immunol (2012)
24:253–65. doi:10.1093/intimm/dxr126
131. Vodovotz Y, Chesler L, Chong H, Kim SJ, Simpson JT, De Graff W, et al. Regulation of transforming growth factor beta1 by nitric oxide. Cancer Res (1999)
59:2142–9.
132. Forlenza M, Fink IR, Raes G, Wiegertjes GF. Heterogeneity of macrophage activation in fish. Dev Comp Immunol (2011) 35:1246–55. doi:10.1016/j.dci.2011.
03.008
Conflict of Interest Statement: The author declares that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.

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Arginase evolution

Received: 21 August 2014; paper pending published: 23 September 2014; accepted: 13
October 2014; published online: 07 November 2014.
Citation: Dzik JM (2014) Evolutionary roots of arginase expression and regulation.
Front. Immunol. 5:544. doi: 10.3389/fimmu.2014.00544
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Dzik. This is an open-access article distributed under the terms of the
Creative Commons Attribution License (CC BY). The use, distribution or reproduction
in other forums is permitted, provided the original author(s) or licensor are credited
and that the original publication in this journal is cited, in accordance with accepted
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comply with these terms.

November 2014 | Volume 5 | Article 544 | 41

OPINION ARTICLE
published: 04 December 2014
doi: 10.3389/fimmu.2014.00620

Macrophage: SHIP of Immunity
Charles D. Mills 1 *, Anita C. Thomas 2 , Laurel L. Lenz 3 and Markus Munder 4
1

BioMedical Consultants, Marine on St. Croix, MN, USA
Bristol Heart Institute and Bristol CardioVascular, University of Bristol, Bristol, UK
3
Immunology and Microbiology Department, University of Colorado School of Medicine, Aurora, CO, USA
4
Third Department of Medicine, University Medical Center, Johannes Gutenberg University, Mainz, Germany
*Correspondence: mills002@umn.edu
2

Edited by:
Claudia Kemper, King’s College London, UK
Reviewed by:
Geanncarlo Lugo-Villarino, Centre National de la Recherche Scientifique, France
Keywords: macrophage, M1, M2, nitric oxide, iNOS, arginase, wound, cancer

Immunology. Why does it exist? Two
words. Cure disease. People get diseases.
“Test tubes” do not. People fund immunologists for solutions to their health
problems. But, immunologists often study
leukocytes in test tubes – the laboratory – away from diseases. Why? Because
much can be learned from analyzing cellular biochemistry and behaviors in vitro
that cannot be ascertained when leukocytes are in animals. At the same time,
isolated leukocyte reactions often do not
reflect how the immune system operates as a unit. So, it is critical to verify
in vitro observations in vivo. Among leukocytes, macrophages are the central initiating and directing element in immune
systems, and serve this role through four
basic “SHIP” functions in vivo: Sample;
Heal; Inhibit; and Present (antigen) (1–4).
The polar-opposite functions of Heal (M2type) and Inhibit (M1-type) can have profoundly different effects on host survival,
and require unique and major changes
in macrophage metabolism and physiology. In turn, macrophage populations are
necessarily heterogeneous as they adapt
to protect hosts in different ways: they
exhibit “plasticity.” Some have focused on
measuring ever-expanding lists of cell surface or various other “markers” (mostly
in vitro) to try and sub-type macrophages.
But, the “heterogeneity” created by such
studies can be “illusory” because there are
many more markers than there are functions (e.g., M1/inhibit and M2/heal). Thus,
it is important to focus on classifying
macrophages by functions, such as SHIP,
to navigate through a “sea of plasticity.”
And, thereby realize the enormous potential of macrophages/innate immunity for
improving health.

BASIC MACROPHAGE FUNCTIONS
IN VIVO
The earliest in vivo SHIP function observed
in macrophages was their ability to “sample” by ingesting items in their surroundings (5, 6). Through sampling,
macrophages routinely receive “self ” signals that instruct them to repair or replace
lost or effete cells and intercellular matrices.
The heal-type function of macrophages is
now called M2 [(7), reviewed in Ref. (2)].
Following infection (or trauma), M2/healtype macrophages can rapidly switch to
become M1/inhibit-type, to promote host
defense (1). M2/heal responses are mediated by ornithine, and other growthpromoting molecules (8, 9). M1/inhibit
is mediated by nitric oxide (NO) and
other molecules that promote cellular
killing activity (10, 11). Fascinatingly, both
ornithine and NO arise from one amino
acid: arginine (12).
The biochemical basis for the M2/heal
function of macrophages was discovered
before the M1/inhibit function (8, 12). As
illustrated in Figure 1A (top), in sterile
wounds, macrophages produce ornithine
(a precursor of polyamines and collagen
for repair) as healing proceeds (Green – M2
dominance) (13). Around the same time, it
was observed that macrophages in growing tumors were also the M2/ornithineproducing type (Figure 1A middle). This
latter finding provided a biological explanation for the association of intratumor
macrophages with tumor growth (14).
M2/heal-type macrophages have since
been shown to also dominate in human
tumors, and are associated with poor survival (15–18).
The biochemical basis of how M1/
inhibit-type macrophages kill pathogens

Frontiers in Immunology | Molecular Innate Immunity

(or, abnormal “self ”) also came from the
study of wounds and cancer described
above, as well as other studies [(19–
26), reviewed in Ref. (3, 12)]. As mentioned, macrophages have a unique ability to switch from making the heal molecule, ornithine, in vivo to making the
killer molecule, NO (1). Such a switch is
shown in Figure 1A (top). For a brief
period following wounding, a high concentration of NO is present (M1 activity, Red), which can protect the wound
if infectious agents have been introduced
(13). It is not clear exactly what stimuli
cause this injury-induced NO production,
though neutrophils are also involved (27,
28). If the wound is sterile, macrophage
NO production stops, neutrophil emigration ends, and wound macrophages make
ornithine (M2 activity) as mentioned.
Another example of the key protective
function of macrophages making NO is
shown in Figure 1A (bottom). It can be
seen that if a host is vaccinated against
the tumor shown in Figure 1A (middle),
implantation of the tumor causes intratumor macrophages to make a large quantity
of NO that helps cause tumor rejection
(12). Macrophage NO is also an important
defense against a variety of infectious diseases (29). That M2/ornithine or M1/NO
are important effector molecules are supported by studies showing that interference with these activities in vivo alters
healing or host protection [reviewed in
Ref. (12)]. Conversely, overexpression
of M1/inhibit responses is associated
with conditions such as atherosclerosis and arthritis, while M2-type contribute to chronic infections, promotion of tumor growth, and allergies (3,
29). Together, these results demonstrated

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Mills et al.

Macrophage SHIP of Immunity

STERILE WOUND
A

Resident Macrophages M2/M1

From: Albina et al
1990
J. Immunol.
144:3877

GROWING TUMOR

From: Mills et al
1992
J. Immunol.
149:2709

TUMOR REJECTION

0 1-2

4

8

12

Days

M2

B
Take one
Cell

Heal

Resident Macrophages

Hours

Add
LPS or
IFNARG
TGFTNF
IL-6

IL-10

ARG
Changing
NOS
Mixtures
of Cytokines TNF
and Products IL-6
IL-10

0

BIOCHEMICAL AND PHYSIOLOGIC
HOST ELEMENTS THAT INFLUENCE
HOW MACROPHAGES PROTECT HOSTS

4

Macrophage SHIP functions (sample, heal,
inhibit, and present) are regulated by integration of a variety of endogenous (e.g.,
host-derived and resident microbiota) and
exogenous signals (1, 43). For example, in
the absence of infection or injury, TGF-β
helps maintain macrophages in the routine M2/heal mode (7). Other host-derived
molecules, such as oxidized LDL, can stimulate M1-type responses that contribute to
atherosclerosis (4, 44). Following infection
or injury, certain PAMPs and DAMPs stimulate macrophages to switch from M2/heal
to M1/inhibit mode (35–37). IFN-γ was
shown to be the primary T cell product
that further amplifies M1/inhibit activity
(45). Later, macrophage IL-12 was found
to be a key cytokine (along with increased
Class II MHC expression) that stimulates
IFN-γ production by T cells (41, 42, 46).
Macrophages have also been reported to
secrete IFN-γ upon stimulation via IL-12
and IL-18 (47) or via CD40 (48), which
might further enhance M1 polarization
through auto- or paracrine activity. Not
all pathogens stimulate macrophages to
switch from M2/heal to M1/inhibit, and
some seem to suppress such a switch. In
this circumstance, M2-type macrophages
can stimulate T cells to make very different cytokines (such as IL-4, IL-13, and

NOS
IL-12
TNF
IL-6
IL-10

16
Ac!vated Macrophages

M1
Inhibit

FIGURE 1 | (A). Predominance of M1 or M2-dominant macrophage responses in vivo in different types
of inflammation. Top: a wound results in a strong, but brief, shift from local macrophages making
primarily ornithine to making NO. Wound macrophages then become M2-dominant to aid in wound
healing. Middle: a growing tumor elicits macrophages, and macrophage production of NO is
suppressed and ornithine production predominates. Bottom: if a host is vaccinated against a tumor,
implantation of the tumor results in T cell-dependent M1 dominance and the tumor is rejected. (B) M2
to M1-type macrophage conversion and the illusion of “subsets.” Top: resident macrophages primarily
produce ornithine via arginase (Arg). Middle: upon stimulation, macrophage production of iNOS/NO
increases. And the production of other cytokines, etc., changes during this time. Bottom: after 16 h,
macrophage NO production increases further and ornithine production declines.

two core functions that result from
macrophages sampling their environment,
and that affect health in very important, and opposite, ways: the M1/inhibit
response and the M2/heal response.
Inhibit-type macrophages and healtype macrophages were specifically
renamed M1 and M2 because these
macrophage responses [or dendritic cells1

first. They directly sense Pathogen or
Damage – Associated Molecular Patterns
(PAMP or DAMP) that can initiate M1
or M2-type responses (35–39). Subsequently, macrophages can stimulate T cells
(that cannot recognize antigens directly),
and thereby further amplify M1 or M2
responses. This “secondary” type of T celldriven response (macrophage “activation”)
was discovered by Mackaness and colleagues using mice preimmunized to Listeria and other pathogens (40). It was not
known at this time that macrophages were
actually responsible for initially activating
T cells (41, 42). The recent revelation about
the central role of macrophages in immune
responses caused a sea change in understanding how immune responses occur and
are regulated in vivo (1, 7).

(30–32)] were also found to stimulate T
cells to make Th1-type (cellular-based), or
Th2-type (antibody-based) cytokines (7),
respectively. This fourth SHIP function of
present (antigen) is only expressed in vertebrates (1). Although T cells can produce molecules that “activate” or “alternatively activate” macrophages (33, 34),
macrophages evolved first and respond

1 Both macrophages and dendritic cells can direct the type of T cell response. So, the word “macrophage” will be used here to refer to both. Readers are directed elsewhere
for discussions of macrophages and dendritic cells.

www.frontiersin.org

December 2014 | Volume 5 | Article 620 | 43

Mills et al.

TGF-β) that cause B cells to become
antibody-producing plasma cells (7, 29).
These same cytokines also inhibit the
M2 to M1 switch, and thus can amplify
M2/heal activity (1). Of course, because
there are many different pathogens invading different locales of hosts, there are
always mixtures of M1/Th1 or M2/Th2type responses as disease regression or progression occurs. In this connection, it is
now known that tissue macrophages can
arise from local renewal or from the blood
(1, 4). The ontogeny of M1 and M2type macrophages is not yet clear, and is
beyond the scope of this article. Recent
advances in metabolomics, defining resident microbiota, other areas, are opening
up new horizons for understanding the
myriad signals that regulate “immunity”
(43). Though more is to be known, the
aforementioned results have established
important biochemical and physiologic
elements that influence how macrophages
serve to protect (or fail to protect) against
infectious or other threats to host homeostasis.

IN VITRO VERSUS IN VIVO
MACROPHAGE CONUNDRUM
In addition to the basic macrophage functions necessary for life (such as SHIP),
some investigators (primarily working
in vitro) have employed ever-expanding
lists of “markers” for macrophage “activation.” These include: cell surface antigens;
expressed gene products; and other factors,
and have created the notion that there are
many different “varieties” of macrophages
such as “M2 a, b, c,”“regulatory,” and “alternatively activated” macrophages (49–53).
Unlike classifying macrophages by functions (e.g., M1/inhibit or M2/heal), the
use of markers has created subsets without clear functional roles in vivo. Likewise,
defining macrophage populations based on
cytokine production patterns has caveats
that are often overlooked. For example,
macrophage cytokines such as IL-6 are
“inflammatory,” yet they can be found in
almost any site where macrophages are
present (1). Indeed, the very presence of
macrophages is inflammatory that raises
questions about what “anti-inflammatory”
macrophages are (47–49). Efforts to define
macrophage “subsets” based on which
cytokine (or agonist) has been used to stimulate them in vitro (such as IL-4 or IFN-γ)

Macrophage SHIP of Immunity

also leads to confusion since macrophages
do not encounter isolated cytokines in vivo.
Rather, they are constantly receiving hundreds of signals, the integration of which
ultimately defines a cell’s behavior. Furthermore, because a selected cytokine
can elicit a given macrophage reaction
in vitro does not mean it has the same
effect in vivo. For example, adding IL4 to macrophages in vitro does increase
M2-type activity (50). And IL-4 from T
cells or innate cells can upregulate M2type antibody responses (7, 29): what has
been has been termed “alternative activation”). However, it is hard to ascribe
M2-type responses in circumstances such
as sterile wounds or tumors to “alternative activation” because little or no IL-4
is present (54, 55). Using T cell-derived
cytokines to stimulate macrophages in vitro
has also propagated the long-held notion
that T cells are necessary to “activate”
macrophages (23, 24). This perception
runs counter to the observations that
macrophages initiate and direct innate or
adaptive responses (1). Another potential artifact of in vitro cultures is that
macrophages can exhaust critical media
components, and thus behave in ways
(including dying) that are not observed
in vivo where nutrients/other products are
replenished (24).
Finally, the source of the “macrophages”
being studied in vitro varies and has created confusion. Specifically, people studying humans have primarily used monocytes from blood because of convenience.
And doing so has caused some to conclude there are major species differences
in “macrophages,” including that humans
seem less able (or unable) to produce
iNOS/NO or arginase/ornithine (3, 56, 57).
However, comparing monocyte-derived
macrophages to tissue macrophages is
an apples and oranges-type comparison.
When human tissue macrophages have
been examined, they do not appear fundamentally different from those of other
vertebrate species (58).
Thus, a variety of pitfalls can make it difficult to translate results from in vitro cultures to understanding how macrophages
function in vivo. In turn, rather than relying on “markers” or selected culture stimuli to try and define different macrophage
“activation” states (59), it seems prudent
to focus on characterizing macrophages

Frontiers in Immunology | Molecular Innate Immunity

by their known in vivo functions, such as
SHIP (1).

SHIP FUNCTIONS TO NAVIGATE A SEA
OF PLASTICITY
Macrophage SHIP functions are associated
with major differences in their metabolism and physiology (1). And hence, at the
population level, macrophages must display considerable heterogeneity. “Plasticity” usefully describes the unique adaptability of macrophages as they change
from, for example, producing a growthpromoting molecule (ornithine) to producing a growth-inhibiting molecule (NO)
(12, 60). However, for some the concept
of plasticity has morphed into a notion
that macrophages are a fluid cell type
that are always only changing (47–51).
Like they say, “change is good” (humor
intended). But, like changing clothes, it is
not the changing that matters: it is the
result. Perhaps, the clothes help one get
a job, or, get a date, etc. And so it is
with macrophages. As macrophages make
major switches in their metabolism, they
are “changing.” But, the changes in functional properties of macrophages can create illusory heterogeneity as illustrated in
Figure 1B. Specifically, if a population of
resting/resident macrophages (or a single
macrophage, left) receives appropriate signals (e.g., LPS and/or IFN-γ) and commits
to switching from M2/heal to M1/inhibit
dominant activity, it takes the cell(s) several
hours to accomplish this major change in
metabolism. In turn, at any given time there
will be a variety of different macrophages
expressing different M2 and (increasingly
in this example) M1-type activity. In turn,
if one examines macrophages (or a single macrophage) at any given time there
will be intermediate phenotypes in terms of
marker or cytokine expression. Also often
overlooked is that M1-type macrophages
produce non-specific killer molecules (like
NO) that inhibit or kill macrophages too
(24). In turn, analysis of whole populations can create the additional illusion
that M1-type have converted back to M2type, when actually, they are dead/missing
(1) In turn, examining macrophage populations (particularly in vitro) can create impressions of reversible plasticity or
heterogeneity, but which are not based
on what functions the macrophages have
(e.g., M1/inhibit or M2/heal) (49). Thus,

December 2014 | Volume 5 | Article 620 | 44

Mills et al.

heterogeneity (or plasticity) is a means to
an end. The “end” immunologists should
strive for is identifying macrophages by
their health-impacting functions (1).

SUMMARY
Immunology has and will continue to
cure important diseases. And, the ability to culture macrophages in vitro, the
expanding power of “transcriptome”analysis to examine thousands of genes, the
capability of analyzing single macrophages,
and other new technologies are providing necessary new information about the
cellular biochemistry and physiology of
leukocytes. But, as demonstrated here with
macrophages, overemphasis on ambiguous
“markers,” or analyzing whole populations
of macrophages that are changing their
functions, can create an illusion – a “sea of
plasticity.” Therefore, to navigate this sea,
it is critical to focus on SHIP functions
(e.g., sample, heal, inhibit, and present)
that importantly affect health. Doing so
will help unleash the tremendous potential for usefully modulating innate immunity/macrophages against a variety of conditions ranging from cancer to atherosclerosis. To cure disease.

ACKNOWLEDGMENTS
We appreciate the excellent and unselfish
input we have received from researchers in
forming this paper. Namely, because we are
all on the“same team”: the goal being better
health.

REFERENCES
1. Mills CD, Ley K. M1 and M2 macrophages: the
chicken and the egg of immunity. J Innate Immun
(2014) 6:716–26. doi:10.1159/000364945
2. Wynn TA, Chawla A, Pollard JW. Macrophage
biology in development, homeostasis and disease. Nature (2013) 496:445–55. doi:10.1038/
nature12034
3. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev Immunol
(2012) 32:463–88. doi:10.1615/CritRevImmunol.
v32.i6.10
4. Geissmann F, Manz MG, Jung S, Sieweke MH,
Merad M, Ley K. Development of monocytes,
macrophages, and dendritic cells. Science (2010)
327:656–61. doi:10.1126/science.1178331
5. Cooper EL. eCAM: Darwin and Metchnikoff. Evid
Based Complement Alternat Med (2009) 6:421–2.
doi:10.1093/ecam/nep194
6. Dzik JM. The ancestry and cumulative evolution
of immune reactions. Acta Biochim Pol (2010)
57:443–66.
7. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill
AM. M-1/M-2 macrophages and the Th1/Th2

www.frontiersin.org

Macrophage SHIP of Immunity

paradigm. J Immunol (2000) 164:6166–73. doi:10.
4049/jimmunol.164.12.6166
8. Morris SM. Recent advances in arginine metabolism: roles and regulation of the arginases. Br
J Pharmacol (2009) 6:922–30. doi:10.1111/j.14765381.2009.00278.x
9. Munder M. Arginase: an emerging key player in
the mammalian immune system. Br J Pharmacol
(2009) 158:638–51. doi:10.1111/j.1476-5381.2009.
00291.x
10. Hibbs JB, Taintor RR, Vavrin Z, Rachlin EM.
Nitric oxide: a cytotoxic activated macrophage
effector molecule. Biochem Biophys Res Commun
(1988) 157:87–94. doi:10.1016/S0006-291X(88)
80015-9
11. Nathan CF. Nitric oxide as a secretory
product of mammalian cells. FASEB J (1992)
12:3051–64.
12. Mills CD. Macrophage arginine metabolism to
ornithine/urea or nitric oxide/citrulline: a life or
death issue. Crit Rev Immunol (2001) 21:399–426.
doi:10.1615/CritRevImmunol.v21.i5.10
13. Albina JE, Mills CD, Henry WL Jr, Caldwell MD.
Temporal expression of different pathways of 1arginine metabolism in healing wounds. J Immunol
(1990) 144:3877–80.
14. Mills CD, Shearer J, Evans R, Caldwell MD.
Macrophage arginine metabolism and the inhibition or stimulation of cancer. J Immunol (1992)
149:2709–14.
15. Komohara Y, Jinushi M, Takeya M. Clinical significance of macrophage heterogeneity in human
malignant tumors. Cancer Sci (2014) 105:1–8.
doi:10.1111/cas.12314
16. Noy
R, Pollard
JW.
Tumor-associated
macrophages: from mechanisms to therapy.
Immunity (2014) 41:49-61. doi:10.1016/j.immuni.
2014.06.010
17. Tang X, Mo C, Wang Y, Wei D, Xiao H. Anti-tumour
strategies aiming to target tumour-associated
macrophages. Immunology (2012) 138:93–104.
doi:10.1111/imm.12023
18. Weigert A, Brune B. Nitric oxide, apoptosis and
macrophage polarization during tumor progression. Nitric Oxide (2008) 2:95–102. doi:10.1016/j.
niox.2008.04.021
19. Nathan C. Role of iNOS in human host defense.
Science (2006) 312:1874–5. doi:10.1126/science.
312.5782.1874b
20. Hibbs JB. Infection and nitric oxide. J Infect Dis
(2002) 185:S9–17. doi:10.1086/338005
21. Vogel SN, Johnson D, Perera PY, Medvedev A, Lariviere L, Qureshi ST, et al. Cutting edge: functional
characterization of the effect of the C3H/HeJ defect
in mice that lack an Lpsn gene: in vivo evidence for
a dominant negative mutation. J Immunol (1999)
162:5666–70.
22. Reiss CS, Komatsu T. Does nitric oxide play a
critical role in viral infections? J Virol (1998)
78:4547–51.
23. Stevens RB, Ansite JD, Mills CD, Lokeh A, Rossini
TJ, Saxena M, et al. Nitric oxide mediates early dysfunction of rat and mouse islets after transplantation. Transplantation (1996) 12:1740–9. doi:10.
1097/00007890-199606270-00014
24. Albina JE, Caldwell MS, Henry WL, Mills CD. Regulation of macrophage functions by L-arginine. J
Exp Med (1989) 169:1021–9. doi:10.1084/jem.169.
3.1021

25. Nathan CF. Secretory products of macrophages.
J Clin Invest (1987) 79:319–26. doi:10.1172/
JCI112815
26. Hibbs JB, Vavrin Z, Taintor RR. L-arginine
is required for expression of the activated
macrophage effector mechanism causing selective
metabolic inhibition in target cells. J Immunol
(1987) 138:550–65.
27. Li Y, Karlin A, Loike JD, Silverstein SC. Determination of the critical concentration of neutrophils required to block bacterial growth in tissues. J Exp Med (2004) 200:613–22. doi:10.1084/
jem.20040725
28. Filipin L, Moreira AJ, Marroni NP, Xavier RM.
Nitric oxide and repair of skeletal muscle injury.
Nitric Oxide (2009) 3:157–63. doi:10.1016/j.niox.
2009.08.002
29. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat
Rev Immunol (2011) 11:723–37. doi:10.1038/
nri3073
30. Geissmann F, Gordon S, Hume DA, Mowat DA,
Randolph GJ. Unravelling mononuclear phagocyte
heterogeneity. Nat Rev Immunol (2010) 10:453–60.
doi:10.1038/nri2784
31. Hume DA. Macrophages as APC and the dendritic cell myth. J Immunol (2008) 181:5829–35.
doi:10.4049/jimmunol.181.9.5829
32. Steinman RM, Banchereau J. Taking dendritic cells
into medicine. Nature (2007) 449:419–26. doi:10.
1038/nature06175
33. Hsieh CS, Macatonia SE, O’Garra A, Murphy KM. T cell genetic background determines
default T helper phenotype development in vitro. J
Exp Med (1995) 181:713–21. doi:10.1084/jem.181.
2.713
34. O’Garra A, Murphy KM. Role of cytokines in
determining T-lymphocyte function. Curr Opin
Immunol (1994) 6:458–66. doi:10.1016/09527915(94)90128-7
35. Beutler BA. TLRs and innate immunity. Blood
(2009) 113:1399–407. doi:10.1182/blood-200807-019307
36. Kawai T, Akira S. Toll-like receptors and their
crosstalk with other innate receptors in infection and immunity. Immunity (2011) 34:637–50.
doi:10.1016/j.immuni.2011.05.006
37. Janeway CA Jr, Medzhitov R. Innate immune
recognition. Annu Rev Immunol (2002)
20:197–216.
doi:10.1146/annurev.immunol.
20.083001.084359
38. Srikrishna G, Freeze HH. Endogenous damageassociated molecular pattern molecules at the
crossroads of inflammation and cancer. Neoplasia
(2009) 11:615–28. doi:10.1593/neo.09284
39. Matzinger P. An innate sense of danger. Ann N
Y Acad Sci (2002) 961:341–2. doi:10.1111/j.17496632.2002.tb03118.x
40. Mackaness GB. The immunologic basis of acquired
cellular resistance. J Exp Med (1964) 120:105–20.
doi:10.1084/jem.120.1.105
41. Shevach EM, Rosenthal AM. Function of
macrophages in antigen recognition by guinea pig
T lymphocytes. II. Role of the macrophage in
the regulation of genetic control of the immune
response. J Exp Med (1973) 138:1213–29. doi:10.
1084/jem.138.5.1213
42. Unanue ER. Antigen-presenting function of
the macrophage. Annu Rev Immunol (1984)

December 2014 | Volume 5 | Article 620 | 45

Mills et al.

43.

44.

45.

46.

47.

48.

2:395–428.
doi:10.1146/annurev.iy.02.040184.
002143
Dorrestein PC, Mazmanian SK, Knight R. Finding the missing links among metabolites, microbes,
and the host. Immunity (2014) 40:824–32. doi:10.
1016/j.immuni.2014.05.015
Bae YS, Lee JH, Choi SH, Kim S, Almazan
F, Witztum JL, et al. Macrophages generate
reactive oxygen species in response to minimally oxidized low-density lipoprotein: toll-like
receptor 4- and spleen tyrosine kinase-dependent
activation of NADPH oxidase 2. Circ Res
(2009) 104:210–8. doi:10.1161/CIRCRESAHA.
108.181040
Nathan CF. Macrophage activation: some
questions. Ann Inst Pasteur Immunol (1986)
137:345–51. doi:10.1016/S0771-050X(86)80051-1
Hsieh CS, Macatonia SE, Tripp CS, Wolf SF,
O’Garra A, Murphy KM. Development of TH1
CD4+ T cells through IL-12 produced by Listeriainduced macrophages. Science (1993) 260:547–9.
doi:10.1126/science.8097338
Munder M, Mallo M, Eichmann K, Modolell M. Murine macrophages secrete interferon gamma upon combined stimulation
with interleukin (IL)-12 and IL-18: a novel
pathway of autocrine macrophage activation. J Exp Med (1998) 12:2103–8.
doi:10.1084/jem.187.12.2103
Buhtoiarov IN, Lum H, Berke G, Paulnock
DM, Sondel PM, Rakhmilevich AL. CD40 ligation activates murine macrophage via an
IFN-gamma-dependent mechanism resulting in
tumor cell destruction in vitro. J Immunol
(2005) 174:6013–22. doi:10.4049/jimmunol.174.
10.6013

Macrophage SHIP of Immunity

49. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation. Nat Rev Immunol
(2008) 8:958–69. doi:10.1038/nri2448
50. Fleming
BD, Mosser
DM.
Regulatory
macrophages: setting the threshold for therapy. Eur J Immunol (2011) 41:2498–502.
doi:10.1002/eji.201141717
51. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine system in diverse
forms of macrophage activation and polarization.
Trends Immunol (2004) 25:677–86. doi:10.1016/j.
it.2004.09.015
52. Stein M, Keshav S, Harris N, Gordon S. Interleukin
4 potently enhances murine macrophage mannose
receptor activity: a marker of alternative immunologic macrophage activation. J Exp Med (1992)
176:287–92. doi:10.1084/jem.176.1.287
53. Martinez FO, Gordon S, Locati M, Mantovani A.
Transcriptional profiling of the human monocyteto-macrophage differentiation and polarization:
new molecules and patterns of gene expression. J Immunol (2006) 177:7303–11. doi:10.4049/
jimmunol.177.10.7303
54. Daley JM, Brancato SK, Thomay AA, Reichner
JS, Albina JE. The phenotype of murine wound
macrophages. J Leukoc Biol (2010) 87:59–67. doi:
10.1189/jlb.0409236
55. Wynn TA, Ramalingam TR. Mechanisms of
fibrosis: therapeutic translation for fibrotic disease.
Nat Med (2012) 18:1028–40. doi:10.1038/nm.2807
56. Bogdan C. Species differences in macrophage NO
production are important. Nat Immunol (2002)
3:102. doi:10.1038/ni0202-102a
57. Fang FC, Nathan C. Man is not a mouse:
reply. J Leukoc Biol (2007) 81:580. doi:10.1189/jlb.
1206715

Frontiers in Immunology | Molecular Innate Immunity

58. Thomas A, Matilla J. Of mice and men:
arginine metabolism in macrophages. Front
Immunol (2014) 5:479. doi:10.3389/fimmu.2014.
00479
59. Murray PJ, Allen JE, Biswas SK, Fisher EA,
Gilroy DW, Goerdt S, et al. Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41:14–20.
doi:10.1016/j.immuni.2014.06.008
60. Stout RD, Suttles J. Functional plasticity of
macrophages: reversible adaptation to changing microenvironments. J Leukoc Biol (2004)
76:509–13. doi:10.1189/jlb.0504272

Conflict of Interest Statement: The authors declare
that the research was conducted in the absence of any
commercial or financial relationships that could be
construed as a potential conflict of interest.
Received: 29 October 2014; accepted: 20 November 2014;
published online: 04 December 2014.
Citation: Mills CD, Thomas AC, Lenz LL and Munder
M (2014) Macrophage: SHIP of Immunity. Front.
Immunol. 5:620. doi: 10.3389/fimmu.2014.00620
This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology.
Copyright © 2014 Mills, Thomas, Lenz and Munder.
This is an open-access article distributed under the terms
of the Creative Commons Attribution License (CC BY).
The use, distribution or reproduction in other forums is
permitted, provided the original author(s) or licensor are
credited and that the original publication in this journal
is cited, in accordance with accepted academic practice.
No use, distribution or reproduction is permitted which
does not comply with these terms.

December 2014 | Volume 5 | Article 620 | 46

REVIEW ARTICLE
published: 17 October 2014
doi: 10.3389/fimmu.2014.00514

From monocytes to M1/M2 macrophages: phenotypical vs.
functional differentiation
Paola Italiani * and Diana Boraschi
Laboratory of Innate Immunity and Cytokines, Institute of Protein Biochemistry, National Research Council, Napoli, Italy

Edited by:
Klaus Ley, La Jolla Institute for Allergy
and Immunology, USA
Reviewed by:
Fulvio D’Acquisto, Queen Mary
University of London, UK
Gaurav K. Gupta, Harvard Medical
School, USA
*Correspondence:
Paola Italiani , Laboratory of Innate
Immunity and Cytokines, Institute of
Protein Biochemistry, National
Research Council, Via Pietro
Castellino 111, Napoli 80131, Italy
e-mail: italianipaola@gmail.com

Studies on monocyte and macrophage biology and differentiation have revealed the
pleiotropic activities of these cells. Macrophages are tissue sentinels that maintain tissue integrity by eliminating/repairing damaged cells and matrices. In this M2-like mode,
they can also promote tumor growth. Conversely, M1-like macrophages are key effector
cells for the elimination of pathogens, virally infected, and cancer cells. Macrophage differentiation from monocytes occurs in the tissue in concomitance with the acquisition of
a functional phenotype that depends on microenvironmental signals, thereby accounting
for the many and apparently opposed macrophage functions. Many questions arise. When
monocytes differentiate into macrophages in a tissue (concomitantly adopting a specific
functional program, M1 or M2), do they all die during the inflammatory reaction, or do some
of them survive? Do those that survive become quiescent tissue macrophages, able to
react as naïve cells to a new challenge? Or, do monocyte-derived tissue macrophages conserve a “memory” of their past inflammatory activation? This review will address some
of these important questions under the general framework of the role of monocytes and
macrophages in the initiation, development, resolution, and chronicization of inflammation.
Keywords: monocytes, monocyte-derived macrophages, tissue-resident macrophages, functional phenotypes,
inflammation

INTRODUCTION
In the healthy organism, the innate immune system provides the
first line of defense against external or internal danger signals, by
initiating a protective inflammatory response that develops during
time through different phases, from initiation and full inflammation, to resolution and re-establishment of tissue integrity. The
first phase of an inflammatory response is aimed at destroying
pathogens, and is followed by a phase in which dead and dying
cells, damaged extracellular matrix material, and cellular debris
are removed, to end up with a recovery phase in which the tissue
is repaired and restored to a healthy fully functional condition. In
fact, if the defense against harmful threats is a priority for avoiding
tissue damage, maintaining homeostasis (i.e., maintaining tissue
morphology and tissue function) is the ultimate goal of a tissue
in multicellular organisms (1). In this perspective, inflammation
presumably evolved as an adaptive response to tissue malfunction
or homeostatic imbalance (2). Thus, while the disease state is a displacement from homeostasis, inflammation is the tissue response
for restoring homeostasis. However, since the inflammatory activities are potentially harmful to the host, these need to be tightly
controlled to avoid excessive tissue damage (3).
The mononuclear phagocyte system (MPS) plays major roles
in development, scavenging, inflammation, and anti-pathogen
defenses, both by the direct elimination of foreign agents and
in organizing each different phase of the inflammatory process
(4). Under the term, MPS are grouped lineage-committed bone
marrow precursors, circulating monocytes, resident macrophages,
and dendritic cells (DC) (5). The development, homeostatic
maintenance, proliferation, differentiation, and function of the

www.frontiersin.org

MPS are regulated by the growth factors colony-stimulating factor (CSF)-1 and interleukin (IL)-34, the second ligand for the
CSF-1R (6, 7).
The issue of heterogeneity in the MPS still leads to a confusion
and debate about DC as truly distinct cells from macrophages,
with separate lineage and functions (8). In fact, macrophages and
myeloid DC possibly represent alternative differentiation options
of bone marrow progenitors and blood monocytes (9), with overlapping functions and marker expression. Reviewing this issue
is beyond the scope of this essay [we refer the reader to recent
excellent reviews on the topic; (9–11)], and will only focus on
monocytes and their relationship with macrophages.
The traditional view of the MPS suggests that recruited monocytes (that become macrophages in tissues) are key players during
inflammation and pathogen challenge, whereas tissue-resident
macrophages have important roles in development, tissue homeostasis, and the resolution of inflammation. A basic concept of
the MPS is that blood monocytes are precursors that replace tissue macrophages within a single developmental lineage (4). This
dogma needs now to be revised in the light of new evidence that
macrophages are endowed with self-renewal capacity and can populate tissues before birth, deriving from early hematopoiesis in the
yolk sac (12, 13). The discovery of new macrophage progenitors
of embryonic origin forces us to reassess definitions, functions,
and cell–cell relationships within the MPS. We can synthesize it in
three key new questions:
1. Are monocytes more than circulating precursors and can they
have effector functions?

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Italiani and Boraschi

2. Is there a functional difference between the monocyte-derived
macrophages and the yolk sac-derived self-renewing resident
macrophages?
3. What is the relationship between monocytes and macrophages
and which are their mutual roles in different phases of
inflammatory reactions?
Another new perspective arises from the description of
macrophage polarization, i.e., the ability of macrophages to
acquire different functional phenotypes, enabling them to steer
adaptive immunity in different directions. This highlights the
central role of macrophages in immune defense, overturning
the long-held notion that macrophages need to be activated by
T-cells (14).
This review will summarize what has been so far investigated
and established on monocyte/macrophage biology, highlighting
what remains outstanding, and which questions are still unanswered. We will consider key studies that have been carried out in
mice, with reference to the human situation when data are available. We will review the various aspects (monocyte recruitment,
monocyte functions, macrophage polarization) before (homeostatic conditions), during (inflammatory reaction), and after a
damaging event (resolution/repair).

MONOCYTES
MONOCYTE DEVELOPMENT AND HETEROGENEITY

Monocytes are a group of cells circulating in the blood, bone
marrow, and spleen, and constituting ~10% of the total leukocytes in human beings and only 2–4% in mice. They have typical morphological features, such as irregular cell shape, oval- or
kidney-shaped nucleus, cytoplasmic vesicles, and high cytoplasmto-nucleus ratio. Monocytes can remain in the circulation for up
to 1–2 days, after which time, if they have not been recruited into
a tissue for facing a danger, they die and are removed. Monocytes originate in the bone marrow from hematopoietic stem cells
(HSCs) and develop through a series of sequential differentiation stages: the common myeloid progenitor (CMP) (15), the
granulocyte-macrophage progenitor (GMP) (15), the common
macrophage and DC precursor (MDP) (16), and finally the committed monocyte progenitor (cMoP), a recently identified bone
marrow precursor that differs from MDP as it lacks CD135 expression (17). MDP gives rise also to common DC progenitors (CDP),
whose differentiation potential is restricted to the DC lineage
(18). Monocytes have been considered as the systemic reservoir
of myeloid precursors for renewal of tissue macrophages and DC.
However, many DC and macrophage subpopulations [for example, lymphoid organ DC, plasmacytoid DC, skin Langerhans cells
(LC), and brain microglia] originate from the MDP independent
of monocytes (11, 18), and in some cases, they can even develop
directly from the bone marrow (19).
Homeostatic control of monocyte/macrophage development is
mostly influenced by CSF-1 (also known as M-CSF), produced by
stromal cells within the blood and in tissues (20). Mature mononuclear phagocytes in turn express CSF-1 receptors (CSF-1R) and
remove circulating CSF-1, allowing a feedback loop responsible for monocyte proliferation decrease (21, 22). Recently, the
cytokine IL-34 has been identified as able to bind and signal

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

through the CSF-1R (6, 23). Unlike broadly expressed CSF-1,
IL-34 expression is restricted to the epidermis and central nervous system (24), where it supports the steady-state proliferation
of macrophages (LC and microglia, respectively). Granulocytemacrophage colony-stimulating factor (GM-CSF) is another factor involved in the development of mononuclear phagocytes but
only during the inflammatory state and not under homeostatic
conditions (25, 26).
Abundant experimental evidence indicates that recruited
monocytes are innate effectors of the inflammatory response to
microbes, and they kill pathogens via phagocytosis, production
of reactive oxygen species (ROS), nitric oxide (NO), myeloperoxidase, and inflammatory cytokines (27). In some circumstances,
monocytes can trigger and polarize T-cell responses (27, 28) and
may also contribute to angiogenesis and atherogenesis (29).
Human peripheral blood monocytes are not a homogeneous
population. Monocyte heterogeneity was first reported with
the identification of a minor population of CD16 (FcγRIII)expressing cells within circulating human monocytes (30). In
recent years, investigators have identified three functional subsets of human monocytes, the characterization of which is still
in its infancy. Likewise, it is still unclear which are the specific
roles that they exert in homeostasis and inflammation in vivo,
in comparison with those of the previously described classically
and alternatively activated macrophages (see below). The new
nomenclature that groups monocytes into three subsets, based
on the expression of the surface markers CD14 and CD16, has
recently been approved by the Nomenclature Committee of the
International Union of Immunologic Societies (31). Based on
this nomenclature, the major population of human monocytes
(90%) with high CD14 but no CD16 expression (CD14++ CD16−
or CD14+ CD16− ) are termed classical monocytes, whereas the
minor population of human monocytes (10%) is further subdivided into the intermediate subset, with low CD16 and high
CD14 (CD14++ CD16+ or CD14+ CD16+ ), and the non-classical
subset, with high CD16 but with relatively lower CD14 expression
(CD14+ CD16++ or CD14dim CD16+ ) (31). In this review, we refer
only to the main difference, terming classical monocytes simply as
CD14+ , and non-classical as CD16+ .
Over the recent years, an increasing amount of knowledge
has been gained in the field of monocyte subpopulations. Many
authors demonstrated that the three subsets express different transcriptomes (32–38), although discrepancies between studies were
evident. These discrepancies may be due to differences in cell isolation methodology and in the purity of the cell populations isolated,
and the microarray methodologies, which use different amounts
of total RNA for the hybridization, different probes to identify
the genes, and even distinct solid supports for the probes (39).
However, there is stronger agreement for the proximity of relationship between the intermediate and non-classical monocyte
subsets, while the classical subset is the most distant subset (36).
The close relationship between intermediate and non-classical
monocytes suggests a direct developmental relationship between
them, although this has yet to be formally proven. Also, it needs to
be clarified how the characteristics previously ascribed to CD16+
monocytes are distributed between intermediate and non-classical
subsets (36). Recent data suggested a sequential developmental

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Monocyte to macrophage differentiation

relationship between the two subsets based on the observation
that, in time course studies in inflammatory diseases, an increase
in the intermediate monocytes is followed by an increase of
non-classical monocytes (40).
The physiological role of the monocyte subsets in vivo is not
fully defined. They might have different roles during the homeostasis, immune defense/inflammation, and tissue repair, in terms
of their capacity to become activated and secrete inflammatory
cytokines in response to different stimuli, antigen processing and
presentation, pro-angiogenic and patrolling behavior. The phenotypic and functional differences between the monocyte subsets were recently discussed in an exhaustive review (41). The
authors of this review report a complete and referenced list
of studies on bacterial and viral infections, autoimmune diseases, and inflammatory conditions, in which an expansion of
CD16+ cells in respect to other subsets has been observed. In
general terms, both human classical and intermediate monocytes
have inflammatory properties reminiscent of the murine Ly6C+
monocytes (also termed “inflammatory” monocytes) (42), while
non-classical monocytes display patrolling properties similar to
those of murine Ly6C− monocytes (also termed “alternative” or
“patrolling” monocytes) (43). Both human and mouse inflammatory monocytes express high levels of the chemokine receptor
CCR2 and low levels of the chemokine receptor CX3CR1, whereas
patrolling monocytes show a reverse pattern. Accordingly, inflammatory monocytes respond to the chemokine CCL2 that mediates
Ly6C+ /CD14+ monocyte recruitment to inflammatory sites (44),
while patrolling monocytes respond to CX3C-chemokine ligand
1 [CX3CL1, the human fractalkine and mouse neurotactin; (45)],
a chemokine present both as soluble protein and as membranebound chemokine form that is expressed on endothelial cells and
in tissues. Overall, it is clear that the subsets between human being
and mouse are similar but not identical (42, 46). Table 1 summarizes the main features of monocytes in human beings and mice.
Of note, there is a clear difference in the proportion of the two

monocyte subsets, as Ly6C− cells represent about half of the circulating monocytes in mice, whereas CD16+ monocytes account for
less than 15% in human beings (30). However, Ziegler-Heitbrock
hypothesized that the higher proportion of the Ly6C− in mouse
blood could be due to stressful blood drawing (cardiac puncture
under terminal anesthesia) that mobilizes these monocytes from
the marginal pool (46). This hypothesis still needs experimental
proof.
To date, a relevant question that is still open concerns the origin
of the various monocyte subpopulations. It should be kept in mind
that the majority of current knowledge derives from mouse studies. It is unknown if the monocyte subpopulations are end stages of
different differentiation paths of a common precursor, or whether
they represent subsequent maturation stages in a common path of
differentiation, where the intermediate subset could be a phenotypical and/or developmental intermediate between the classical
and non-classical subsets. The latter hypothesis seems to be the
most reliable. While initial studies suggested that Ly6C+ cells were
recruited under inflammatory conditions and did not serve as precursors to Ly6C− cells [which in turn were originally considered
the immediate precursors of resident macrophages; (43)], recent
evidence suggests that, in steady state, Ly6C+ monocytes are precursors of Ly6C− monocytes (48, 49), as shown in experiments in
which grafted Ly6C+ monocytes spontaneously differentiated into
Ly6C− in the blood of recipient mice (48). This conversion can also
occur in the bone marrow, where Ly6C+ monocytes apparently
return in the absence of inflammation (47–49). More recently, it
has been suggested that CSF-1R signaling was required for the
maturation of monocytes from Ly6C+ to Ly6C− , as blockade of
this receptor leads to decrease in the number of Ly6C− cells, shortens their lifespan (48), and concomitantly increases the number
of Ly6C+ monocytes (50).
It has also been observed that development of the Ly6C− population depends on the transcription factor NR4A1 (Nurr77) (51).
NR4A1 deletion alters the number of Ly6C− monocytes in the

Table 1 | Human and murine monocyte subsets.
Species

Subseta

% In WB

% In blood

Half-life

Markers

monocytes
Human

Classical

~10%

85%

1–2 days

CD14++ CD16−

Chemokine

Other surface

receptors

markers

CCR2+ CX3CR1−

CD62L+ , CD64− , MHC

Phagocytosis,

class II+ , CD163+

inflammatory effectors

CD62L+ , CD64− , MHC

Inflammatory effectors

being
Intermediate

5%

–

CD14++ CD16+

CCR2− CX3CR1+

Main functions

class II++ , CD163+
Non-classical

10%

–

CD14+ CD16++

CCR2− CX3CR1+

CD62L− , CD64+ , MHC

Patrolling, antiviral role

class II++ , CD163−
Mouse

Ly6Clow
Ly6Chigh

4%

~60%
~40%

18–20 h
5–7 days

CD11b+ CD115+ CCR2+ CX3CR1−

F4/80+ , CD62L− , MHC

Ly6C+

class IIb , CD43+

Inflammatory effectors

CD11b+ CD115+ CCR2− CX3CR1+

F4/80+ , CD62L+ , MHC

Patrolling, tissue repair

Ly6C−

class IIb , CD43−

Phagocytosis,

a

Work by Sunderkötter et al. (47) characterized a population of Ly6Cmed monocytes with intermediate features between Ly6C+ and Ly6C− . These are not included in

the table, because this population remains poorly characterized in terms of both phenotype and function.
b

Inducible.

WB, whole blood; Ly6C, lymphocyte antigen 6 complex; CCR2, chemokine (C-C motif) receptor 2; CX3CR1, CX3C-chemokine receptor 1.

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Italiani and Boraschi

bone marrow but not in blood or spleen (52), but does not alter
the number of macrophages within tissues (53). This suggests that
either Ly6C− monocytes can develop from MDP within the bone
marrow, or that Ly6C− monocytes are a functional end stage. In
this regard, given that Ly6C− monocytes exhibit a long steady-state
half-life of 5–7 days [which in the absence of their renewal from
Ly6C+ monocytes can extend to 2 weeks; (48)] compared to ~8 h
for Ly6C+ cells, Ly6C− monocytes might be considered as terminally differentiated blood-resident macrophages or “vasculature
macrophages,” rather than bona fide monocytes (48). Indeed, the
primary function of these cells seems to be that of patrolling the
vascular endothelium and monitoring its integrity (45, 51). Conversely, and in parallel with human CD14+ cells, Ly6C+ monocytes
because of their short half-life are unlikely to have other functions,
and thus are more likely to be the direct precursors of the tissue macrophages/peripheral mononuclear phagocytes described
in the original MPS model.
FUNCTION OF MONOCYTE SUBSETS DURING HOMEOSTASIS
(CLASSICAL VS. ANTIGEN-PRESENTING TISSUE MONOCYTES VS.
PATROLLING MONOCYTES)

The original concept of MPS implicated that classical monocytes are recruited in the tissue to become tissue-resident
macrophages in homeostatic conditions, and inflammatory activated macrophages during an infection (27, 54). We will examine
more in detail the role of recruited cells during the inflammatory response later, while here we will focus on the recruitment of
monocytes in homeostasis and their contribution to maintaining
the pool of tissue macrophages. In order to avoid misunderstandings, it is important to agree on the definition of monocyte. In
our view, bona fide monocytes are restricted to the blood compartment, and to the bone marrow and spleen (55), where they
wait to be released in the blood. For obvious reasons, in both
these compartments, monocytes should not initiate any inflammatory reaction, but they must be ready to be recruited into
the blood first and subsequently to all organs and tissues. A
phenomenon was recently reported, termed “anticipatory inflammation,” whereby Ly6C+ classical monocytes are released from
the bone marrow in diurnal rhythmic waves under the control
of circadian gene Bmal1 (or Arntl) (56) to provide an adequate
innate response to environmental challenges that are expected
to occur with a evolutionarily predicted frequency. Despite new
evidence supports the view that Ly6C+ classical monocytes are
not precursors of resident macrophages in all tissues and during certain types of inflammation (see below), it is clear that
circulating monocytes contribute to the repopulation of tissueresident macrophages under homeostatic conditions in tissues
like the lamina propria of the small intestine and healthy skin.
Studies based on functional and lineage tracing and adoptive
transfer have revealed that Ly6C+ monocytes are precursors of
intestinal macrophages that have a short half-life of only 3 weeks
(57–59). Conversely, in the dermis are present both resident dermal macrophages and monocyte-derived macrophages (60, 61). A
recent work suggests that the number of macrophages is partially
replenished by monocytes also in the heart (62) and in the lung
(63). It is unknown why some tissue macrophages are constantly
maintained by circulating monocytes, whereas other populations

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

are independent on circulating monocytes (see below). The notion
that monocyte-derived macrophages derive from Ly6C+ cells suggests that the repopulation/maintenance of resident macrophages
in steady-state conditions follows the same mechanism as that
occurring during inflammation.
The function of Ly6C+ monocytes in circulation remains
poorly defined. In the attempt to identify an effective role of
monocytes in the blood in homeostatic conditions (besides being
precursor cells), a recent work has suggested a distinct surveillance phenotype for Ly6C+ monocytes (64). These monocytes
can enter non-lymphoid organs without obligatory differentiation
into macrophages or DC. The authors propose that these monocytes can upregulate MHC class II expression and subsequently
recirculate to lymph nodes, where they are able to present antigens
to T-cells. Considering that these cells retain a monocyte-like gene
expression profile, the authors term them “tissue monocytes” (64).
This study contributes to revising the role of circulating monocytes, suggesting that they are not only precursors of macrophages
but also effector cells.
Regarding the role of the Ly6C− subset in the blood in steadystate conditions, intravital microscopy studies have established
that these cells display a “patrolling” phenotype, being able to
crawl on the luminal surface of the vascular endothelium (45,
51). This patrolling behavior, along with the ability to phagocytose endothelial-associated particles, suggests that a primary role
of these monocytes is sensing and scanning the endothelial surface
for damage and/or the presence of pathogens (51). The patrolling
monocytes mainly respond via Toll-like receptor 7 (TLR7) to local
danger signals (while they are poorly responsive to bacterial products such as LPS) by producing inflammatory mediators (51). They
are able to induce the intravascular recruitment of neutrophils,
which trigger endothelial necrosis, and subsequently they clear the
resulting debris (51). A similar patrolling feature and TLR7/TLR8dependent reactivity were also detected in human CD14+ CD16++
monocytes (35).
Consistent with their functional role of surveillance of the
endothelium integrity and with the fact that they are terminally
differentiated cells, we agree with the view that Ly6C− /CD16+
can be considered as the tissue-resident macrophages of the blood.
Regarding their ability to produce inflammatory factors, we speculate that the patrolling monocytes have a higher activation threshold than Ly6C+ monocytes; therefore, they should be able to
produce an amount of inflammatory cytokines and chemokines
sufficient for coordinating the repair of a damaged endothelium,
but not enough to initiate a strong inflammatory reaction.
A summary of the roles of monocyte subsets in steady state vs.
inflammatory conditions is reported in Table 2.

MACROPHAGES
TISSUE-RESIDENT MACROPHAGE DEVELOPMENT IN STEADY STATE:
EMBRYONIC ORIGIN VS. MONOCYTE DERIVATION

Resident macrophages are heterogeneous and versatile cells found
in virtually all tissues of adult mammals, where they can represent
up to 10–15% of the total cell number in quiescent conditions.
This number can increase further in response to inflammatory stimuli. The specialization of macrophages in particular
microenvironments explains their heterogeneity. Macrophages

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Monocyte to macrophage differentiation

take different names according to their tissue location, such as
osteoclasts (bone) (see Box 1), alveolar macrophages (lung),
microglial cells (CNS), histiocytes (connective tissue), Kupffer
cells (liver), and LC (skin). These populations have such highly
different transcriptional profiles that they could be considered
as many different and unique classes of macrophages (74). On
the other hand, the functions of macrophages are the same in
all tissues. They are key players in tissue development (by shaping the tissue architecture), in immune response to pathogens (by
generating and resolving the inflammatory reaction), in surveillance and monitoring of tissue changes (by acting as sentinel and
effector cells), and especially in maintenance of tissue homeostasis
(by clearing apoptotic or senescent cells, and by remodeling and
repairing tissues).

Table 2 | Functions of monocyte subsets in steady state and
inflammatory conditions.
Subset

Ly6C+

Function
Steady-state conditions

Inflammation

Replenishment of

Differentiation in M1-like

monocyte-derived

functional phenotype and

macrophages in the tissue (gut, initiation of the inflammatory
skin, heart, and lung)

response

Differentiation in Ly6C− cells in

Antigen uptake in the tissue,

the blood and in the bone

recirculation to lymph nodes,

marrow

antigen presentation in lymph
nodes (“tissue monocytes”)

Ly6C−

Patrolling and surveillance of

Promotion of healing in

the luminal surface of the

ischemic myocardium, and

endothelium

tissue repair during infection

Sensing viral nucleic acids

with Listeria monocytogenes

The view that tissue macrophages originate from circulating
peripheral blood monocytes that migrate into tissues under a variety of stimuli, proposed and strongly supported by van Furth in
the 1970s (4, 75, 76), needs to be reconsidered. In addition to
a wealth of old data (77, 78), two new pieces of evidence have
further weakened the view that monocytes are the precursors
of tissue macrophages in steady-state conditions: (1) the finding of the macrophage origin from embryonic progenitors that
seed developing tissues before birth and give rise to fetal tissue
macrophages (79) and (2) the self-maintaining ability of tissueresident macrophages through local proliferation in adulthood
(13). The latter finding will be discussed hereafter.
Two main phases of embryonic hematopoiesis have been
described in the mouse: primitive hematopoiesis and definitive
hematopoiesis. The former takes place in the ectoderm of the
yolk sac and gives rise to macrophages without going through
a monocytic progenitor. The latter takes place in the fetal liver,
which is initially seeded by hematopoietic progenitors from the
yolk sac and subsequently by HSCs from endothelium of the
aorta-gonads-mesonephros (80, 81). The fetal liver subsequently
becomes the source of definitive hematopoiesis that generates circulating monocytes during embryogenesis. Spleen and bone marrow are also colonized via the circulatory system by hematopoietic
progenitors that will ultimately differentiate there. After birth,
upon bone formation, hematopoiesis passes from the fetal liver
to the bone marrow. The definitive bone marrow hematopoiesis is
the source of both Ly6C+ and Ly6C− circulating monocytes, from
which resident tissue macrophages were thought to derive (10).
The human embryonic hematopoietic system is organized
roughly in the same way as in the mouse (82), and early studies
propose that macrophages could arise in the embryo independent
of bone marrow progenitors in human beings [for more extensive
reading, see Ref. (83, 84)]. In summary, macrophages in fetal and
adult tissues derive from at least three sources: yolk sac (giving rise
to some tissue-resident yolk sac-derived macrophages), fetal liver

Box 1 A hint on osteoclasts.
Osteoclasts are multinuclear giant cells with a hematopoietic origin, commonly known as bone macrophages. They function in bone resorption and are involved in a normal skeletal development, growth, and modeling, for the maintenance of its integrity throughout life, and for
remodeling through calcium metabolism (65). Moreover, osteoclasts are able to interact with the hematopoietic system and the adaptive
immune system (66). Excessive bone loss mediated by osteoclasts plays a major role in certain pathologic conditions, such as rheumatoid
arthritis (RA) and osteoporosis (67, 68). On the other hand, insufficient bone resorption due to the lack of functional osteoclasts (as in CSF-1R
knock-out mice) leads to excessive bone apposition and osteopetrosis (69).
Osteoclasts really seem a class of macrophages on their own. They are generated from mononuclear phagocyte lineage progenitors in the
bone marrow, and their differentiation from an osteoclast precursor (PreOC) depends on CSF-1 and the engagement of receptor activator
of nuclear factor-κB (RANK) and its ligand (RANKL), a specific osteoclast differentiation factor (70). Recently, it has been shown that also
IL-34 is involved in the osteoclast development (71).
Osteoclasts can differentiate in vitro from a cell population named monocyte-derived multipotential cells (MOMCs), which seem to originate
from circulating CD14+ monocytes (72). In vitro induction of MOMCs from circulating CD14+ monocytes apparently requires their binding
to fibronectin, and exposure to soluble factor(s) derived from peripheral blood CD14dim monocytes (72).
Thus, culture of unfractionated peripheral blood monocytes with M-CSF and RANKL is sufficient to induce their differentiation into
osteoclasts, and it has been assumed that osteoclast precursors are monocytes, although this has not been shown in vivo.
The question arises as to why osteoclasts, unlike other macrophages, have their own lineage of commitment and differentiation. Possibly,
the reason may lie in the fact that they are phylogenically closely linked to the presence of bone, a tissue that develops late as compared to other organs and tissues during embryonic/fetal development, as in fact vertebrates are the most recent phylogenic step in the
evolution (73).

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Italiani and Boraschi

(giving rise to fetal liver-derived macrophages), and bone marrow
(giving rise to tissue-resident bone marrow-derived macrophages
and inflammatory bone marrow-derived macrophages, see below)
(Figure 1). The primitive yolk sac-derived macrophages have
two distinct characteristics: (1) their pattern of differentiation
does not go through a monocytic intermediate state but they
directly become mature macrophages in fetal tissues (85) and
(2) unlike macrophages derived from definitive c-Myb-dependent
hematopoiesis, they are independent of the transcriptional factor
c-Myb during development, while depending on the transcriptional factor PU.1 (12).
Based on different experimental approaches, from lineage tracing (12, 48) to experiments carried out in parabiotic mice (64,
86), it is evident that monocytes do not contribute or contribute only minimally to the maintenance of peripheral tissueresident macrophages in steady-state conditions in many adult
tissues. Fate-mapping experiments have shown that the adult
microglial cell population is exclusively derived from yolk sac
progenitors (87, 88), whereas for LC in adult skin it was clearly
demonstrated a mixed origin, from the yolk sac and from the
fetal liver (12, 89). Moreover, using Myb-deficient mice that
lack development of HSCs, followed by transplantation with
genetically dissimilar bone marrow together with fate mapping,
it has been observed that yolk sac macrophages can generate
macrophages with a characteristically high expression of the F4/80
marker (F4/80 bright macrophages) in brain (microglia), skin
(LC), liver (Kupffer cells), pancreas, and spleen (12). In kidney and lung, tissue-resident macrophages have a double origin,
encompassing F4/80high macrophages, derived from yolk sac, and
F4/80low macrophages, which have a hematopoietic origin and are
continuously replaced by bone marrow-derived progenitors (12).

FIGURE 1 | Origin of tissue-resident macrophages in the mouse. In adult
tissues, macrophages derive from three sources. The first is the yolk sac in
the embryo, where primitive hematopoiesis occurs giving rise to progenitors
that seed tissues with F4/80high macrophages. Later during fetal development,
hematopoiesis shifts from the yolk sac to the fetal liver (that seems to
contribute to the LC pool in the skin, possibly through a yolk sac-derived
progenitor). It is unknown whether other resident macrophages in other
tissues may also derive from fetal liver hematopoiesis. The third source is the
bone marrow, where definitive hematopoiesis occurs in the fetus and in the

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

Moreover, F4/80high shares a common gene signature with yolk sac
macrophages, unlike F4/80low cells, as shown by global transcriptional analysis (12). Also, for splenic red pulp macrophages, alveolar, and peritoneal macrophages, an embryonic origin has been
confirmed, rather than a monocyte origin (48). All these experiments show that early embryonic progenitor-derived macrophages
can persist in tissues to adulthood. As mentioned previously, an
exception is the gut, which contains a large population of resident macrophages that are all blood monocyte-derived cells, in
steady-state conditions (57). How the mutual contribution of yolk
sac-derived macrophages and fetal liver-derived monocytes is regulated in each tissue is unknown, and likewise it is not known
how these two distinct populations of macrophages are functionally and ontogenically related. Regarding how much yolk sac
progenitors contribute to originating adult tissue macrophages vs.
fetal liver hematopoiesis, there are different opinions. One hypothesis is that fetal liver-derived monocytes proliferate and differentiate into adult tissue macrophages markedly diluting the population of yolk sac-derived macrophages (e.g., in lung and heart). This
hypothesis stems from the observation that generation of yolk sacderived macrophages does not go through a monocytic intermediate, therefore being in contrast with normal adult hematopoiesis,
while a fetal liver origin for tissue macrophages would be reminiscent of the adult scenario in inflammation (90). Conversely, others
believe that all tissue macrophages derive from yolk sac during the
embryonic development, and circulating monocytes do not seed
the majority of the adult tissues in mice (except kidney and lung)
(12, 91) (Figure 2). This concept is strengthened by findings in
human beings, where a complete loss of CD16+ monocytes seems
to be of little consequence (92), and many tissue macrophage populations appear to be intact in patients with monocytopenia caused

adult, giving rise to monocytes and to monocyte-derived F4/80low
macrophages. Expression of murine F4/80 (the human EMR1) is an
insufficient marker to discriminate between monocyte-derived macrophages
and tissue-resident macrophages. It seems that Ly6C+ monocytes are the
precursors of tissue macrophages, while the exact contribution of Ly6C−
monocytes remains unclear. HSC, hematopoietic stem cell; GMP,
granulocyte-macrophage progenitor; MDP, macrophage–dendritic cell
progenitor; LC, Langerhans cell; CSF-1, colony-stimulating factor 1; IL-34,
interleukin 34.

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FIGURE 2 | Distribution of tissue-resident macrophages and
monocyte-derived macrophages in tissues and organs. Monocyte
contribution to resident macrophages is highly tissue-dependent and varies
from no contribution for brain microglia and epidermal LC to complete
monocyte origin for intestinal lamina propria macrophages. The tissues
listed in the middle are those at the center of ongoing controversy (see the
main text), and for which a mixed contribution is probable. Here, we define
yolk sac-derived macrophages as tissue-resident macrophages, and both
fetal liver-derived macrophages and bone marrow-derived macrophages as
monocyte-derived macrophages (considering that bone marrow
hematopoiesis derives from fetal liver hematopoiesis). LC, Langerhans
cells; LP, lamina propria; RP, red pulp.

by immune deficiency syndromes (93, 94). In conclusion, to which
extent different populations of yolk sac-derived macrophages may
be later replaced by fetal liver-derived macrophages or monocytes,
and how yolk sac-derived tissue-resident macrophages can proliferate locally through life to maintain their own pool independently
of adult monocyte input, these issues remain a matter of debate
(90, 91, 95). Thus, three main issues arise from all these findings:
1. The origin of adult macrophages in steady-state conditions can
vary considerably between tissues.
2. The exact role of the patrolling Ly6C− monocytes remains
unclear, while Ly6C+ monocytes are recruited predominantly
to sites of infection or injury, or to the organs and tissues that
have continuous cyclic recruitment of macrophages (such as
the uterus), or that are exposed to microbiota (such as the gut
and the skin).
3. Due to some limits and weaknesses of the published studies
(whole blood irradiation or other myelo-ablative treatments,
parabiotic mice, engrafted bone marrow or monocytes, adoptive transfer of radiolabeled cells, Cre-loxP-based fate mapping,
CCR2 or CSF-1 inactivation, etc.) (9, 90), none of such studies
provides conclusive evidence against a role for monocytes in
tissue macrophage homeostasis. Thus, more efficient and specific fate-mapping models of yolk sac-derived macrophages and
fetal liver-derived monocytes are needed, along with further
investigation, to determine which tissue macrophage populations are constantly replenished by circulating monocytes and
which are not.
SELF-RENEWAL/PROLIFERATION CAPACITY OF TISSUE-RESIDENT
MACROPHAGES IN HOMEOSTASIS

Given that it is currently not possible to discriminate the
two populations of tissue macrophages (yolk sac-derived and

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Monocyte to macrophage differentiation

monocyte-derived) during homeostasis, we will report their ability
to proliferate without considering them as distinct subpopulations. In any case, we will bear in mind the notion that the tissue
macrophages can maintain their number in the absence of monocyte precursors both in steady-state conditions (12, 48, 64, 86) as
well as in genetically or experimentally monocytopenic situations
(94, 95).
It is important to clarify the difference between self-renewal
and proliferative capacity. As stated by Sieweke and Allen (13), in
immunology, self-renewal is understood as a replacement of a certain cell population, while in stem cell research as the capacity to
generate with a cell division a daughter cell showing the same identity as the parental cell. Local proliferation of tissue macrophages
can be considered as self-renewal in both senses [see Ref. (12,
13)], since macrophages can proliferate without change of their
differentiated phenotype (96). Having said that, recent evidence
demonstrated that macrophages within the adult tissues self-renew
via proliferation in homeostatic conditions rather than through an
influx of progenitors. This has been shown for LC, which are able
to proliferate (97) both in human beings (98) and in mice (99),
for brain microglia (19), resident peritoneal macrophages (100),
and alveolar macrophages (101). The self-renewal process is regulated by growth factors and cytokines such as CSF-1 and GM-CSF
(Box 2).
Resident macrophages can proliferate at low levels in steadystate conditions, but proliferation rates strongly increase after
macrophage depletion (86) or under inflammatory challenge (13).
Regarding their proliferative ability, Ginhoux and Jung (90) raise
the interesting question as to “whether all macrophages within
a tissue possess equal self-renewal potential, or whether there
are macrophage subpopulations that differ in their capacity of
survival and proliferation, which would imply the existence of
macrophages subpopulations with stem cell-like features.” The
question arises from observations that physiological or experimental depletion of LC leads to a clonal expansion of LC by adjacent
proliferative cell clusters with stem cell-like features (120), and
that among lung macrophages, different cells can proliferate to
maintain the population (87). To solve this issue, there is evidence that a macrophage that had previously divided has the same
probability of entering the cell cycle as a cell that had not, suggesting the same proliferative ability for all macrophages (87).
This is consistent with the observation that macrophages genetically modified to have an indefinite self-renewal potential can be
efficiently cloned (96).
During inflammation, things are quite different, especially because the tissue is enriched with monocyte-derived
macrophages. We will discuss later the replenishment of tissue
macrophages by monocyte-derived macrophages and their ability
to proliferate.
TISSUE MACROPHAGE FUNCTIONS

Table 3 summarizes the functions of resident macrophages in
the main body tissues. These functions, mirroring different phenotypes (74, 143), are specific because depending on different
tissue microenvironments. Different tissues define different phenotypes of both resident macrophages and monocyte-derived
macrophages recruited from the reservoirs of blood, spleen, and

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Box 2 Factors driving monocyte/macrophage self-renewal, proliferation, and functional differentiation.
Macrophage colony-stimulating factor (M-CSF, also known as CSF-1) and granulocyte-macrophage colony-stimulating factor (GM-CSF) drive
the monocyte/macrophage development, differentiation, and proliferation along with cytokines such as IL-4 (102) and the recently discovered IL-34 (103). Macrophages and circulating monocytes express the CSF-1 receptor (CSF-1R) (42, 43), and mouse deficient in CSF-1R
(osteopetropic mice, op/op) have a decreased number of monocytes in the bone marrow and in circulation, in addition to a decrease in
osteoclasts (69, 104). Experimentally blocking CSF-1R with antibodies leads to a reduction in Ly6C− monocytes (69) and to an associated
increase in Ly6C+ monocytes, suggesting the involvement of CSF-1 in the maturation of monocytes from Ly6C+ to Ly6C− (48, 104). CSF-1 is
constitutively produced by mesenchymal cells (105) and is detectable in circulation in resting conditions (20). Under homeostatic conditions
(106), CSF-1 promotes monocyte development and macrophage proliferation (107), which is controlled in a negative feedback loop. In fact,
mature mononuclear phagocytes express high level of CSF-1R and are responsible for the clearance of CSF-1. The decreased CSF-1 levels
lead to a decrease in mononuclear cell proliferation, thereby maintaining the cell number to normal levels both systemically and locally [(21,
22); a model of CSF-1-dependent local homeostasis of macrophage density has been described by Jenkins and Hume (9)]. Thus, elevated
production of CSF-1 can drive both an increased proliferation of resident macrophages and an increased recruitment of monocytes (103, 108)
via macrophage production of CCL2 (109). CSF-1 deficiency in mice affects distinct tissues by different degrees, ranging from marked cell
loss in the gut, kidney, peritoneal cavity, and in circulation, as compared to liver (86). CSF-1 is also involved in the proliferation of splenic red
pulp macrophages and bone marrow macrophages (110). GM-CSF is also critical for macrophage homeostasis and proliferation, especially
in the lung (111) and in the peritoneal cavity in vivo (112), but it is less important in hematopoiesis, and, therefore, for monocyte development
(113). GM-CSF can support monocyte expansion and differentiation in vitro (25, 114), and it seems to be mainly involved in induction of
hematopoiesis during inflammation rather than in homeostasis (115, 116). CSF-1 and GM-CSF are also involved in monocyte/macrophage
functional differentiation programs: CSF-1 stimulation leads to a homeostatic or anti-inflammatory M2-like phenotype (25, 117, 118), whereas
GM-CSF leads to an M1-like inflammatory phenotype (25, 117–119). Thus, CSF-1 stimulation represents a default homeostatic/M2 pathway
of monocyte development (119). In summary, CSF-1 is mainly involved in self-renewal of tissue macrophages, consistent with its role in
M2 polarization, while GM-CSF is involved in proliferation of monocyte-derived inflammatory macrophages, consistent with its role in M1
polarization.

Table 3 | Macrophage functions and the pathological consequences of their anomalous activation in the main tissues.
Macrophages (MΦ)

Tissue

Microglia

Brain

Functions

Pathology

Brian development (121), immune surveillance,

Neurodegeneration (123)

synaptic remodeling (122)
Osteoclasts

Bone

Bone modeling and remodeling, bone resorption

Osteoporosis, osteopetrosis, arthritis (126)

(124), support to hematopoiesis (125)
Heart MΦ

Heart and vasculature Surveillance

Atherosclerosis (127)

Kupffer cells

Liver

Toxin removal, lipid metabolism, iron recycling,

Fibrosis (130), impaired erythrocyte

erythrocyte clearance, clearance of microbes, and

clearance (131)

cell debris from blood (128, 129)
Alveolar MΦ

Lung

Surfactant clearance, surveillance for inhaled

Alveolar proteinosis (133)

pathogens (132)
Adipose tissue-associated

Adipose tissue

MΦ

Metabolism, adipogenesis, adaptive

Obesity, diabetes, insulin resistance, loss

thermogenesis (134)

of adaptive thermogenesis (131)

Bone marrow MΦ

Bone marrow

Reservoir of monocytes, waste disposal (131)

Disruption of hematopoiesis (131)

Intestinal MΦ

Gut

Tolerance to microbiota, defense against pathogens,

Inflammatory bowel disease (136)

intestinal homeostasis (135)
Langerhans cells

Skin

Marginal zone MΦ, red

Spleen

pulp MΦ
Inflammatory MΦa

Healing

MΦb

All tissues

All tissues

Immune surveillance (137)

Insufficient healing, fibrosis (138)

Erythrocyte clearance, iron processing, capture of

Impaired iron recycling and erythrocyte

microbes from blood (139)

clearance (140)

Defense against pathogens, protection against

Chronic inflammation, tissue damage,

dangerous stimuli (141)

autoimmunity (91)

Branched morphology, angiogenesis (142)

Cancer, fibrosis, epithelial hyperplasia (91)

a

Also known as inflammatory macrophages or M1 macrophages.

b

Also known as deactivated or M2 macrophages.

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bone marrow (10), phenotypes that are necessary for the tissuespecific needs of defending, maintaining, and regaining homeostasis (144). These homeostatic functions may be altered by
chronic insults, which may lead to an anomalous prolongation/amplification of the macrophage attempt to regain homeostasis and to a consequent causal association between macrophages
and diseases (Table 3). In pathological conditions, the distinction
between tissue-resident macrophages and recruited inflammatory macrophages has not yet been possible. For an in-depth
analysis of these issues, the reader can refer to recent exhaustive
reviews (99, 144, 145). Apart from tissue-specific functions, tissue
macrophages share a series of common functions encompassing clearance of cell debris, immune surveillance, wound healing,
defense against pathogens, and the initiation and resolution of
inflammation. In this review, we will only focus on the role of
macrophages in inflammatory responses, considering their capacity to polarize into different functional phenotypes in response
to the tissue microenvironmental changes that occur during the
different phases of an inflammatory response. This polarization
process is based on the M1–M2 paradigm (see below).

PLASTICITY OF MONOCYTES/MACROPHAGES DURING
INFLAMMATORY REACTION
MONOCYTE RECRUITMENT DURING THE INFLAMMATORY REACTION:
INFLAMMATORY MONOCYTE-DERIVED MACROPHAGES VS.
TISSUE-RESIDENT MACROPHAGES AND THEIR PROLIFERATIVE ABILITY

During the first phases of an inflammatory reaction, there is
in the tissue an increase of the effector cell number, necessary
for increasing the immune defensive firepower. These cells are

Monocyte to macrophage differentiation

monocyte-derived macrophages. The concomitant drastic loss of
resident macrophages, due to tissue adherence, emigration, or
death, is a phenomenon termed “the macrophages disappearance
reaction” (146), and it is especially evident for peritoneal and alveolar macrophages. To cope with the need of increasing the number
of effector cells, two strategies come into play.
First is the recruitment of blood monocytes, driven by resident
macrophages alongside with other tissue cells. Recruited blood
monocytes are a source of inflammatory macrophages, which
take the name of bone marrow-derived or monocyte-derived
inflammatory macrophages. The other strategy is the increase of
tissue-resident macrophage proliferation by enhancement of their
self-renewal ability.
Central to the issue of monocyte recruitment is the difference in
monocyte subset trafficking. Such differences have been observed
to occur during acute and chronic inflammation in mice, and
underline the fact that the monocyte subsets are under the control of distinct trafficking mechanisms, with the classical subset
being recruited via CCR2 and the non-classical one utilizing a
CXCR1-dependent pathway (see Box 3).
In a model of Listeria monocytogenes infection, non-classical
monocytes (Ly6C− ) extravasate rapidly within 1 h, invade the surrounding tissues, and develop a very early inflammatory response
by producing chemokines responsible for recruiting other effectors cells (granulocytes, NK cells, T-cells), and cytokines such
as TNF-α (central to macrophage-mediated inflammation and
innate responses) (45). This inflammatory response is transient,
and 8 h after infection, the main producers of inflammatory
cytokines in the tissue are the classical monocytes (Ly6C+ ). As

Box 3 Chemokines and monocyte recruitment mechanisms.
The two main chemokines and related receptors involved in the inflammation-dependent recruitment of the Ly6C+ and Ly6C− monocyte
subsets from the blood, bone marrow, and spleen, are CCL2/CCR2 and CX3CL1/CX3CR1, respectively (45, 46). Fibroblasts, epithelial, and
endothelial cells produce CCL2 in response to inflammatory cytokines or microbial molecules, and generate a high level of this chemokine
in the inflamed tissue (to allow egress of monocytes from the blood and entry in the tissues) and/or in blood (to allow entry of bone marrow
monocytes) (44, 147). Recently, it has been proposed that both mesenchymal cells and progenitor cells closely apposed to bone marrow
vessels can produce CCL2 in inflammatory situations, to allowing the egress of monocytes from the tissue and their subsequent entry
into the blood (148). During a bacterial infection, Ly6C+ monocytes require CCR2 for being recruited from the bone marrow into the blood
(149). In mice lacking CX3CR1, a reduction of patrolling by Ly6C− was observed (45), and a reduction of their number in infracted heart
(150), suggesting an impaired recruitment from the blood. Genetic destruction of CCR2 reduces the accumulation of both Ly6C+ and Ly6C−
monocytes in injured skeletal muscle, but it does not alter the recruitment of Ly6C− monocytes in the heart after myocardial infarction
(150). A reduction was also observed in skin wounds on the first day from injury, when Ly6C+ cells are those principally involved in the early
repair phases, but not during the late stage of tissue repair, when Ly6C− cells are dominant (151). These studies underline the importance
of monocyte recruitment from blood to the tissue in the injured cardiac or skeletal muscle. Regarding the role of in situ differentiation,
in addition to the data mentioned above (150), a reduction of Ly6C− monocytes has been observed also in the blood of CCR2-deficient
mice, despite the fact that they do not express this receptor (48). Macrophage accumulation in skin wounds is also reduced in mice lacking
CX3CR1 (152). CX3CL1 and CX3CR1 provide a survival or anti-apoptotic signal to Ly6C− cells (153). Two models have been proposed for
the CCL2-dependent Ly6C+ cell recruitment from the bone marrow: CCL2 increases monocyte chemokinesis and contact with blood vessels; CCL2 associates with tissue glycosaminoglycans and forms a gradient driving monocytes to exit the bone marrow for entering into
circulation (54). Intravenous administration of CCL2 leads to the mobilization of monocytes into the circulation, which is consistent with a
role for peripheral CCL2 production responsible for replenishment of circulating monocytes from bone marrow (154).
Under steady-state conditions, the release of Ly6C+ and Ly6C− monocytes from bone marrow depends on two genes, the circadian clock
gene Bmal1 for Ly6C+ cells (55), and the G-coupled receptor for sphingosine-1-phosphate S1PR5 for Ly6C− monocytes (155). In mice with
myeloid cell-restricted Bmal 1-deficiency, the rhythmic release of CCL2 was ablated along with monocyte pools. Thus, myeloid cells produce
low diurnal levels of CCL2 in a circadian fashion, and CCL2 in turn stimulates the release of CCR2-expressing monocytes from the bone
marrow into the blood. On the other hand, in S1PR5-deficient mice, Ly6C− monocytes are retained in the bone marrow and are not released
in the blood and spleen.

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Italiani and Boraschi

previously mentioned (45), it has been observed that the two
subsets of monocytes differentiate into two distinct cells types.
Ly6C− patrolling monocytes initiate a macrophage differentiation
program that resembles that of M2 macrophages (see below), while
Ly6C+ monocytes differentiate into DC-like cells that resemble
Tip-DC (45). However, in other systems, this double recruitment of different monocyte subsets has not been observed. Only
Ly6C+ monocytes were observed to migrate to the injured tissue in a model of skeletal muscle injury and be responsible for
early inflammatory responses (156). Generally, classical monocytes infiltrate inflamed tissues more robustly than their nonclassical counterparts, and their number is significantly increased
in the circulation during systemic or chronic infection (27). After
engulfing dying cells in the tissue, the recruited classical monocytes differentiate into cells that resemble Ly6C− monocytes, and
become involved in tissue repair mechanisms (156). Likewise, in
a mouse model of sterile wound (subcutaneous polyvinyl alcohol sponge implantation), it has been recently demonstrated that
Ly6C+ monocytes recruited from the circulation into the skin
acquired an inflammatory function and, despite time of maturation was long, they matured into Ly6C− macrophages with
repair functions (157). Yet, another situation is that of myocardial infarction, during which both monocyte subsets appear to
home to the same tissue at different stages of inflammation
(150). Specifically, the Ly6C+ subset first infiltrates the infracted
heart and exhibits inflammatory functions, while the Ly6C− subset is recruited at a later stage and promotes tissue healing by
expressing high amounts of vascular endothelial growth factor, exhibiting angiogenic capacity, and promoting deposition of
collagen (150).
In atherosclerosis, as a model of chronic inflammation, both
monocyte subsets are recruited at the same time to the activated endothelium/plaques, and healing seems to be correlated
with a reduction in total monocyte recruitment (158). However,
it was recently demonstrated that the maintenance and accumulation of monocyte-derived macrophages in atherosclerotic plaques
mainly depend on local proliferation of bone marrow-derived
macrophages rather than on the influx of circulating monocytes
(127, 159). In an atopic dermatitis model and in experimental
autoimmune encephalomyelitis, a massive proliferation of LC and
microglia cells has been observed (160, 161), despite a significant monocyte influx (161). In the peritoneal cavity and in the
lung, where the macrophage disappearance phenomenon occurs
upon bacterial and virus insults, the few remaining macrophages
are responsible for repopulating the tissue (86, 101, 106). Similarly, in the context of Th2-mediated immunity against nematode infection, IL-4 drives tissue-resident macrophage expansion
in the pleural cavity in the absence of peripheral monocyte
recruitment (102).
Proliferation of macrophages is observed in a variety of human
diseases [see Ref. (91)], including tumor-associated macrophages
in solid tumors (162), and adipose tissue-associated macrophages
in obesity (163).
In this context, a question is still open. Having established
that monocytes are recruited into tissues during an inflammatory event, to what extent are they capable to differentiate in
tissue macrophages and to proliferate? As proposed by Jenkins

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Monocyte to macrophage differentiation

and Hume, the negligible contribution of monocytes to the pool
of resident macrophages could be due to the fact that monocyte recruitment is specifically aiming at providing a population
of functionally differentiated cells needed for resolving an acute
inflammatory event, rather than being triggered by the homeostatic need of maintaining the autonomous pool of resident
macrophages (9). This view is supported by another interesting
hypothesis, i.e., in inflammatory conditions, monocyte-derived
macrophages are mostly end-type killer cells, as the non-specific
toxic molecules they produce will also cause their own death (164).
The gastrointestinal tract provides evidence in favor of this
hypothesis. In the gut, blood monocytes are constantly recruited
to the tissue where they contribute to maintaining the resident macrophage population, but during an inflammatory event
they re-program their differentiation plan toward adopting an
inflammatory phenotype (57, 165).
Thus, we should consider that monocyte-derived macrophages
adopt different and opposing phenotypes based on microenvironmental signals. Adoption of a phenotype or another depends
on the time by which the sequential waves of recruited Ly6C+
monocytes reach the tissue during the course of the inflammatory reaction, since the incoming monocytes will find a different
microenvironment in different phases of the reaction. In this context, it is conceivable that monocytes entering the tissue at later
times could find conditions favorable to adopting an M2-like phenotype (see above), thereby becoming tissue macrophages over
time.
Inflammatory monocyte-derived macrophages (12, 86) and tissue monocytes (64) can be phenotypically and functionally distinguished from resident macrophages in many tissues. In the central
nervous system, inflammatory monocyte-derived macrophages
do not contribute to the resident population (161). In contrast, fate-mapping experiments revealed that monocyte-derived
macrophages recruited to the peritoneal cavity upon thioglycollate injection differentiate into resident macrophages and persist
over time (48). The fraction of monocyte-derived macrophages
that do not die upon inflammation and become tissue-resident
macrophages share gene profiling with resident macrophages (45,
64, 165), but there is no information as to whether they are
functionally different or not.
The accumulation of inflammatory monocytes in an inflamed
tissue is due to their influx from blood rather than by their proliferative ability, and in fact inflammatory signals of microbial origin
generally prevent their proliferation. An exception to this general
paradigm comes from a recent study that has demonstrated that
also inflammatory monocyte-derived macrophages can proliferate at certain stages during the resolution of zymosan-induced
peritonitis (106).
All these findings are summarized in Figure 3.
Finally, two issues should be reminded:
1. The precise nature and extent of the contribution of monocytederived macrophages to tissue macrophages could depend on
how, and to which extent, inflammation or its cause has affected
the tissue-resident macrophages. In this view, as proposed by
Ginhoux and Jung (90), tissue-resident macrophages are more
involved in tissue macrophage repopulation after mild injury,

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Italiani and Boraschi

FIGURE 3 | Schematic representation of monocyte and macrophage
populations in homeostasis and inflammation. Under homeostatic
conditions (left panel), Ly6C+ monocytes derive from the bone marrow and
circulate via the blood into the tissue. A minor fraction of these cells lose
Ly6C expression and become Ly6C− monocytes in the blood or in the bone
marrow where some of them might return in the absence of inflammation.
Ly6C+ blood monocytes enter tissues and become either macrophages, for
example, in the gut, lung, and dermis (monocyte-derived macrophages or
monocyte-derived tissue-resident macrophages). Some tissue
macrophages derive directly from yolk sac during the embryogenesis (e.g.,
LC, microglia, liver Kupffer cells, and alveolar macrophages), are long lived,
and are mainly maintained by self-renewal (tissue-resident macrophages).
Ly6C− monocytes act as resident macrophages of the vasculature,
patrolling, and monitoring the endothelial surface in the blood vessel lumen.
In the figure, the presence of “trained” macrophages is also considered,
which we define as “memory macrophages,” i.e., the tissue macrophages
that retain the memory of a previous inflammation and are in a quiescent
state in the tissue. During an inflammatory reaction (right panel), the
number of blood Ly6C+ monocytes recruited to an inflamed tissue increases
considerably. The large majority of these cells gives rise to the inflammatory
monocyte-derived macrophages, while some of them do not differentiate

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Monocyte to macrophage differentiation

into macrophages and remain monocyte-like cells, are able to take up
antigens, and to migrate to the draining lymph nodes (tissue monocytes).
These are the antigen-uptaking and -presenting cells of the tissue. During
inflammation, all macrophages (tissue-resident macrophages,
monocyte-derived tissue macrophages, inflammatory monocyte-derived
macrophages) are activated and differentiate into M1-like inflammatory cells
following interaction with pathogenic and damaged signals/insults in
surrounding microenvironment. These cells produce a series of cytokines
and other inflammatory factors. Tissue-resident macrophages increase their
capacity of proliferation to compensate the loss of macrophages caused by
the inflammatory reaction. Recent evidence demonstrates that also
inflammatory monocyte-derived macrophages are able to proliferate in a late
phase of the inflammatory reaction. Memory macrophages are important
players in the inflammatory reaction, as they can react to inflammatory
stimuli with a faster and stronger inflammatory cytokine production. The role
of circulating Ly6C− cells during an inflammatory reaction is not fully
identified. They probably remain in the blood vessels as sentinels, and in
some cases they could enter in the tissue, as it has been reported in the
case of myocardial infarction, to take up a repair function. HSC,
hematopoietic stem cell; cMoP, common monocyte progenitor; Ly6C,
lymphocyte antigen 6 complex; LC, Langerhans cells; MΦ, macrophages.

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while monocyte-derived macrophages are more involved in
severe inflammatory injuries.
2. In the mouse it seems that non-classical monocytes contribute
to the resident macrophage population. It is possible (although
there is little evidence in this respect) that when non-classical
monocytes are recruited in the inflamed tissue, they may differentiate into alternatively activated macrophages, while classical
monocytes would give rise to classically activated macrophages.
In this context, the developmental relationship between the
different monocyte subsets and the different macrophage functional phenotypes has yet to be fully and formally proven. No
evidence in this sense has been generated yet in human being
(see below).
MACROPHAGE DIFFERENTIATION AND FUNCTIONAL PHENOTYPES

Macrophage polarization occurs through different activation programs by which macrophages carry out their defensive functions.
In this way, macrophages become able to respond with appropriate functions in distinct contexts, functional diversity becoming the key feature of these cells. Essentially, macrophages can
modify their metabolic functions from a heal/growth promoting
setting (M2 macrophages), to a killing/inhibitory capacity (M1
macrophages) (145, 166). The main difference between these cells
is that in M2 macrophages the arginine metabolism is shifted to
ornithine and polyamines, while in M1 cells it is shifted to NO
and citrulline (166). M2-produced ornithine can promote cell
proliferation and repair through polyamine and collagen synthesis, fibrosis and other tissue remodeling functions (167), while
M1-produced NO is an important effector molecule with microbicidal activity and cell proliferation inhibitory capacity (168).
Interestingly, polyamine production per se has been reported to
be a driver of M2 polarization (169), and M2 is the normal
“default” program adopted by resident macrophages (170). Moreover, M1 and M2 macrophages have distinct features in terms
of chemokine production profiles (171), and iron and glucose
metabolism (172, 173).
The description of macrophages polarization is leading immunologists to take a step back and revise their concept on how
the immune system works (14, 145). The M1 and M2 definition was formulated by mirroring the Th1/Th2 polarization
concept. However, this definition might be misleading in that
it may suggest that Th1/Th2 cells do instruct M1/M2 polarization, whereas it is now obvious that the reverse is true, i.e.,
macrophages are initiating and directing T-cell polarization. Since
it has been shown that distinct populations of macrophages promote and control CD4+ T-cell-dependent type 1 and type 2
immune inflammatory responses (i.e., those against viruses and
bacteria, and against multicellular parasites, respectively), not
surprisingly they have been termed M1 and M2 (166). Thus,
considering that macrophages recognize pathogens directly (174,
175), while T-cell do not, and considering that T-cells proliferate through interaction with macrophages (176), it is logical
to think that macrophages are the cells that initiate and direct
T-cell response, and that the adaptive immune response needs
the triggering and guidance of innate immunity (14). Notably,
M1 and M2 macrophage activities do not need the presence of
lymphocytes (145). The fact that T-cell-derived cytokines such

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Monocyte to macrophage differentiation

as IFN-γ and IL-4 may amplify macrophage polarization (see
below) should not deceive us into believing that macrophage
polarization cannot occur without lymphocytes, as this is not
the case. In vitro, macrophages are activated toward an M1 functional program by infectious microorganism-related molecules
(e.g., the gram-negative product LPS) and by inflammationrelated cytokines TNF-α or IFN-γ, alone or in combination. M1
macrophages are characterized in vitro by an IL-12hi IL-23hi IL10lo phenotype; are efficient producers of toxic effector molecules
(ROS and NO) and inflammatory cytokines (IL-1β, TNF, IL6); participate as inducers and effector cells in polarized Th1
responses; and mediate resistance against intracellular parasites
and tumors (177). Conversely, M2-like polarization has been
observed in vitro in response to the Th2-related cytokines IL-4
or IL-13 (178), to the concomitant triggering of Fcγ receptors
and Toll-like receptors (TLR), to immune complexes, and to
anti-inflammatory molecules such as IL-10, TGF-β, and glucocorticoids (115). The variety of functional programs adopted by
macrophages in response to the stimuli listed above has been
termed M2a (IL-4 and IL-13; alternative inflammation), M2b
(immune complexes, FcγR/TLR triggering), and M2c (IL-10, TGFβ, glucocorticoids; deactivation) (178, 179). M2 cells are characterized in vitro by an IL-12lo IL-23lo IL-10hi TGF-βhi phenotype and
generally have high levels of scavenger, mannose, and galactosetype receptors. In general, these macrophages take part in polarized
Th2 responses, allergy, parasites clearance, dampening of inflammation, tissue remodeling, angiogenesis, immunoregulation, and
tumor promotion (180).
Macrophage taxonomy is an attempt to rationally categorize an
extended variety of cell functions. Indeed, the M1/M2 paradigm
is a limited attempt to define the complexity and plasticity of
mononuclear phagocytes. In vivo, macrophages can adopt a variety of functional phenotypes depending on subtle and continuous
changes in the tissue microenvironment. So, the M1/M2 polarization of macrophage functions may be taken as a simplified conceptual framework describing a continuum of diverse functional
states, of which M1 and M2 activation states are not ontogenically defined subsets but represent the extremes (180–182). In this
regard, Mosser and Edwards (181) have suggested a macrophage
classification that takes into account the three functions of these
cells in maintaining homeostasis: host defense, wound healing,
and immune regulation. Classifying macrophages according to
these functions provides three basic macrophage populations:
classically activated macrophages, wound-healing macrophages,
and regulatory macrophages (183, 184). The authors believe that
this classification also helps to illustrate how macrophages can
evolve to exhibit characteristics that are shared by more than one
macrophage population (181).
Without going into details [for which we refer the reader
to other reviews; (14, 185)], the M1/M2 classification in different subsets can create the illusory perception of a heterogeneity,
which is not proven to exist in vivo. Thus, as already mentioned
above, it is logical to hypothesize that the subsets are varying mixtures of M1- and M2-type macrophages, as observed in the lung
and in the peritoneal cavity, where tissue-specific variations in
the balance of M1- and M2-type responses have been revealed
(74, 186). This situation has also been observed in pathological

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conditions, where macrophages can develop mixed M1 and M2
phenotypes (187, 188). Moreover, it has also been proposed to consider the heterogeneity of macrophage functions as a consequence
of interaction with different immunological pathways (e.g., interaction with different growth and survival factors, interaction with
lymphoid and myeloid cytokines, interaction with pathogens, resolution), rather than attributing them to distinct macrophages
subsets (185).
In summary, the initial inflammatory response is carried out
by activated macrophages in classical or alternative modality
(depending on the triggering events), aiming at eliminating invading microbes by promoting the inflammatory response. Then, the
resolution phase is carried out by macrophages in deactivated
modality, unresponsive to inflammatory stimuli, and active in the
elimination of the injured cells and tissue components, in promoting angiogenesis, cell proliferation, matrix deposition, and in
general in tissue remodeling. The mechanisms that account for
macrophage deactivation play a key role in maintaining homeostasis and keeping the immune response under control (189).
Both innate and adaptive signals can influence the macrophage
functional phenotype, which can have potentially dangerous consequences if not appropriately regulated. For example, classically
activated M1 macrophages can cause damage to host tissues,
predispose surrounding tissue to neoplastic transformation and
influence glucose metabolism by promoting insulin resistance.
Macrophages that are normally involved in wound healing can
promote fibrosis, exacerbate allergic responses, and be exploited by
pathogens for intracellular survival. These M2-type macrophages
can contribute to the progression of neoplasia by promoting tumor
survival (see Table 3).
CURRENT HYPOTHESIS ON MACROPHAGE POLARIZATION

Plasticity and flexibility are key features of macrophages and of
their activation states. A controversial issue is whether a phenotypic and functional evolution of macrophages occurs in vivo, and
how it happens. As mentioned above, it has been observed in mice
that the M1 to M2 switch during the progression of the inflammatory response enables macrophages to perform different activities
in the different phases of the reaction. The controversy refers to
the mechanisms underlying this switch, i.e., whether M1 and M2
macrophages are phenotypically distinct subpopulations that can
serve different functions in different phases of an inflammatory
reaction (45, 150), or the same cells can shift from one to another
functional phenotype in response to microenvironmental signals
(156, 157).
Several hypotheses are attempting to explain the issue. A first
hypothesis is that different subsets of monocytes or macrophages
can adopt a different functional phenotype. Thus, Ly6C+ monocytes and/or monocyte-derived macrophages in the tissue become
M1 macrophages, and Ly6C− monocytes and/or tissue-resident
macrophages become M2 macrophages. It is possible that resident macrophages maintain cytoprotective and reparative functions, whereas macrophages derived from circulating inflammatory monocytes perform mainly M1 type functions. This hypothesis is not fully supported by the studies previously cited, where in
different situations it was possible to observe both the differentiation of Ly6C+ cells in M1 and of Ly6C− cells in M2 (45, 150) and

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Monocyte to macrophage differentiation

the transdifferentiation from Ly6C+ M1 cells to Ly6C− M2 cells
(156, 157).
A second hypothesis is that there are sequential waves of
monocyte recruitment into a tissue throughout the course of
an inflammatory reaction. Therefore, monocytes recruited into
the tissue at different times encounter different microenvironments with different signals that can polarize them in M1 during early phases and in M2 in late phases (156). In this case,
cytokines and other microenvironmental signals in the tissue play
a key role in determining the different functional phenotypes
of macrophages. Although the role of cytokines in steering the
macrophage functional phenotypes has been proven in vitro (179),
the situation could be very different in vivo, where M2 activity
is strongly increased in sterile wounds (157) or injured kidney
(190) in absence of Th2-like cytokines IL-4 or IL-13 (which in
any case do not induce the typical M2 phenotype, i.e., the deactivated healing/repairing functional phenotype). In these cases, M2
macrophages derive largely from M1 macrophages, with monocytes recruited from the circulation first acquiring an inflammatory phenotype, and then persisting in the tissue and maturing
into repair macrophages.
Based on the latter data, a third hypothesis is that polarized
macrophage populations can switch one to the other in response
to different conditions. Data from in vitro studies demonstrate
that human monocytes can acquire the phenotype of polarized
M1 macrophages and then mature into M2 repair macrophages
upon exposure in culture to sequential changes in the microenvironmental conditions (191). Other studies demonstrated that
M2 macrophages are reprogramed to express M1 genes following
exposure to TLR ligands or IFN-γ (192, 193).
A related question is whether both tissue-resident macrophages
and monocyte-derived macrophages can polarize in M1 or/and
M2 functional phenotypes. We have described above that tissue
macrophages have basically an M2-like phenotype, whereas infiltrating recruited monocytes differentiate in M1 or M2 depending
on the tissue conditions. For instance, it has been shown that
tissue-resident macrophages, rather than recruited monocytes, are
alternatively activated in the tissue during infection with Litomosoides sigmodondis (102). Also, recruited monocytes can be
directly polarized into an anti-inflammatory M2 phenotype by
basophil-derived IL-4, in order to alleviate allergic inflammation in the skin (194). Although it is not possible discriminating between tissue-resident and monocyte-derived macrophages
in steady-state conditions, it seems that alternatively activated
tissue macrophages have a transcriptional profile and phenotype different from that of alternatively activated monocytederived macrophages, with the latter having immunoregulatory
properties (195).
It should be considered that in vitro studies do not fully
recapitulate in vivo differentiation for two main reasons:
1. These studies are generally based on an heterogeneous population of monocytes, encompassing all the blood subsets (Ly6C+
and Ly6C− in the mouse, and CD14+ and CD16+ in human
being), thus it cannot be defined whether upon different stimuli
the same cells can pass from a phenotype to another or whether
different subsets are activated in response to different stimuli.

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2. While M2 macrophages can convert to the M1 phenotype, the
reverse generally does not occur, or it may only occur in particular conditions (e.g., in very mild inflammatory responses).
In fact, M1 is probably an end-stage killer cell that dies during
the inflammatory response, possibly succumbing to its own
NO production, as it was demonstrated in vitro (196). So, their
selective death may give the impression that they convert in
M2 cells, which in fact proportionally increase (145). It seems
that M1 vs. M2 polarization correlates with the capacity of
macrophages to produce NO (166) as opposed to the important
M2 driver TGF-β (164, 197, 198), thus the decrease in NOproducing macrophages would increase TGF-β production and
amplify M2 polarization.
There are cases in which a phenotypic switch in the macrophage
population occurs over time, often associated with pathology (91,
141). Three specific examples of this phenotypic switch are the
following:
endotoxin tolerance, an altered state of responsiveness to secondary stimulation with LPS, resulting in a global and sustained
switch of the gene expression program from an inflammatory
M1 signature to an anti-inflammatory phenotype (199);
obesity-induced insulin resistance or type 2 diabetes, and atherosclerosis lesions. These are metabolic syndromes that can lead
to a switch in the phenotype of adipose tissue macrophages
from wound healing (as in healthy non-obese human beings)
to classically activated macrophages (200, 201);
cancer, where the tumor-infiltrating classically activated
macrophages have the potential to contribute to the earliest stages
of neoplasia (202–204), and then, as the tumor progresses, can
progressively differentiate to a regulatory phenotype and eventually become cells that share the characteristics of both regulatory
and wound-healing macrophages (181).
Although the pathology provides the proof-of-principle that
macrophages can undergo dynamic transitions between different functional states, it is possible that a mixture of M1/M2
phenotypes underlies these conditions (14, 145, 166). In the
past few years, gene expression profiling techniques and genetic
approaches have been used to cast some light on the plasticity of macrophage activation. The commonly held view is
that macrophage polarization is driven by cues in the tissue
microenvironment, which can include cytokines, growth factors, and microorganism-associated molecular patterns. These
signals are thought to dictate a transcriptional response that
shapes the phenotype and function of macrophages based on the
physiological or pathological context. Progress has been made
in defining the molecular mechanism underlying macrophage
polarization, including signaling pathways, miRNA, epigenetic
modification, post-transcriptional regulators, and transcriptional
factors (189, 205–207). However, the data are still incomplete
and far from being systematic, and our knowledge of the
mechanistic basis of macrophage diversity in different tissues
or in response to changing environment is to a large extent
unknown.

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Monocyte to macrophage differentiation

POST-INFLAMMATION FATE OF
MONOCYTES/MACROPHAGES
ANTIGEN PRESENTATION IN NON-LYMPHOID ORGANS

The capacity of taking up and presenting antigen (i.e., the linking function between innate and adaptive immunity) is one of
the most important features of tissue macrophages (208). It has
been mentioned above that some monocytes that enter the tissue
during inflammation do not differentiate into macrophages, and
are able to take up antigen in the tissue and carry it to lymph
nodes where they can present it to naïve T-cells (64). In addition to this population of monocyte-like cells, tissue macrophages
are also able to present antigen, despite the fact that they do
not recirculate to lymph nodes after antigen uptake. That tissue
macrophages are highly phagocytic and can take up microorganisms and other matter in the tissue is well known, as this is their
major function both in homeostasis and during inflammation.
That antigen presentation may occur also in non-lymphoid organs
has been suggested by several experimental evidence describing
antigen-specific local activation and expansion of primed T-cells,
but not of naïve T-cells (209–215). Based on this evidence, the
hypothesis proposed by Ley is that initial priming of naïve Tcells occurs in the lymph node (to which antigen-loaded tissue
monocytes recirculate), but that the full activation and effector
functions of T-cells occur in the tissue where the inflammatory
reaction is taking place, upon the productive interaction and formation of immunological synapse between primed T-cells and the
antigen-presenting tissue macrophages (the difference between
monocyte-derived tissue DC and tissue macrophages is bleared,
as they seem to be not much more than slightly different functional differentiation states from a common precursor). Most
likely, the inflammatory monocyte-derived cells with an M1-like
functional phenotype are the antigen-presenting cells (APC) that
induce activation/polarization of effector Th1 and Th17 cells upon
production of IL-12 and IL-23, respectively, and in a TNFRSF
and TNFSF-dependent fashion (but independent of CD80, CD86,
and CD28 co-stimulation). Likewise, M2-like tissue macrophages,
which produce TGF-β and express the αVβ8 integrin are likely
involved in the polarization of iTreg cells, whereas their role in
Th2 polarization is less clear (208).
FATE OF ACTIVATED RESIDENT MACROPHAGES AND RECRUITED
MONOCYTES: PROLIFERATION, REPLACEMENT, AND M2-LIKE
POLARIZATION

Based on what described above, the cell populations present in the
tissue during the acute phase of an inflammatory reaction are the
following:
• Tissue-resident macrophages and monocyte-derived macrophages.
These, after initial recognition of microbial or damageassociated molecules, drive the influx of blood-derived monocytes, which will become inflammatory macrophages. Their role
in initiating the inflammatory reaction possibly depends on the
nature and grade of challenge.
• Monocyte-derived macrophages, newly recruited and rapidly
occupying the inflammatory lesion, becoming the majority of
the macrophages present in the tissue. These cells induce the

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Monocyte to macrophage differentiation

inflammatory response by differentiating in the M1 functional
phenotype.
• Tissue monocytes, the recently described cells that can take up
antigens in the tissue and move to lymph nodes, where they are
able to present antigens to naïve T-cells.
• Memory macrophages, or trained monocytes, cells functionally
programed by a previously stimulus for either enhanced (training) or decreased (tolerance) cytokine production, depending
on the type and concentration of the stimulus they encountered [(216); see below]. Here, we consider them as a kind
of resident inflammatory monocyte-derived macrophage, able
to react in a faster and stronger manner compared to other
macrophages.
A summary of the different macrophage types and of their fate
after the acute inflammatory phase is given in Figure 4.
In general, tissue-resident macrophages are maintained locally
by proliferative self-renewal (100, 106), and retain an M2-like phenotype, for example, in the peritoneal cavity, brain, and lung (86,
100, 161). The fate of monocyte-derived resident macrophages is
hard to follow, considering that it is not possible to fully discriminate between them. However, we may hypothesize that they have
the same fate of tissue-resident macrophages, i.e., they maintain
an M2-like phenotype and a low self-renewal capacity. A number
of cells of both populations probably die during inflammation,
the extent of their survival possibly depending on the nature and
magnitude of the insult.
Generally, the inflammatory monocyte-derived macrophages
are polarized toward M1, and the majority of them dies, killed
by their own NO production (see above). In an experimental acute lung injury model, these cells undergo Fas-mediated
death, while the resident alveolar cells persist (217). From that,
we can argue that M1 likely is a terminal differentiation phenotype. However, there are reports that they can also undergo in situ
phenotype conversation to become tissue-resident macrophages
either during inflammation or after experimental deletion of tissue
macrophages (48, 86). This underlines the notion that macrophage
polarization is both transient and plastic.
The survival in the tissue of inflammatory monocyte-derived
macrophages raises important questions that need to be answered.
Do monocyte-derived tissue macrophages conserve a “memory” of their past inflammatory activation, thereby becoming
memory macrophages? And, do tissue macrophages resume their
previous functional phenotype in response to a new inflammatory
challenge? Or, do they react as naïve cells?
Memory macrophages (also recently termed “trained monocytes”) have been described, which retain a memory of past challenges (see below). Their fate in the tissue is, however, unknown,
since no long-term experiments have been performed in mammals. It is possible that a part of them dies after reacting to a
new inflammatory challenge. If some of them survive (again, this
possibly depends on the type and magnitude of the new challenge), they would probably behave like inflammatory monocytederived macrophages, i.e., they could become M2-like cells, having
a low level of self-renewal, and may also form a new population of memory macrophages that retain the memory of multiple
challenges.

www.frontiersin.org

FIGURE 4 | Fate of the different monocyte/macrophage populations in
the tissue during the post-inflammatory phase. Tissue-resident
macrophages are in general maintained locally by proliferative self-renewal,
and retain an M2-like functional phenotype. The same situation is
hypothesized for monocyte-derived resident macrophages, since it is not
possible to fully discriminate between the two populations. A number of
cells of these two populations probably die during the inflammatory
reaction. Inflammatory monocyte-derived macrophages can die killed by
the NO they have produced, and the surviving cells can undergo in situ
phenotype conversion and become M2-like tissue-resident macrophages.
In addition, a number of these cells can conserve a “memory” of their past
inflammatory activation, and become trained monocytes/memory
macrophages. Monocytes recruited from the blood during the
post-inflammatory phase can lose the expression of Ly6C and become
Ly6C− cells, subsequently differentiating in M2 macrophages. They may
also become memory macrophages. Memory macrophages that are
present in the tissue, reminiscent of previous inflammatory events, would
probably behave like naïve macrophages upon a new inflammatory
challenge, except for a much quicker reaction, and will, therefore, mostly
die or generate M2-like macrophages or again memory macrophages. Their
life span in the tissue is presently unknown.

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Italiani and Boraschi

Another population that should be considered is that of monocytes recruited from the blood during the post-inflammatory
phase. It is possible that these cells lose Ly6C expression when
in the tissue, thereby becoming Ly6C− cells that subsequently
differentiate in M2 macrophages.
MEMORY MACROPHAGES

It is long known that innate immune responses are higher to a
secondary infection/challenge, and that this higher reactive occurs
whether the new challenge is the same or different from the first
one (cross-protection). An old example is that of mouse peritoneal
macrophages from BCG-infected mice that have little/no activity 7 days after infection, and acquire significant citocydal activity
upon in vitro challenge with LPS or with a wealth of other stimuli,
while naïve macrophages do not (218). Recently, this phenomenon has been re-named trained innate immunity (219). Innate
memory plays an important defensive role in organisms lacking adaptive immunity, such as plants and invertebrates, but it
is evident also in vertebrates lacking functional T and B lymphocytes (220). In these animals, this innate memory mechanism was
shown to involve innate immune cells with low turnover [such as
macrophages and NK cells; (221, 222)] that would be responsible
for improved pathogen recognition through pathogen recognition
receptors, and for an enhanced protective inflammatory response
(223, 224). NK cells could generate a memory response to viruses,
while macrophages retain memory of both bacterial and viral challenges. A logical possibility is that the microorganisms encountered by the host on a regular basis may serve to differentiating
and continually renewing a pool of memory-like macrophages
with enhanced reactivity to infectious challenges. The molecular
mechanisms responsible for shifting macrophages toward a memory status have not yet been elucidated. Putative mechanisms may
involve differences in the monocyte/macrophage population (i.e.,
CD14+ and CD16− ) or changes in the expression of lectin receptors on cell membrane (221), or in the functional phenotype (e.g.,
phagocytosis or protein production), but all are probably underlain by epigenetic reprograming that, through modification of
DNA, post-translational modifications of histones (methylation),
or microRNA, regulates gene expression by inducing dynamic
alterations in the chromatin structure (220). Establishment of
macrophage memory, depending on the experienced challenges,
is likely to rely on epigenetic changes, as these can be at the basis
of a rapid evolution of responsiveness and adaptation to incurring events, thereby allowing to surviving to new environmental
threats (220, 225). Efficacy of many vaccines probably implies the
induction of non-specific macrophage memory that contributes
to the increased resistance to infections. Research in the field of
memory macrophages needs a thorough re-assessment of a large
body of old evidence accumulated in the past decades in the areas
of macrophage activation and of adjuvanticity.

CONCLUSION
An increasing amount of evidence supports four revolutionary
concepts/discoveries on monocytes/macrophages that will force
the researcher to rewrite the books of immunology:
1. The embryonic origin of tissue-resident macrophages, which
raises the need to better understand the features/properties

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

of monocytes (that are no longer simple precursors of tissue
macrophages), and those of macrophages, which are capable of
self-renewal without loss of their differentiated cellular identity.
2. The capacity of monocytes/macrophages to polarize into distinct
functional phenotypes able to initiate and direct virtually all
immune responses, including adaptive ones.
3. The notion of innate memory, an old concept that has been
recently revived with the description of the so-called trained
innate immunity.
4. The importance of macrophage-mediated antigen presentation
in tissue responses, with the identification of antigen-uptaking,
recirculating, and presenting “tissue monocytes,” and with the
notion that tissue macrophages are probably the major APC
upon a second challenge at the tissue level, without need of
recirculation to the lymph nodes.
The central role of monocytes/macrophages in this new view of
immunity implies that innate immunity has a major role in inducing and modulating adaptive immunity (including the induction
of polarized T-cell responses), while on the other hand taking
advantage of adaptive immune mechanisms (e.g., T-cell-derived
cytokines) for modulating its own activity. Thus, new knowledge
on macrophage biology and functions will have a direct impact
on our understanding of immune responses and on the design
of novel therapeutic strategies. For this reason, it is necessary
to overcome several experimental obstacles that delay the full
understanding of the new dynamics and relationships within the
immune system, and that have been identified by the researchers
cited in the review.
For example, to date, transcriptome analysis of monocyte subsets has been done at the basal unstimulated level, showing dramatic differences consistent with a different functional repertoire
for the three types of human monocytes. Circulating monocytes
are most likely “quiescent” (their quiescent status is needed in
order to avoid developing a deleterious intravascular inflammation), while their effector functions only develop after relocation
and activation in the tissue. Thus, the true role of the different
monocyte subsets could be only understood after activation, and
the stimulus-induced transcriptome of these cells will be required.
Further, the models of inflammation used to test the proliferative capacity of resident macrophages have so far been limited
to one or two rounds of tissue repopulation or relatively acute
periods of infection/inflammation. This obviously cannot provide
reliable information on the long-term capacity of macrophage
self-renewal. Moreover, when studying the plasticity and interchangeability of M1 and M2 macrophages, since mixed M1/M2
phenotypes can be found especially in pathological conditions, it is
capital to focus not only on populations but also either at the single
cell level or by lineage-tracking studies (e.g., with mice expressing
Cre ricombinase under the iNOS or arginase promoters, to track
M1 and M2 lineages, respectively). Precautions need to be taken
when drastic experimental procedures such as monocyte depletion or parabiosis are used to study macrophages self-renewal.
These treatments can alter the concentration of circulating CSF1 and CSFR1 signaling, which are important for self-renewal of
resident macrophages under homeostatic conditions, and critical for differentiation of monocytes into tissue macrophages.

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Italiani and Boraschi

Likewise, precautions and appropriate controls need to be implemented when using CCR2-deficient mouse for studying monocyte
recruitment to the tissue, since the CCL2/CCR2 chemokine system is also responsible of the release of monocytes from bone
marrow. Thus, the lack of recruitment of monocytes from the
blood to the tissue could be due to lack of release of monocytes
from bone marrow to the blood, where circulating monocytes are
decreased.
Our final recommendation, therefore, is probably obvious, but
it is anyway important to state it again. We need to re-evaluate
patiently and critically a huge body of experimental evidence that
is already present in the literature. In particular, we need to overcome the lack of consensus in defining and describing the different
macrophage phenotypes (226). Many old studies have already generated information that, in light of our present knowledge, can
become very important and help us to clarify the general picture.
Second recommendation is that of designing experiments very
carefully, keeping in mind that the immune system is redundant
and that the same factor can have different activities, and that the
same activity can be carried out by different factors. Third recommendation: monocytes and macrophages are never isolated in the
body, and what they do and what they become are totally influenced by the surrounding cells and tissue. In vitro systems may only
partially reproduce this complexity. Last recommendation: consider evolution as an incommensurable and most precious source
of information that can greatly help us understand the ontology
and behavior of monocytes and macrophages. Common mechanisms are many, and also species-specific differences exist, thus
we should be able to pick up the relevant common information
without, however, forgetting that human being is not a mouse or
a mosquito.

AUTHOR CONTRIBUTIONS
Paola Italiani wrote the paper; Diana Boraschi contributed to
writing and critically revised the paper.

ACKNOWLEDGMENTS
The authors wish to thank Dr. Charles Mills for helpful discussion and for his continuous assistance in writing this review.
They also thank Elfi Töpfer for figure design and realization. This
work was supported by HUMUNITY (FP7-PEOPLE-INT-2012
GA no. 316383), BioCog (FP7-HEALTH-2013-INNOVATION1 GA no. 602461), and by the grant 2011-2114 of Fondazione
Cariplo (Milano, Italy).

REFERENCES
1. Matzinger P. Friendly and dangerous signals: is the tissue in control? Nat
Immunol (2007) 8:11–3. doi:10.1038/ni0107-11
2. Medzhitov R. Origin and physiological roles of inflammation. Nature (2008)
454:428–35. doi:10.1038/nature07201
3. Nathan C. Points of control in inflammation. Nature (2002) 420:846–52.
doi:10.1038/nature01320
4. van Furth R, Cohn ZA. The origin and kinetics of mononuclear phagocytes.
J Exp Med (1968) 128:415–35. doi:10.1084/jem.128.3.415
5. Taylor PR, Gordon S. Monocytes heterogeneity and innate immunity. Immunity (2003) 19:2–4. doi:10.1016/S1074-7613(03)00178-X
6. Lin H, Lee E, Hestir K, Leo C, Huang M, Bosch E, et al. Discovery of a cytokine
and its receptor by functional screening of the extracellular proteome. Science
(2008) 320:807–11. doi:10.1126/science.1154370

www.frontiersin.org

Monocyte to macrophage differentiation

7. Hume DA, MacDonald KP. Therapeutic applications of macrophage colonystimulating factor-1 (CSF-1) and antagonists of CSF-1 receptor (CSF-1R) signaling. Blood (2012) 119:1810–20. doi:10.1182/blood-2011-09-379214
8. Geissmann F, Gordon S, Hume DA, Mowat AM, Randolph GJ. Unravelling
mononuclear phagocyte heterogeneity. Nat Rev Immunol (2010) 10:453–60.
doi:10.1038/nri2784
9. Jenkins SJ, Hume DA. Homeostasis in the mononuclear phagocyte system.
Trends Immunol (2014) 35:358–67. doi:10.1016/j.it.2014.06.006
10. Geissmann F, Manz MG, Jung S, Sieweke MH, Merad M, Ley K. Development
of monocytes, macrophages, and dendritic cells. Science (2010) 327:656–61.
doi:10.1126/science.1178331
11. Boltjes A, van Wijk F. Human dendritic cell functional specialization in steadystate and inflammation. Front Immunol (2014) 5:131. doi:10.3389/fimmu.
2014.00131
12. Schulz C, Gomez Perdiguero E, Chorro L, Szabo-Rogers H, Cagnard N, Kierdorf K, et al. A lineage of myeloid cells independent of Myb and hematopoietic
stem cells. Science (2012) 336:86–90. doi:10.1126/science.1219179
13. Sieweke MH, Allen JE. Beyond stem cells: self-renewal of differentiated
macrophages. Science (2013) 342:1242974. doi:10.1126/science.1242974
14. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity. J Innate Immun (2014) 6(6). doi:10.1159/000364945
15. Akashi K, Trever D, Miyamoto T, Weissman IL. A clonogenic common myeloid
progenitor that gives rise to all myeloid lineages. Nature (2000) 404:193–7.
doi:10.1038/35004599
16. Fogg DK, Sibon C, Miled C, Jung S, Aucouturier P, Littman DR, et al. A clonogenic bone marrow progenitor specific for macrophages and dendritic cells.
Science (2006) 311:83–7. doi:10.1126/science.1117729
17. Hettinger J, Richards DM, Hansson J, Barra MM, Joschko AC, Krijgsveld M,
et al. Origin of monocytes and macrophages in a committed progenitor. Nat
Immunol (2013) 14:821–30. doi:10.1038/ni.2638
18. Liu K, Victora GD, Schwickert TA, Guermonprez P, Meredith MM, Yao K, et al.
In vivo analysis of dendritic cell development and homeostasis. Science (2009)
324:392–7. doi:10.1126/science.1170540
19. Ajami B, Bennett JL, Krieger C, Tetzlaff W, Rossi FM. Local self-renewal can
sustain CNS microglia maintenance and function throughout adult life. Nat
Neurosci (2007) 10:1538–43. doi:10.1038/nn2014
20. Hamilton JA. Colony-stimulating factor in inflammation and autoimmunity.
Nat Rev Immunol (2008) 8:533–44. doi:10.1038/nri2356
21. Bartocci A, Mastrogiannis DS, Migliorati G, Stockert RJ, Wolkoff AW, Stanley ER. Macrophages specifically regulate the concentration of their own
growth factor in the circulation. Proc Natl Acad Sci U S A (1987) 84:6179–83.
doi:10.1073/pnas.84.17.6179
22. Tushinski RJ, Oliver IT, Guilbert LJ, Tynan PW, Warner JR, Stanley ER.
Survival of mononuclear phagocytes depends on a lineage-specific growth
factor that the differentiated cells selectively destroy. Cell (1982) 28:71–81.
doi:10.1016/0092-8674(82)90376-2
23. Ma X, Lin WY, Chen Y, Stawicki S, Mukhyala K, Wu Y, et al. Structural basis for
the dual recognition of helical cytokines IL-34 and CSF-1 by CSF-1R. Structure
(2012) 20:676–87. doi:10.1016/j.str.2012.02.010
24. Wang Y, Szretter KJ, Vermi W, Gilfillan S, Rossini C, Cella M, et al. IL34 is a tissue-restricted ligand of CSF1R required for the development of
Langerhans cells and microglia. Nat Immunol (2012) 13:753–60. doi:10.1038/
ni.2360
25. Burgess AW, Metcalf D. The nature and action of granulocyte-macrophage
colony stimulating factors. Blood (1980) 56:947–58.
26. Gasson J. Molecular physiology of granulocyte-macrophage colony stimulating
factor. Blood (1991) 77:1131–45.
27. Serbina NV, Jia T, Hohl TM, Pamer EG. Monocyte-mediated defense against
microbial pathogens. Annu Rev Immunol (2008) 26:421–52. doi:10.1146/
annurev.immunol.26.021607.090326
28. Evans HG, Gullick NJ, Kelly S, Pitzalis C, Lord GM, Kirkham BW, et al.
In vivo activated monocytes from the site of inflammation in humans specifically promote Th17 responses. Proc Natl Acad Sci U S A (2009) 106:6232–7.
doi:10.1073/pnas.0808144106
29. Avraham-Davidi I, Yona S, Grunewald M, Landsman L, Cochain C, Silvestre
JS, et al. On-site education of VEGF-recruited monocytes improves their performance as angiogenic and arteriogenic accessory cells. J Exp Med (2013)
210:2611–25. doi:10.1084/jem.20120690

October 2014 | Volume 5 | Article 514 | 63

Italiani and Boraschi

30. Passlick B, Flieger D, Ziegler-Heitbrock HW. Identification and characterization of a novel monocyte subpopulation in human peripheral blood. Blood
(1989) 74:2527–34.
31. Ziegler-Heitbrock L, Ancuta P, Crowe S, Dalod M, Grau V, Hart DN, et al.
Nomenclature of monocytes and dendritic cells in blood. Blood (2010)
116:e74–80. doi:10.1182/blood-2010-02-258558
32. Mobley JL, Leininger M, Madore S, Baginski TJ, Renkiewicz R. Genetic evidence of a functional monocyte dichotomy. Inflammation (2007) 30:189–97.
doi:10.1007/s10753-007-9036-0
33. Zhao C, Zhang H, Wong WC, Sem X, Han H, Ong SM, et al. Identification of
novel functional differences in monocyte subsets using proteomic and transcriptomic methods. J Proteome Res (2009) 8:4028–38. doi:10.1021/pr900364p
34. Ancuta P, Liu KY, Misra V, Wacleche VS, Gosselin A, Zhou X, et al. Transcriptional profiling reveals developmental relationship and distinct biological functions of CD16+ and CD16- monocyte subsets. BMC Genomics (2009) 10:403.
doi:10.1186/1471-2164-10-403
35. Cros J, Cagnard N, Woollard K, Patey N, Zhang SY, Senechal B, et al. Human
CD14dim monocytes patrol and sense nucleic acids and viruses via TLR7
and TLR8 receptors. Immunity (2010) 33:375–86. doi:10.1016/j.immuni.2010.
08.012
36. Wong KL, Tai JJ, Wong WC, Han H, Sem X, Yeap WH, et al. Gene expression profiling reveals the defining features of the classical, intermediate, and
nonclassical human monocyte subsets. Blood (2011) 118:e16–31. doi:10.1182/
blood-2010-12-326355
37. Zawada AM, Rogacev KS, Rotter B, Winter P, Marell RR, Fliser D, et al. SuperSAGE evidence for CD14++CD16+ monocytes as a third monocyte subset.
Blood (2011) 118:e50–61. doi:10.1182/blood-2011-01-326827
38. Frankenberger M, Hofer TP, Marei A, Dayyani F, Schewe S, Strasser C, et al.
Transcript profiling of CD16-positive monocytes reveals a unique molecular
fingerprint. Eur J Immunol (2012) 42:957–74. doi:10.1002/eji.201141907
39. Barnes M, Freudenberg J, Thompson S, Aronow B, Pavlidis P. Experimental
comparison and cross-validation of the Affymetrix and Illumina gene expression analysis platforms. Nucleic Acids Res (2005) 33:5914–23. doi:10.1093/nar/
gki890
40. Ziegler-Heitbrock L, Hofer TP. Toward a refined definition of monocyte subsets. Front Immunol (2013) 4:23. doi:10.3389/fimmu.2013.00023
41. Wong KL, Yeap WH, Tai JJ, Ong SM, Dang TM, Wong SC. The three human
monocyte subsets: implications for health and disease. Immunol Res (2012)
53:41–57. doi:10.1007/s12026-012-8297-3
42. Ingersoll MA, Spanbroek R, Lottaz C, Gautier EL, Frankenberger R, Hoffman R, et al. Comparison of gene expression profiles between human and
mouse monocyte subsets. Blood (2010) 115:e10–9. doi:10.1182/blood-200907-235028
43. Geissmann F, Jung S, Littman DR. Blood monocytes consist of two principal subsets with distinct migratory properties. Immunity (2003) 19:71–82.
doi:10.1016/S1074-7613(03)00174-2
44. Tsou CL, Peters W, Si Y, Slaymaker S, Aslanian AM, Weisberg SP, et al. Critical roles for CCR2 and MPC-3 in monocyte mobilization from bone marrow and recruitment to inflammatory sites. J Clin Invest (2007) 117:902–9.
doi:10.1172/JCI29919
45. Auffray C, Fogg D, Garfa M, Elain G, Join-Lambert O, Kayal S, et al. Monitoring of blood vessels and tissues by a population of monocytes with patrolling
behavior. Science (2007) 317:666–70. doi:10.1126/science.1142883
46. Ziegler-Heitbrock L. Reprint of: monocyte subsets in man and other species.
Cell Immunol (2014). doi:10.1016/j.cellimm.2014.06.008
47. Sunderkötter C, Nikolic T, Dillon MJ, Van Rooijen N, Stehling M, Drevets DA,
et al. Subpopulations of mouse blood monocytes differ in maturation stage and
inflammatory response. J Immunol (2004) 172:4410–7. doi:10.4049/jimmunol.
172.7.4410
48. Yona S, Kim KW, Wolf Y, Mildner A, Varol D, Breker M, et al. Fate mapping
reveals origins and dynamics of monocytes and tissue macrophages under
homeostasis. Immunity (2013) 38:79–91. doi:10.1016/j.immuni.2012.12.001
49. Varol C, Landsman L, Fogg DK, Greenshtein L, Gildor B, Margalit R, et al.
Monocytes give rise to mucosal, but not splenic, conventional dendritic cells.
J Exp Med (2007) 204:171–80. doi:10.1084/jem.20061011
50. MacDonald KP, Palmer JS, Cronau S, Seppanen E, Olver S, Raffelt NC, et al.
An antibody against the colony-stimulating factor 1 receptor depletes the resident subset of monocytes and tissue- and tumor-associated macrophages but

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

does not inhibit inflammation. Blood (2010) 116:3955–63. doi:10.1182/blood2010-02-266296
51. Carlin LM, Stamatiades EG, Auffray C, Hanna RN, Glover L, Vizcay-Barrena
G, et al. Nr4a1-dependent Ly6C(low) monocytes monitor endothelial cells
and orchestrate their disposal. Cell (2013) 153:362–75. doi:10.1016/j.cell.2013.
03.010
52. Hanna RN, Carlin LM, Hubbeling HG, Nackiewicz D, Green AM, Punt JA,
et al. The transcription factor NR4A1 (Nur77) controls bone marrow differentiation and the survival of Ly6C- monocytes. Nat Immunol (2011) 12:778–85.
doi:10.1038/ni.2063
53. Varga T, Mounier R, Gogolak P, Poliska S, Chazaud B, Nagy L. Tissue LyC6macrophages are generated in the absence of circulating LyC6- monocytes and
Nur77 in a model of muscle regeneration. J Immunol (2013) 191:5695–701.
doi:10.4049/jimmunol.1301445
54. Shi C, Pamer EG. Monocyte recruitment during infection and inflammation.
Nat Rev Immunol (2011) 11:762–74. doi:10.1038/nri3070
55. Swirski FK, Nahrendorf M, Etzrodt M, Wildgruber M, Cortez-Retamozo V,
Panizzi P, et al. Identification of splenic reservoir monocytes and their deployment to inflammatory sites. Science (2009) 325:612–6. doi:10.1126/science.
1175202
56. Nguyen KD, Fentress SJ, Qiu Y, Yun K, Cox JS, Chawla A. Circadian gene Bmal1
regulates diurnal oscillations of Ly6C(hi) inflammatory monocytes. Science
(2013) 341:1483–8. doi:10.1126/science.1240636
57. Bain CC, Scott CL, Uronen-Hansson H, Gudjonsson S, Jansson O, Grip O, et al.
Resident and pro-inflammatory macrophages in the colon represent alternative context-dependent fates of the same Ly6Chi monocyte precursors. Mucosal
Immunol (2013) 6:498–510. doi:10.1038/mi.2012.89
58. Rivollier A, He J, Kole A, Valatas V, Kelsall BL. Inflammation switches the differentiation program of Ly6Chi monocytes from anti-inflammatory macrophages
to inflammatory dendritic cells in the colon. J Exp Med (2012) 209:139–55.
doi:10.1084/jem.20101387
59. Jaensson E, Uronen-Hansson H, Pabst O, Eksteen B, Tian J, Coombes JL, et al.
Small intestinal CD103+ dendritic cells display unique functional properties
that are conserved between mice and humans. J Exp Med (2008) 205:2139–49.
doi:10.1084/jem.20080414
60. Tamoutounour S, Guilliams M, Montanana Sanchis F, Liu H, Terhorst D, Malosse C, et al. Origins and functional specialization of macrophages and of
conventional and monocyte-derived dendritic cells in mouse skin. Immunity
(2013) 39:925–38. doi:10.1016/j.immuni.2013.10.004
61. Abtin A, Jain R, Mitchell AJ, Roediger B, Brzoska AJ, Tikoo S, et al. Perivascular
macrophages mediate neutrophil recruitment during bacterial skin infection.
Nat Immunol (2014) 15:45–53. doi:10.1038/ni.2769
62. Epelman S, Lavine KJ, Beaudin AE, Sojka DK, Carrero JA, Calderon B, et al.
Embryonic and adult-derived resident cardiac macrophages are maintained
through distinct mechanisms at steady state and during inflammation. Immunity (2014) 40:91–104. doi:10.1016/j.immuni.2013.11.019
63. Guilliams M, De Kleer I, Henri S, Post S, Vanhoutte L, De Prijck S, et al. Alveolar
macrophages develop from fetal monocytes that differentiate into long-lived
cells in the first week of life via GM-CSF. J Exp Med (2013) 210:1977–92.
doi:10.1084/jem.20131199
64. Jakubzick C, Gautier EL, Gibbings SL, Sojka DK, Schlitzer A, Johnson TE,
et al. Minimal differentiation of classical monocytes as they survey steady-state
tissues and transport antigen to lymph nodes. Immunity (2013) 39:599–610.
doi:10.1016/j.immuni.2013.08.007
65. Capulli M, Paone R, Rucci N. Osteoblast and osteocyte: games without frontiers. Arch Biochem Biophys (2014) 561C:3–12. doi:10.1016/j.abb.2014.05.003
66. Charles JF, Aliprantis AO. Osteoclasts: more than “bone eaters”. Trends Mol Med
(2014) 20:449–59. doi:10.1016/j.molmed.2014.06.001
67. Kikuta J, Ishii M. Osteoclast migration, differentiation and function: novel
therapeutic target for rheumatic diseases. Rheumatology (Oxford) (2013)
52:226–34. doi:10.1093/rheumatology/kes259
68. Boyle WJ, Simonet WS, Lacey DL. Osteoclast differentiation and activation.
Nature (2003) 423:337–42. doi:10.1038/nature01658
69. Dai XM, Ryan GR, Hapel AJ, Dominguez MG, Russell RG, Kapp S, et al.
Targeted disruption of the mouse colony-stimulating factor 1 receptor gene
results in osteopetrosis, mononuclear phagocyte deficiency, increased primitive
progenitor cell frequencies, and reproductive defects. Blood (2002) 99:111–20.
doi:10.1182/blood.V99.1.111

October 2014 | Volume 5 | Article 514 | 64

Italiani and Boraschi

70. Massey HM, Flanagan AM. Human osteoclasts derive from CD14-positive
monocytes. Br J Haematol (1999) 106:167–70. doi:10.1046/j.1365-2141.1999.
01491.x
71. Chen Z, Buki K, Vääräniemi J, Gu G, Väänänen HK. The critical role of IL-34
in osteoclastogenesis. PLoS One (2011) 6:e18689. doi:10.1371/journal.pone.
0018689
72. Matsuzaki K, Udagawa N, Takahashi N, Yamaguchi K, Yasuda H, Shima N, et al.
Osteoclast differentiation factor (ODF) induces osteoclast-like cell formation
in human peripheral blood mononuclear cell cultures. Biochem Biophys Res
Commun (1998) 246:199–204. doi:10.1006/bbrc.1998.8586
73. Hayashi S, Yamane T, Miyamoto A, Hemmi H, Tagaya H, Tanio Y, et al. Commitment and differentiation of stem cells to the osteoclast lineage. Biochem Cell
Biol (1998) 76:911–22. doi:10.1139/o98-099
74. Gautier EL, Shay T, Miller J, Greter M, Jakubzick C, Ivanov S, et al. Geneexpression profiles and transcriptional regulatory pathways that underlie the
identity and diversity of mouse tissue macrophages. Nat Immunol (2012)
13:1118–28. doi:10.1038/ni.2419
75. van Furth R. Origin and kinetics of monocytes and macrophages. Semin Hematol (1970) 7:125–41.
76. van Furth R, Cohn ZA, Hirsch JG, Humphrey JH, Spector WG, Langevoort
HL. The mononuclear phagocyte system: a new classification of macrophages,
monocytes, and their precursor cells. Bull World Health Organ (1972)
46:845–52.
77. Volkman A. The origin and fate of the monocyte. Ser Haematol (1970) 3:69–92.
78. Volkman A. The function of the monocytes. Bibl Haematol (1968) 29:86–97.
79. Epelman S, Lavine KJ, Randolph GJ. Origin and function of tissue
macrophages. Immunity (2014) 41:21–35. doi:10.1016/j.immuni.2014.06.013
80. Orkin SH, Zon LI. Hematopoiesis: an evolving paradigm for stem cell biology.
Cell (2008) 132:631–44. doi:10.1016/j.cell.2008.01.025
81. Cumano A, Godin I. Ontogeny of the hematopoietic system. Annu Rev Immunol
(2007) 25:745–85. doi:10.1146/annurev.immunol.25.022106.141538
82. Tavian M, Péault B. Embryonic development of the human hematopoietic
system. Int J Dev Biol (2005) 49:243–50. doi:10.1387/ijdb.041957mt
83. Enzan H. Electron microscopic studies of macrophages in early human yolk
sacs. Acta Pathol Jpn (1986) 36:49–64.
84. Migliaccio G, Migliaccio AR, Petti S, Mavilio F, Russo G, Lazzaro D, et al.
Human embryonic hemopoiesis. Kinetics of progenitors and precursors
underlying the yolk sac – liver transition. J Clin Invest (1986) 78:51–60.
doi:10.1172/JCI112572
85. Takahashi K, Yamamura F, Naito M. Differentiation, maturation, and proliferation of macrophages in the mouse yolk sac: a light-microscopic, enzymecytochemical, immunohistochemical, and ultrastructural study. J Leukoc Biol
(1989) 45:87–96.
86. Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, et al. Tissueresident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity (2013) 38:792–804.
doi:10.1016/j.immuni.2013.04.004
87. Ginhoux F, Greter M, Leboeuf M, Nandi S, See P, Gokhan S, et al. Fate mapping analysis reveals that adult microglia derive from primitive macrophages.
Science (2010) 330:841–5. doi:10.1126/science.1194637
88. Kierdorf K, Erny D, Goldmann T, Sander V, Schulz C, Perdiguero EG,
et al. Microglia emerge from erythromyeloid precursors via Pu.1- and Irf8dependent pathways. Nat Neurosci (2013) 16:273–80. doi:10.1038/nn.3318
89. Hoeffel G, Wang Y, Greter M, See P, Teo P, Malleret B, et al. Adult Langerhans
cells derive predominantly from embryonic fetal liver monocytes with a minor
contribution of yolk sac-derived macrophages. J Exp Med (2012) 209:1167–81.
doi:10.1084/jem.20120340
90. Ginhoux F, Jung S. Monocytes and macrophages: development pathways
and tissue homeostasis. Nat Rev Immunol (2014) 14:392–404. doi:10.1038/
nri3671
91. Wynn TA, Chawla A, Polland JW. Macrophages biology in development, homeostasis and disease. Nature (2013) 496:445–55. doi:10.1038/nature12034
92. Frankenberger M, Ekici AB, Angstwurm MW, Hoffmann H, Hofer TP, Heimbeck I, et al. A defect of CD16-positive monocytes can occur without disease.
Immunobiology (2013) 218:169–74. doi:10.1016/j.imbio.2012.02.013
93. Hambleton S, Salem S, Bustamante J, Bigley V, Boisson-Dupuis S, Azevedo
J, et al. IRF8 mutations and human dendritic-cell immunodeficiency. N Engl
J Med (2011) 365:127–38. doi:10.1056/NEJMoa1100066

www.frontiersin.org

Monocyte to macrophage differentiation

94. Bigley V, Haniffa M, Doulatov S, Wang XN, Dickinson R, McGovern N, et al.
The human syndrome of dendritic cell, monocyte, B and NK lymphoid deficiency. J Exp Med (2011) 208:227–34. doi:10.1084/jem.20101459
95. Randolph GJ. Macrophages in Marseille. Immunity (2013) 38:619–21. doi:10.
1016/j.immuni.2013.04.002
96. Aziz A, Soucie S, Sarrazin S, Sieweke MH. MafB/c-Maf deficiency enables selfrenewal of differentiated functional macrophages. Science (2009) 326:867–71.
doi:10.1126/science.1176056
97. Czernielewski JM, Demarchez M. Further evidence for the self-reproducing
capacity of Langerhans cells in human skin. J Invest Dermatol (1987) 88:17–20.
doi:10.1111/1523-1747.ep12464659
98. Kanitakis J, Petruzzo P, Dubernard JM. Turnover of epidermal Langerhans’
cells. N Engl J Med (2004) 351:2661–2. doi:10.1056/NEJM200412163512523
99. Merad M, Manz MG, Karsunky H, Wagers A, Peters W, Charo I, et al. Langerhans cells renew in the skin throughout life under steady-state conditions. Nat
Immunol (2002) 3:1135–41. doi:10.1038/ni852
100. Davies LC, Rosas M, Smith PJ, Fraser DJ, Jones SA, Taylor PR. A quantifiable
proliferative burst of tissue macrophages restores homeostatic macrophage
populations after acute inflammation. Eur J Immunol (2011) 41:2155–64.
doi:10.1002/eji.201141817
101. Coggle JE, Tarling JD. The proliferation kinetics of pulmonary alveolar
macrophages. J Leukoc Biol (1984) 35:317–27.
102. Jenkins SJ, Ruckerl D, Cook PC, Jones LH, Finkelman FD, van Rooijen
N, et al. Local macrophages proliferation, rather than recruitment from
the blood, is a signature of TH 2 inflammation. Science (2011) 332:1284–8.
doi:10.1126/science.1204351
103. Jenkins SJ, Ruckerl D, Thomas GD, Hewitson JP, Duncan S, Brombacher F,
et al. IL-4 directly signals tissue-resident macrophages to proliferate beyond
homeostatic levels controlled by CSF-1. J Exp Med (2013) 210:2477–91.
doi:10.1084/jem.20121999
104. Hibbs ML, Quilici C, Kountouri N, Seymour JF, Armes JE, Burgess AW, et al.
Mice lacking three myeloid colony-stimulating factors (G-CSF, GM-CSF, and
M-CSF) still produce macrophages and granulocytes and mount an inflammatory response in a sterile model of peritonitis. J Immunol (2007) 178:6435–43.
doi:10.4049/jimmunol.178.10.6435
105. Chitu V, Stanley ER. Colony-stimulating factor-1 in immunity and inflammation. Curr Opin Immunol (2006) 18:39–48. doi:10.1016/j.coi.2005.11.006
106. Davies LC, Rosas M, Jenkins SJ, Liao CT, Scurr MJ, Brombacher F, et al.
Distinct bone marrow-derived and tissue-resident macrophage lineages proliferate at key stages during inflammation. Nat Commun (2013) 4:1886.
doi:10.1038/ncomms2877
107. Hume DA. Differentiation and heterogeneity in the mononuclear phagocyte
system. Mucosal Immunol (2008) 1:432–41. doi:10.1038/mi.2008.36
108. Tagliani E, Shi C, Nancy P, Tay CS, Pamer EG, Erlebacher A. Coordinate regulation of tissue macrophage and dendritic cell population dynamics by CSF-1.
J Exp Med (2011) 208:1901–16. doi:10.1084/jem.20110866
109. Ryan GR, Dai XM, Dominguez MG, Tong W, Chuan F, Chisholm
O, et al. Rescue of the colony-stimulating factor 1 (CSF-1)-nullizygous
mouse (Csf1(op)/Csf1(op)) phenotype with a CSF-1 transgene and identification of sites of local CSF-1 synthesis. Blood (2001) 98:74–84. doi:10.1182/
blood.V98.1.74
110. Shibata Y, Berclaz PY, Chroneos ZC, Yoshida M, Whitsett JA, Trapnell BC. GMCSF regulates alveolar macrophage differentiation and innate immunity in the
lung through PU.1. Immunity (2001) 15:557–67. doi:10.1016/S1074-7613(01)
00218-7
111. Koenen HJ, Smit MJ, Simmelink MM, Schuurman B, Beelen RH, Meijer S.
Effect of intraperitoneal administration of granulocyte/macrophage-colonystimulating factor in rats on omental milky-spot composition and tumoricidal
activity in vivo and in vitro. Cancer Immunol Immunother (1996) 42:310–6.
doi:10.1007/s002620050288
112. Lieschke GJ, Grail D, Hodgson G, Metcalf D, Stanley E, Cheers C, et al. Mice
lacking granulocyte colony-stimulating factor have chronic neutropenia, granulocyte and macrophage progenitor cell deficiency, and impaired neutrophil
mobilization. Blood (1994) 84:1737–46.
113. Stanley E, Lieschke GJ, Grail D, Metcalf D, Hodgson G, Gall JA, et al. Granulocyte/macrophage colony-stimulating factor-deficient mice show no major perturbation of hematopoiesis but develop a characteristic pulmonary pathology.
Proc Natl Acad Sci U S A (1994) 91:5592–6. doi:10.1073/pnas.91.12.5592

October 2014 | Volume 5 | Article 514 | 65

Italiani and Boraschi

114. Tadokoro CE, de Almeida Abrahamsohn I. Bone marrow-derived macrophages
grown in GM-CSF or M-CSF differ in their ability to produce IL-12 and
to induce IFN-gamma production after stimulation with Trypanosoma cruzi
antigens. Immunol Lett (2001) 77:31–8. doi:10.1016/S0165-2478(01)00197-3
115. El-Behi M, Ciric B, Dai H, Yan Y, Cullimore M, Safavi F, et al. The encephalitogenicity of T(H)17 cells is dependent on IL-1- and IL-23-induced production
of the cytokine GM-CSF. Nat Immunol (2011) 12:568–75. doi:10.1038/ni.2031
116. Fleetwood AJ, Dinh H, Cook AD, Hertzog PJ, Hamilton JA. GM-CSF- and MCSF-dependent macrophage phenotypes display differential dependence on
type I interferon signaling. J Leukoc Biol (2009) 86:411–21. doi:10.1189/jlb.
1108702
117. Fleetwood AJ, Lawrence T, Hamilton JA, Cook AD. Granulocyte-macrophage
colony-stimulating factor (CSF) and macrophage CSF-dependent macrophage
phenotypes display differences in cytokine profiles and transcription factor
activities: implications for CSF blockade in inflammation. J Immunol (2007)
178:5245–52. doi:10.4049/jimmunol.178.8.5245
118. Verreck FA, de Boer T, Langenberg DM, Hoeve MA, Kramer M,Vaisberg E, et al.
Human IL-23-producing type 1 macrophages promote but IL-10-producing
type 2 macrophages subvert immunity to (myco)bacteria. Proc Natl Acad Sci U
S A (2004) 101:4560–5. doi:10.1073/pnas.0400983101
119. Martinez FO, Gordon S, Locati M, Mantovani A. Transcriptional profiling of
the human monocyte-to-macrophage differentiation and polarization: new
molecules and patterns of gene expression. J Immunol (2006) 177:7303–11.
doi:10.4049/jimmunol.177.10.7303
120. Ghigo C, Mondor I, Jorquera A, Nowak J, Wienert S, Zahner SP, et al. Multicolor
fate mapping of Langerhans cell homeostasis. J Exp Med (2013) 210:1657–64.
doi:10.1084/jem.20130403
121. Paolicelli RC. Synaptic pruning by microglia is necessary for normal brain
development. Science (2011) 333:1456–8. doi:10.1126/science.1202529
122. London A, Cohen M, Schwartz M. Microglia and monocyte-derived
macrophages: functionally distinct populations that act in concert in CNS plasticity and repair. Front Cell Neurosci (2013) 7:34. doi:10.3389/fncel.2013.00034
123. Bogie JF, Stinissen P, Hendriks JJ. Macrophages subsets and microglia in multiple sclerosis. Acta Neurophatol (2014) 128:191–213. doi:10.1007/s00401-0141310-2
124. Pollard JW. Trophic macrophages in development and disease. Nat Rev
Immunol (2009) 9:259–70. doi:10.1038/nri2528
125. Blin-Wakkach C, Rouleau M, Wakkach A. Roles of osteoclasts in the control
of medullary hematopoietic niches. Arch Biochem Biophys (2014) 561C:29–37.
doi:10.1016/j.abb.2014.06.032
126. Kleyer A, Schett G. Arthritis and bone loss: a hen and egg story. Curr Opin
Rheumatol (2014) 26:80–4. doi:10.1097/BOR.0000000000000007
127. Randolph GJ. Mechanisms that regulate macrophage burden in atherosclerosis.
Circ Res (2014) 114:1757–71. doi:10.116/CIRCRESAHA.114.301174
128. Klein I, Cornejo JC, Polakos NK, John B, Wuensch SA, Topham DJ, et al. Kupffer cell heterogeneity: functional properties of bone marrow derived and sessile
hepatic macrophages. Blood (2007) 110:4077–85. doi:10.1182/blood-2007-02073841
129. Ganz T. Macrophages and systemic iron homeostasis. J Innate Immun (2012)
4:446–53. doi:10.1159/000336423
130. Sánchez-Valle V, Chávez-Tapia NC, Uribe M, Méndez-Sánchez N. Role of
oxidative stress and molecular changes in liver fibrosis: a review. Curr Med
Chem (2012) 19:4850–60. doi:10.2174/092986712803341520
131. Davies LC, Jenkins SJ, Allen JE, Taylor PR. Tissue-resident macrophages. Nat
Immunol (2013) 14:986–95. doi:10.1038/ni.2705
132. Maus UA, Koay MA, Delbeck T, Mack M, Ermert M, Ermert L, et al. Role of
resident alveolar macrophages in leukocyte traffic into the alveolar air space
of intact mice. Am J Physiol Lung Cell Mol Physiol (2002) 282:L1245–52.
doi:10.1152/ajplung.00453.2001
133. Carey B, Trapnell BC. The molecular basis of pulmonary alveolar proteinosis.
Clin Immunol (2010) 135:223–35. doi:10.1016/j.clin.2010.02.017
134. Nguyen KD, Qiu Y, Cui X, Goh YP, Mwangi J, David T, et al. Alternatively activated macrophages produce catecholamines to sustain adaptive thermogenesis.
Nature (2011) 480:104–8. doi:10.1038/nature10653
135. Zigmond E, Jung S. Intestinal macrophages: well educated exceptions from the
rule. Trends Immunol (2013) 34:162–8. doi:10.1016/j.it.2013.02.001
136. Wallace KL, Zheng LB, Kanazawa Y, Shih DQ. Immunopathology of inflammatory bowel disease. World J Gastroenterol (2014) 20:6–21. doi:10.3748/wjg.
v20.i1.6

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

137. Chorro L, Geissmann F. Development and homeostasis of “resident” myeloid
cells: the case of the Langerhans cell. Trends Immunol (2010) 31:438–45.
doi:10.1016/j.it.2010.09.003
138. Novak ML, Koh TJ. Phenotypic transitions of macrophages orchestrate tissue
repair. Am J Pathol (2013) 183:1352–63. doi:10.1016/j.ajpath.2013.06.034
139. den Haan JM, Kraal G. Innate immune functions of macrophages subpopulations in the spleen. J Innate Immun (2012) 4:437–45. doi:10.1159/000335216
140. Kohyama M, Ise W, Edelson BT, Wilker PR, Hildner K, Mejia C, et al. Role of
Spi-C in the development of red pulp macrophages and splenic iron homeostasis. Nature (2009) 457:318–21. doi:10.1038/nature07472
141. Labonte AC, Tosello-Tramopont AC, Hahn YS. The role of macrophages polarization in infectious and inflammatory diseases. Mol Cells (2014) 37:275–85.
doi:10.14348/molcells.2014.2374
142. Mantovani A, Biswas SK, Galdiero MR, Sica A, Locati M. Macrophage plasticity
and polarixation in tissue repair and remodelling. J Pathol (2013) 229:176–85.
doi:10.1002/path.4133
143. Hume DA. The complexity of constitutive and inducible gene expression in
mononuclear phagocytes. J Leukoc Biol (2012) 92:433–44. doi:10.1189/jlb.
0312166
144. Lech M, Gröbmayr R, Weidenbush M, Anders HJ. Tissue use resident dendritic
cells and macrophages to maintain homeostasis and to regain homeostasis
upon tissue injury: the immunoregulatory role of changing tissue environments. Mediators Inflamm (2012) 2012:951390. doi:10.1155/2012/951390
145. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32:463–88. doi:10.1615/CritRevImmunol.v32.i6.10
146. Barth MW, Hendrzak JA, Melnicoff MJ, Morahan PS. Review of the
macrophage disappearance reaction. J Leukoc Biol (1995) 57:361–7.
147. Struyf S, Van Collie E, Paemen L, Put W, Lenaerts JP, Proost P, et al. Synergistic induction of MCP-1 and -2 by IL-1β and interferons in fibroblasts and
epithelial cells. J Leukoc Biol (1998) 36:364–72.
148. Shi C, Jia T, Mendez-Ferrer S, Hohl TM, Serbina NV, Lipuma L, et al. Bone marrow mesenchymal stem and progenitor cells induce monocyte emigration in
response to circulating toll-like receptor ligands. Immunity (2011) 34:590–601.
doi:10.1016/j.immuni.2011.02.016
149. Serbina NV, Pamer EG. Monocyte emigration from bone marrow during bacterial infection requires signals mediated by chemokine receptor CCR2. Nat
Immunol (2006) 7:311–7. doi:10.1038/ni1309
150. Nahrendorf M, Swirski FK, Aikawa E, Stangenberg L, Wurdinger T, Figueiredo
JL, et al. The healing myocardium sequentially mobilizes two monocyte subsets
with divergent and complementary functions. J Exp Med (2007) 204:3037–47.
doi:10.1084/jem.20070885
151. Willenborg S, Lucas T, van Loo G, Knipper JA, Krieg T, Haase I, et al. CCR2
recruits an inflammatory macrophage subpopulation critical for angiogenesis in tissue repair. Blood (2012) 120:613–25. doi:10.1182/blood-2012-01403386
152. Ishida Y, Gao JL, Murphy PM. Chemokine receptor CX3CR1 mediates skin
wound healing by promoting macrophage and fibroblast accumulation and
function. J Immunol (2008) 180:569–79. doi:10.4049/jimmunol.180.1.569
153. White GE, Greaves DR. Fractalkine: a survivor’s guide: chemokines as antiapoptotic mediators. Arterioscler Thromb Vasc Biol (2012) 32:589–94. doi:10.
1161/ATVBAHA.111.237412
154. Wang Y, Cui L, Gonsiorek W, Min SH, Anilkumar S, Rosenblum J, et al. CCR2
and CXCR4 regulate peripheral blood monocyte pharmacodynamics and link
to efficacy in experimental autoimmune encephalomyelitis. J Inflamm (Lond)
(2009) 6:32. doi:10.1186/1476-9255-6-32
155. Debien E, Mayol K, Biajoux V, Daussy C, De Aguero MG, Taillardet M, et al.
S1PR5 is pivotal for the homeostasis of patrolling monocytes. Eur J Immunol
(2013) 43:1667–75. doi:10.1002/eji.201343312
156. Arnold L, Henry A, Poron F, Baba-Amer Y, van Rooijen N, Plonquet A, et al.
Inflammatory monocytes recruited after skeletal muscle injury switch into
antiinflammatory macrophages to support myogenesis. J Exp Med (2007)
204:1057–69. doi:10.1084/jem.20070075
157. Crane MJ, Daley JM, van Houtte O, Brancato SK, Henry WL Jr, Albina JE. The
monocyte to macrophage transition in the murine sterile wound. PLoS One
(2014) 9:e86660. doi:10.1371/journal.pone.0086660
158. Gautier EL, Jakubzick C, Randolph GJ. Regulation of the migration and survival
of monocyte subsets by chemokine receptors and its relevance to atherosclerosis. Arterioscler Thromb Vasc Biol (2009) 29:1412–8. doi:10.1161/ATVBAHA.
108.180505

October 2014 | Volume 5 | Article 514 | 66

Italiani and Boraschi

159. Robbins CS, Hilgendorf I, Weber GF, Theurl I, Iwamoto Y, Figueiredo JL, et al.
Local proliferation dominates lesional macrophage accumulation in atherosclerosis. Nat Med (2013) 19:1166–72. doi:10.1038/nm.3258
160. Chorro L, Sarde A, Li M, Woollard KJ, Chambon P, Malissen B, et al. Langerhans cell (LC) proliferation mediates neonatal development, homeostasis, and
inflammation-associated expansion of the epidermal LC network. J Exp Med
(2009) 206:3089–100. doi:10.1084/jem.20091586
161. Ajami B, Bennett JL, Krieger C, McNagny KM, Rossi FM. Infiltrating monocytes trigger EAE progression, but do not contribute to the resident microglia
pool. Nat Neurosci (2011) 14:1142–9. doi:10.1038/nn.2887
162. Tymoszuk P, Evens H, Marzola V, Wachowicz K, Wasmer MH, Datta S,
et al. In situ proliferation contributes to accumulation of tumor-associated
macrophages in spontaneous mammary tumors. Eur J Immunol (2014)
44:2247–62. doi:10.1002/eji.201344304
163. Amano SU, Cohen JL, Vangala P, Tencerova M, Nicoloro SM, Yawe JC, et al.
Local proliferation of macrophages contributes to obesity-associated adipose
tissue inflammation. Cell Metab (2014) 19:162–71. doi:10.1016/j.cmet.2013.
11.017
164. Mills CD. Macrophage arginine metabolism to ornithine/urea or nitric
oxide/citrulline: a life or death issue. Crit Rev Immunol (2001) 21:399–426.
doi:10.1615/CritRevImmunol.v21.i5.10
165. Zigmond E, Varol C, Farache J, Elmaliah E, Satpathy AT, Friedlander G, et al.
Ly6Chi monocytes in the inflamed colon give rise to proinflammatory effector cells and migratory antigen-presenting cells. Immunity (2012) 37:1076–90.
doi:10.1016/j.immuni.2012.08.026
166. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.4049/
jimmunol.164.12.6166
167. Pesce JT, Ramalingam TR, Mentink-Kane MM, Wilson MS, El Kasmi
KC, Smith AM, et al. Arginase-1-expressing macrophages suppress Th2
cytokine-driven inflammation and fibrosis. PLoS Pathog (2009) 5:e1000371.
doi:10.1371/journal.ppat.1000371
168. MacMicking J, Xie QW, Nathan C. Nitric oxide and macrophage function. Annu
Rev Immunol (1997) 15:323–50. doi:10.1146/annurev.immunol.15.1.323
169. Van den Bossche J, Lamers WH, Koehler ES, Geuns JM, Alhonen L, Uimari A,
et al. Pivotal advance: arginase-1-independent polyamine production stimulates the expression of IL-4-induced alternatively activated macrophage markers while inhibiting LPS-induced expression of inflammatory genes. J Leukoc
Biol (2012) 91:685–99. doi:10.1189/jlb.0911453
170. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol (2011) 11:723–37. doi:10.1038/nri3073
171. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine
system in diverse forms of macrophage activation and polarization. Trends
Immunol (2004) 25:677–86. doi:10.1016/j.it.2004.09.015
172. Biswas SK, Mantovani A. Orchestration of metabolism by macrophages. Cell
Metab (2012) 15:432–7. doi:10.1016/j.cmet.2011.11.013
173. Rodríguez-Prados JC, Través PG, Cuenca J, Rico D, Aragonés J, MartínSanz P, et al. Substrate fate in activated macrophages: a comparison between
innate, classic, and alternative activation. J Immunol (2010) 185:605–14.
doi:10.4049/jimmunol.0901698
174. Janeway CA Jr, Medzhitov R. Innate immune recognition. Annu Rev Immunol
(2002) 20:197–216. doi:10.1146/annurev.immunol.20.083001.084359
175. Kawai T, Akira S. Toll-like receptors and their crosstalk with other innate receptors in infection and immunity. Immunity (2011) 34:637–50. doi:10.1016/j.
immuni.2011.05.006
176. Unanue ER. Antigen-presenting function of the macrophage. Annu Rev
Immunol (1984) 2:395–428. doi:10.1146/annurev.iy.02.040184.002143
177. Gordon S, Taylor PR. Monocyte and macrophage heterogeneity. Nat Rev
Immunol (2005) 5:953–64. doi:10.1038/nri1733
178. Gordon S. Alternative activation of macrophages. Nat Rev Immunol (2003)
3:23–35. doi:10.1038/nri978
179. Martinez FO, Sica A, Mantovani A, Locati M. Macrophage activation and polarization. Front Biosci (2008) 13:453–61. doi:10.2741/2692
180. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122:787–95. doi:10.1172/JCI59643
181. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8:958–69. doi:10.1038/nri2448
182. Mantovani A, Sozzani S, Locati M, Allavena P, Sica A. Macrophage polarization:
tumor-associated macrophages as a paradigm for polarized M2 mononuclear

www.frontiersin.org

Monocyte to macrophage differentiation

phagocytes. Trends Immunol (2002) 23:549–55. doi:10.1016/S1471-4906(02)
02302-5
183. Fleming BD, Mosser DM. Regulatory macrophages: setting the threshold for
therapy. Eur J Immunol (2011) 41:2498–502. doi:10.1002/eji.201141717
184. Manjili MH, Wang XY, Abrams S. Evolution of our understanding of myeloid
regulatory cells: from MDSCs to Mregs. Front Immunol (2014) 5:303. doi:10.
3389/fimmu.2014.00303
185. Martinez FO, Gordon S. The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000Prime Rep (2014) 6:13. doi:10.12703/P6-13
186. Okabe Y, Medzhitov R. Tissue-specific signals control reversible program
of localization and functional polarization of macrophages. Cell (2014)
157:832–44. doi:10.1016/j.cell.2014.04.016
187. Pettersen JS, Fuentes-Duculan J, Suárez-Fariñas M, Pierson KC, Pitts-Kiefer A,
Fan L, et al. Tumor-associated macrophages in the cutaneous SCC microenvironment are heterogeneously activated. J Invest Dermatol (2011) 131:1322–30.
doi:10.103/jid.2011.9
188. Vogel DY, Vereyken EJ, Glim JE, Heijnen PD, Moeton M, van der Valk P, et al.
Macrophages in inflammatory multiple sclerosis lesions have an intermediate activation status. J Neuroinflammation (2013) 10:35. doi:10.1186/17422094-10-35
189. Lawrence T, Natoli G. Transcriptional regulation of macrophage polarization: enabling diversity with identity. Nat Rev Immunol (2011) 11:750–61.
doi:10.1038/nri3088
190. Lin SL, Castaño AP, Nowlin BT, Lupher ML Jr, Duffield JS. Bone marrow
Ly6Chigh monocytes are selectively recruited to injured kidney and differentiate into functionally distinct populations. J Immunol (2009) 183:6733–43.
doi:10.4049/jimmunol.0901473
191. Italiani P, Mazza EM, Lucchesi D, Cifola I, Gemelli C, Grande A, et al. Transcriptomic profiling of the development of the inflammatory response in
human monocytes in vitro. PLoS One (2014) 9:e87680. doi:10.1371/journal.
pone.0087680
192. Mylonas KJ, Nair MG, Prieto-Lafuente L, Paape D, Allen JE. Alternatively
activated macrophages elicited by helminth infection can be reprogrammed
to enable microbial killing. J Immunol (2009) 182:3084–94. doi:10.4049/
jimmunol.0803463
193. Stout RD, Jiang C, Matta B, Tietzel I,Watkins SK, Suttles J. Macrophages sequentially change their functional phenotypes in response to changes in microenvironmental influences. J Immunol (2005) 175:342–9. doi:10.4049/jimmunol.
175.1.342
194. Egawa M, Mukai K, Yoshikawa S, Iki M, Mukaida N, Kawano Y, et al. Inflammatory monocytes recruited to allergic skin acquire an anti-inflammatory M2
phenotype via basophil-derived interleukin-4. Immunity (2013) 38:570–80.
doi:10.1016/j.immuni.2012.11.014
195. Gundra UM, Girgis NM, Ruckerl D, Jenkins S, Ward LN, Zurtz ZD, et al.
Alternatively activated macrophages derived from monocytes and tissue
macrophages are phenotypically and functionally distinct. Blood (2014)
123:e110–22. doi:10.1182/blood-2013-08-520619
196. Albina JE, Caldwell MD, Henry WL Jr, Mills CD. Regulation of macrophages
functions by l-arginine. J Exp Med (1989) 169:1021–9. doi:10.1084/jem.169.
3.1021
197. Morris SM. Recent advances in arginine metabolism: roles and regulation of
the arginases. Br J Pharmacol (2009) 157:922–30. doi:10.1111/ji.1476-5381.
2009.00278
198. Vodovoz Y, Bogdan C, Paik J, Xie QW, Nathan C. Mechanisms of suppression
of macrophages nitric oxide release by transforming growth factor beta. J Exp
Med (1993) 178:605–13. doi:10.1084/jem.178.2.605
199. Biswas SK, Lopez-Collazo E. Endotoxin tolerance: new mechanisms, molecules
and clinical significance. Trends Immunol (2009) 30:475–87. doi:10.1016/j.it.
2009.07.009
200. Bastard JP, Maachi M, Lagathu C, Kim MJ, Caron M, Vidal H, et al. Recent
advances in the relationship between obesity, inflammation, and insulin resistance. Eur Cytokine Netw (2006) 17:4–12.
201. Martín-Fuentes P, Civeira F, Recalde D, García-Otín AL, Jarauta E, Marzo
I, et al. Individual variation of scavenger receptor expression in human
macrophages with oxidized low-density lipoprotein is associated with a differential inflammatory response. J Immunol (2007) 179:3242–8. doi:10.4049/
jimmunol.179.5.3242
202. Swann JB, Vesely MD, Silva A, Sharkey J, Akira S, Schreiber RD, et al.
Demonstration of inflammation-induced cancer and cancer immunoediting

October 2014 | Volume 5 | Article 514 | 67

Italiani and Boraschi

203.

204.

205.

206.
207.
208.
209.

210.

211.

212.

213.

214.

215.

216.

217.

during primary tumorigenesis. Proc Natl Acad Sci U S A (2008) 105:652–6.
doi:10.1073/pnas.0708594105
Biswas SK, Mantovani A. Macrophage plasticity and interaction with lymphocyte subsets: cancer as a paradigm. Nat Immunol (2010) 11:889–96.
doi:10.1038/ni.1937
Stout RD, Suttles J. Functional plasticity of macrophages: reversible adaptation to changing microenvironments. J Leukoc Biol (2004) 76:509–13.
doi:10.1189/jlb.0504272
Biswas SK, Chittezhath M, Shalova IN, Lim JY. Macrophage polarization and
plasticity in health and disease. Immunol Res (2012) 53:11–24. doi:10.1007/
s12026-012-8291-9
Medzhitov R, Horng T. Transcriptional control of the inflammatory response.
Nat Rev Immunol (2009) 9:692–703. doi:10.1038/nri2634
Smale ST. Selective transcription in response to an inflammatory stimulus. Cell
(2010) 140:833–44. doi:10.1016/j.cell.2010.01.037
Ley K. The second touch hypothesis: T cell activation, homing and polarization.
F1000Res (2014) 3:37. doi:10.12688/f1000research.3_37.v02
Kawakami N, Nägerl UV, Odoardi F, Bonhoeffer T, Wekerle H, Flügel A.
Live imaging of effector cell trafficking and autoantigen recognition within
the unfolding autoimmune encephalomyelitis lesion. J Exp Med (2005)
201:1805–14. doi:10.1084/jem.20050011
Wakim LM, Waithman J, van Rooijen N, Heath WR, Carbone FR. Dendritic
cell-induced memory T cell activation in nonlymphoid tissues. Science (2008)
319:198–202. doi:10.1126/science.1151869
Muller AJ, Filipe-Santos O, Eberl G, Aebischer T, Späth GF, Bousso P. CD4+ T
Cells rely on a cytokine gradient to control intracellular pathogens beyond sites
of antigen presentation. Immunity (2012) 37:147–57. doi:10.1016/j.immuni.
2012.05.015
Choi JH, Do Y, Cheong C, Koh H, Boscardin SB, Oh YS, et al. Identification
of antigen-presenting dendritic cells in mouse aorta and cardiac valves. J Exp
Med (2009) 206:497–505. doi:10.1084/jem.20082129
Choi JH, Cheong C, Dandamudi DB, Park CG, Rodriguez A, Mehandru S,
et al. Flt3 signaling-dependent dendritic cells protect against atherosclerosis.
Immunity (2011) 35:819–31. doi:10.1016/j.immuni.2011.09.014
Koltsova EK, Garcia Z, Chodaczek G, Landau M, McArdle S, Scott SR, et al.
Dynamic T cell-APC interactions sustain chronic inflammation in atherosclerosis. J Clin Invest (2012) 122:3114–26. doi:10.1172/JCI61758
Lindquist RL, Shakhar G, Dudziak D, Wardemann H, Eisenreich T, Dustin
ML, et al. Visualizing dendritic cell networks in vivo. Nat Immunol (2004)
5:1243–50. doi:10.1038/ni1139
Ifrim DC, Quintin J, Joosten LA, Jacobs C, Jansen T, Jacobs L, et al. Trained
immunity or tolerance: opposing functional programs induced in human
monocytes after engagement of various pattern recognition receptors. Clin
Vaccine Immunol (2014) 21:534–45. doi:10.1128/CVI.00688-13
Janssen WJ, Barthel L, Muldrow A, Oberley-Deegan RE, Kearns MT, Jakubzick
C, et al. Fas determines differential fates of resident and recruited macrophages
during resolution of acute lung injury. Am J Respir Crit Care Med (2011)
184:547–60. doi:10.1164/rccm.201011-1891OC

Frontiers in Immunology | Inflammation

Monocyte to macrophage differentiation

218. Boraschi D, Meltzer MS. Defective tumoridical activity of macrophages
of A/J mice. II. Comparison of the macrophages cytotoxic defect of A7J
mice with that of lipid A-unresponsive C3H/HeJ mice. J Immunol (1979)
122:1592–7.
219. Netea MG, Quintin J, van der Meer JW. Trained immunity: a memory for
innate host defense. Cell Host Microbe (2011) 9:355–61. doi:10.1016/j.chom.
2011.04.006
220. Quintin J, Saeed S, Martens JH, Giamarellos-Bourboulis EJ, Ifrim DC, Logie
C, et al. Candida albicans infection affords protection against reinfection via
functional reprogramming of monocytes. Cell Host Microbe (2012) 12:223–32.
doi:10.1016/j.chom.2012.06.006
221. Bowdish DM, Loffredo MS, Mukhopadhyay S, Mantovani A, Gordon S.
Macrophages receptor implicated in the “adaptive” form of innate immunity.
Microbe Infect (2007) 9:1680–7. doi:10.1016/j.micinf.2007.09.002
222. Vivier E, Raulet DH, Moretta A, Caligiuri MA, Zitvogel L, Lanier LL, et al.
Innate or adaptive immunity? The example of natural killer cells. Science (2011)
331:44–9. doi:10.1126/science.1198687
223. Netea MG. Training innate immunity: the changing concept of immunological memory in innate host defence. Eur J Clin Invest (2013) 43:881–4.
doi:10.1111/eci.12132
224. Quintin J, Cheng SC, van der Meer JW, Netea MG. Innate immune memory: towards a better understanding of host defense mechanisms. Curr Opin
Immunol (2014) 29:1–7. doi:10.1016/j.coi.2014.02.006
225. Ostuni R, Piccolo V, Barozzi I, Polletti S, Termanini A, Bonifacio S, et al.
Latent enhancers activated by stimulation in differentiated cells. Cell (2013)
152:157–71. doi:10.1016/j.cell.2012.12.018
226. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental
guidelines. Immunity (2014) 41:14–20. doi:10.1016/j.immuni.2014.06.008

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 22 August 2014; paper pending published: 23 September 2014; accepted: 02
October 2014; published online: 17 October 2014.
Citation: Italiani P and Boraschi D (2014) From monocytes to M1/M2
macrophages: phenotypical vs. functional differentiation. Front. Immunol. 5:514. doi:
10.3389/fimmu.2014.00514
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Italiani and Boraschi. This is an open-access article distributed under
the terms of the Creative Commons Attribution License (CC BY). The use, distribution
or reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
accepted academic practice. No use, distribution or reproduction is permitted which
does not comply with these terms.

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OPINION ARTICLE
published: 03 December 2014
doi: 10.3389/fimmu.2014.00612

Spatial, temporal, and functional aspects of macrophages
during “The Good, the Bad, and the Ugly” phases
of inflammation
Robert A. Harris*
Applied Immunology and Immunotherapy, Department of Clinical Neurosciences, Karolinska Institutet, Karolinska Hospital, Stockholm, Sweden
*Correspondence: robert.harris@ki.se
Edited by:
Charles Dudley Mills, BioMedical Consultants, USA
Reviewed by:
Thomas E. Morrison, University of Colorado School of Medicine, USA
Markus Munder, University Medicine Mainz, Germany
Keywords: macrophage activation, M1/M2 macrophages, microenvironment, phenotype, immunotherapy, adoptive

In Sergio Leone’s classic western drama
“The Good, the Bad, and the Ugly” the final
scene depicts the three protagonists, each
with their specific personality trait and fastdraw capability, assembled in a graveyard
for a shoot-out. The analogy is thus to
focal sites of inflammation, with different
subpopulations of myeloid cells assembled
within a tissue in proximity to each-other,
but with different functional phenotypes,
associated surface marker expression, and
enacting different functions. The basic
macrophage functional states are described
as pro-inflammatory, anti-inflammatory,
and wound healing, respectively. The M1
versus M2 phenotypic paradigm was first
coined to distinguish macrophage populations and has been instrumental in increasing our knowledge of myeloid biology
(1). Despite more recent suggestions that
there is a continuum of activation states
between the extremes of M1- and M2type responses (2–4), this partly reflects
an over-emphasis on cell surface phenotypes. We should now have the technologies to be able to assess the relevance of
specific cells within specific microenvironments within a given healthy or diseased
tissue. The issue of functionality (irrespective of surface phenotype) and the concept of functional diversity within distinct
microenvironments within a tissue have
been less studied, and is the focus of this
commentary.
Let us first consider in simplistic
terms three stages of an inflammatory
response (Figure 1A): the “Good” noninflammatory phase in which normal tissue homeostasis is maintained by resident
macrophages (green arrows); the “Bad”

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phase in which potentially tissue-damaging
macrophage functionality is initiated due
to damage, infection, or autoimmunity
(pro-inflammatory) or tumor development (anti-inflammatory) (red); and the
“Ugly” phase representing a failure to
down-regulate the initial response that
results in chronic pathogenesis and tissue damage (blue), i.e., an inability to
return to the “Good” phase through healing. The definition of “good” and “bad”
in this sense will depend on the setting –
a pro-inflammatory response to an infection may be desirable, but if uncontrolled
may lead to tissue damage. Likewise, while
anti-inflammatory responses beneficially
modulate autoimmune reactions, they contribute to tumor development. The salient
point is that macrophage function will vary
during these different phases.

MICROENVIRONMENTAL VARIATION IN
INFLAMMATION
The aforementioned basic view reflects the
overall functionality of an inflamed tissue, but does not take into account the
potential compartmentalization of a tissue,
and that these processes may be occurring simultaneously in different areas of
the affected tissue. Consider the image
of a slice of multiple sclerosis subcortical white matter brain tissue (Figure 1B).
Luxol fast blue staining (left) reveals
healthy regions of myelin (dark blue),
focal areas of demyelination (red arrows),
and focal “shadow plaques” that represent remyelination (green arrows) (5). Two
disparate processes – pathogenic demyelination and healing remyelination, occurring side-by-side, implicating that within

these physically separate microenvironments immune cells such as macrophages
might be conducting different processes –
damage or repair (right image). In support
of this, recent histopathological evidence
reveals mixed macrophage phenotypes in
human MS lesions (6).
Understanding of the concept of specific microenvironments within tissues
is increasing, not least within tumor
immunology (7). The ambiguous role of
tumor associated macrophages (TAMs) in
tumor progression is reflected by TAMs
both actively augmenting cancer cell proliferation, invasion, metastasis, and angiogenesis by releasing cytokines, growth factors, enzymes, and angiogenic factors, but
they also kill cancer cells. These varied
activities encompass both M1 and M2
macrophage properties. It is counterintuitive that such diverse tumor-promoting, or
conversely anti-tumoral, activities are performed by a single TAM cell type, so the
existence of distinct TAM subpopulations
associated with different intra-tumoral
microenvironments is predicted (8). The
source of the TAM may also influence their
functions. For example, in gliomas TAM
may be either resident brain microglia or
blood infiltrating macrophages. Molecularly and functionally distinct TAM subpopulations may thus coexist in tumors,
the heterogeneity depending on cancer
type, stage of tumor progression and specific location within the tumor tissue (9,
10). A dynamic“switch”in TAM phenotype
during tumor progression may explain
the mixed activation state of TAM subsets present in different established tumors,
and in certain models a switch from TAM

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Harris

Microenvironmental macrophage functionality

expect this issue to be further addressed in
forthcoming years.

FUNCTION VERSUS FORM DURING
INFLAMMATION

FIGURE 1 | (A) Temporal and microenvironmental macrophage functionality during immune activation.
(B) Left hand panel is reproduced, with permission, from Adams, C. W. M. A Colour Atlas of Multiple
Sclerosis and Other myelin Disorders (©1989 Wolfe Medical Publications Ltd.).

is linked to tumor progression (11). It
is noteworthy that the “switch” that is
often referred to in the literature may
rather reflect a relative predominance in
M1 or M2 cell numbers rather than a
full phenotypical/functional change of a
single cell. Different macrophage populations induced during tumor progression have also been reported to occupy
different microenvironments within the
tumor mass (12). In a murine hepatocellular carcinoma model, the MHC Class
IIhigh TAM population (M1-like) was associated with tumor growth suppression during early tumor growth while MHC IIlow
TAMs (M2-like) dominated as the tumor
progressed (13).
Investigation of the spatial distribution of macrophage phenotypes in human
plaques at different stages of atherosclerosis development also reveals microenvironment variations. M1 is the predominant phenotype in rupture-prone shoulder
regions, and M2 in the adventitia (14).
Likewise, in models of lung inflammation induced by butylated hydroxytoluene
or Mycobacterium tuberculosis there is an
initial M1 activation that progresses to
M2 (15). However, granuloma-associated
macrophages during active infection may
retain an M1 phenotype while nearby
uninfected alveolar macrophages are M2.

Clearly, at various sites of inflammation
then individual subpopulations of cells
contribute to specific microenvironments
in different ways.

INFLAMMATION IS A TEMPORALLY
EVOLVING CIRCUMSTANCE
Another aspect to consider when homogenizing tissues for cell purification and
subsequent FACS or RT-PCR analyses is
not only the geographical gradient of
macrophage activation states that will be
lost but also a temporal element. For
example, given the realization that resident
macrophages and circulating monocytes
are fundamentally different cell types (16,
17), consider the macrophage disappearance reaction (18) in which initial immune
activation within a tissue results in efflux
of resident cells and infiltration of circulating cells, this cellular flux being reversed on
eradication of the offending stimulant. If
one would take a snapshot in the tissue and
sample the phenotype of the macrophages
present at a given time, it is difficult to
be sure whether they are coming or going
from the site of interest. Obviously, this can
impact on their functional relevance to the
ongoing immunological process and especially in our scientific interpretation. Now
that small rodent PET/CT/MRI imaging
is becoming more standardized, one can

Frontiers in Immunology | Molecular Innate Immunity

A host requires basic macrophage functions
in order to survive. These functions are
sometimes less easily measured than other
surrogate markers such as cell surface proteins. Consequently, there is a wide range of
surface markers, cytokines and chemokines
reported to distinguish M1 and M2 activation states (mostly in vitro). However,
the use of such “markers” without parallel assessment of functions can result in
conflicting results.
Variation in published results might
be explained by variation in employed
activation protocol, difference in rodent
strain (19) or human donor to which the
same protocol is applied. Take IL-10, for
example, a prototypic anti-inflammatory
cytokine that can be produced by M2 and
M1 cells. Both the production and lack of
production of IL-10 in M-CSF-stimulated
human M2 monocytes have been described
(21). Reliance on surface marker expression can also be particularly misleading –
if anti-inflammatory M2 states receive an
additional LPS stimulation in vitro, then
while expression of CD86 and MHC II
might become upregulated (“M1”) the IL10 production is actually enhanced (22).
Given the dominant functional role of IL10 as an immunosuppressive cytokine, then
functionally such cells are more potent
M2, despite starting to develop an M1
surface. Similarly, in alcoholic hepatitis,
liver M2 macrophages were determined to
express M1-associated receptors (23). It
would thus seem that there is a necessity to
distinguish between surface and functional
phenotypes.
Clearly, the biological functions in
microenvironments should be more
important than any other phenotype, and
one expects as much functional variation as there is in M1–M2 phenotypic
definition. Even the basic morphology of
activated cells in vitro is reported to be
exact for different phenotypes, yet closer
examination reveals this is not necessarily the case (20, 24, 25). In our own
study in which we applied IL-4, IL-10,
and TGFβ simultaneously, cells of three
different morphologies were apparent in
the same culture well, representing the

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Harris

three distinct morphologies observed if
single cytokines were applied in separate
cultures (26). Does this imply microheterogeneity even in the cell culture medium
containing a mixture of cytokines, such
that the first cytokine receptor ligated
on a cell surface dictates the morphology of that specific cell? Whether the
sequence of activation is of any consequence will depend on whether there are
actually any kinetic effects that impinge
on the final morphology or function.
This may seem a trivial issue but a
recent publication elegantly demonstrated
precisely why this is important by studying human macrophage transcriptomes
using different combinations of activation
stimulants (4).
The challenge is thus how to quantify functional phenotypes in microenvironments. We have a limited range of
markers that can be applied immunohistologically. Detailed knowledge about
expression/regulation of expression for
each macrophage population in each tissue,
within each microenvironment within a tissue, during both resting and inflammatory
states, is currently lacking. Development of
conditional knockout mouse strains lacking resident/peripheral macrophage populations is one approach that is warranted,
for example, to distinguish the relative
roles of infiltrating macrophages and resident microglia during brain tumor development. Alternatively, the use of single
cell laser capture and proteomic or genetic
analyses might be one modern approach
to explore what individual cells within
a particular microenvironment do within
their niche, although function will only be
inferred from these analyses (27, 28). The
basic assumption is that all cells are equal
and that the relative numbers of different
subpopulations will ultimately define the
functional state of the tissue. Increasing
evidence challenges this assumption, and
in our hands co-culture of pre-activated
M1- and M2-type populations demonstrated a clear dominant phenotype of
M2 cells (pre-activated with a combination of IL-4/IL-10/TGFβ) (29). Even if single cells are phenotyped and their relative
numbers are quantified within a tissue or
microenvironment, it remains difficult to
predict the net functional activity or interplay in vivo if they are not functionally
equivalent.

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Microenvironmental macrophage functionality

THERAPEUTIC MANIPULATION OF
CHRONIC INFLAMMATORY
MICROENVIRONMENTS
A final question is whether chronic
inflammatory states such as autoimmune
diseases represent a failure to downregulate pro-inflammatory M1-mediated
tissue destruction (i.e., a deficiency in antiinflammatory M2 function), or whether
this reflects a lack of healing M2 functionality. It follows that a stochastic alteration
of the relative M1/M2 functions within
microenvironments represents a feasible
therapeutic approach. Earlier work indicated that M2 cells accumulate at the
edge of the tissue damage in the setting
of spinal cord injury (30). If large numbers of “therapeutic cells” could be applied
to inflammatory microenvironments (e.g.,
through local stimulation or cell transfer) then they should be able to exert tailored “local” immunomodulatory effects.
In our experience adoptively transferred
pre-activated anti-inflammatory M2 cells
resulted in clinical abrogation of both T1D
(22) and MOG-EAE (26) disease courses.
In addition to a cell therapy approach,
there are new generations of agents aimed
at specific conversion of macrophage phenotypes within microenvironments, such
as TAM conversion to M1 within tumors
using docetaxel and phosphatidylserinetargeting antibody (31). The effectiveness
of this approach was first reported many
years ago (32). The major challenge will
be to access the microenvironments specifically rather than systemically administrating an agent and hoping for its specific
access to the target area.

LOOKING BACK AND LOOKING AHEAD
The last decade has heralded a revolution
in our understanding of immune mechanisms and particularly the critical role of
macrophages in both innate and adaptive
responses. During the coming decade, we
can expect a refinement of this knowledge when the functions of individual
cells and their specific contributions to a
specific microenvironment become better
understood. This may lead to yet further
refinement of macrophage nomenclature
as function supersedes form in importance. The next era may well also herald the successful therapeutic manipulation of inflammatory microenvironments
in order to slow or abrogate inflammatory

disease courses in human beings. One can
be certain that the good macrophages vanquishing the bad macrophages will be a
component aspect, and that restoration
of the damaged, ugly tissue will also be
macrophage-dependent.

REFERENCES
1. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill
AM. M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.
4049/jimmunol.164.12.6166
2. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation. Nat Rev Immunol
(2008) 8:958–69. doi:10.1038/nri2448
3. Murray PJ, Allen JE, Biswas SK, Fisher EA,
Gilroy DW, Goerdt S, et al. Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41:14–20.
doi:10.1016/j.immuni.2014.06.008
4. Xue J, Schmidt SV, Sander J, Draffehn A, Krebs
W, Quester I, et al. Transcriptome-based network analysis reveals a spectrum model of
human macrophage activation. Immunity (2014)
40:274–88. doi:10.1016/j.immuni.2014.01.006
5. Vogel DY, Vereyken EJ, Glim JE, Heijnen PD, Moeton M, van der Valk P, et al. Macrophages in inflammatory multiple sclerosis lesions have an intermediate activation status. J Neuroinflammation (2013)
10:35. doi:10.1186/1742-2094-10-35
6. Franklin RJM. Why does remyelination fail in multiple sclerosis? Nat Rev Neurosci (2002) 3:705–14.
doi:10.1038/nrn917
7. Schouppe E, De Baetselier P, Van Ginderachter
JA, Sarukhan A. Instruction of myeloid cells
by the tumor microenvironment: Open questions on the dynamics and plasticity of different tumor-associated myeloid cell populations.
Oncoimmunology (2012) 1:1135–45. doi:10.4161/
onci.21566
8. Lewis CE, Pollard JW. Distinct role of macrophages
in different tumor microenvironments. Cancer Res
(2006) 66:605–12. doi:10.1158/0008-5472.CAN05-4005
9. Movahedi K, Laoui D, Gysemans C, Baeten M,
Stangé G, Van den Bossche J, et al. Different tumor
microenvironments contain functionally distinct
subsets of macrophages derived from Ly6C(high)
monocytes. Cancer Res (2010) 70:5728–39. doi:10.
1158/0008-5472.CAN-09-4672
10. Laoui D, Van Overmeire E, Di Conza G, Aldeni
C, Keirsse J, Morias Y, et al. Tumor hypoxia
does not drive differentiation of tumor-associated
macrophages but rather fine-tunes the M2like macrophage population. Cancer Res (2014)
74:24–30. doi:10.1158/0008-5472.CAN-13-1196
11. Zaynagetdinov R, Sherrill TP, Polosukhin VV, Han
W, Ausborn JA, McLoed AG, et al. A critical
role for macrophages in promotion of urethaneinduced lung carcinogenesis. J Immunol (2011)
187:5703–11. doi:10.4049/jimmunol.1100558
12. Kuang D-M, Wu Y, Chen N, Cheng J, Zhuang
SM, Zheng L. Tumor-derived hyaluronan induces
formation of immunosuppressive macrophages
through transient early activation of monocytes. Blood (2007) 110:587–95. doi:10.1182/
blood-2007-01-068031

December 2014 | Volume 5 | Article 612 | 71

Harris

13. Wang B, Li Q, Qin L, Zhao S, Wang J, Chen X.
Transition of tumor-associated macrophages from
MHC class II(hi) to MHC class II(low) mediates
tumor progression in mice. BMC Immunol (2011)
12:43. doi:10.1186/1471-2172-12-43
14. Stöger JL, Gijbels MJ, van der Velden S, Manca M,
van der Loos CM, Biessen EA, et al. Distribution
of macrophage polarization markers in human
atherosclerosis. Atherosclerosis (2012) 225:461–8.
doi:10.1016/j.atherosclerosis.2012.09.013
15. Redente EF, Higgins DM, Dwyer-Nield LD, Orme
IM, Gonzalez-Juarrero M, Malkinson AM. Differential polarization of alveolar macrophages and
bone marrow-derived monocytes following chemically and pathogen-induced chronic lung inflammation. J Leukoc Biol (2010) 88:159–68. doi:10.
1189/jlb.0609378
16. Ginhoux F, Greter M, Leboeuf M, Nandi S,
See P, Gokhan S, et al. Fate mapping analysis
reveals that adult microglia derive from primitive macrophages. Science (2010) 330:841–5. doi:
10.1126/science.1194637
17. Hoeffel G, Wang Y, Greter M, See P, Teo P,
Malleret B, et al. Adult Langerhans cells derive
predominantly from embryonic fetal liver monocytes with a minor contribution of yolk sac-derived
macrophages. J Exp Med (2012) 209:1167–81.
doi:10.1084/jem.20120340
18. Barth MW, Hendrzak JA, Melnicoff MJ, Morahan
PS. Review of the macrophage disappearance reaction. J Leukoc Biol (1995) 57:361–7.
19. Andersson A, Kokkola R, Wefer J, ErlandssonHarris H, Harris RA. Differential macrophage
expression of IL-12 and IL-23 upon innate
immune activation defines rat autoimmune susceptibility. J Leukoc Biol (2004) 76:1118–24. doi:
10.1189/jlb.0704385
20. Vogel DY, Glim JE, Stavenuiter AW, Breur M, Heijnen P, Amor S, et al. Human macrophage polarization in vitro: maturation and activation methods
compared. Immunobiology (2014) 219:695–703.
doi:10.1016/j.imbio.2014.05.002
21. Verreck FA, de Boer T, Langenberg DM, van
der Ottenhoff TH. Phenotypic and functional

Microenvironmental macrophage functionality

22.

23.

24.

25.

26.

27.

28.

29.

Frontiers in Immunology | Molecular Innate Immunity

profiling of human proinflammatory type-1
and anti-inflammatory type-2 macrophages in
response to microbial antigens and IFN-gammaand CD40L-mediated costimulation. J Leukoc Biol
(2006) 79:285–93. doi:10.1189/jlb.0105015
Parsa R, Andresen P, Gillett A, Mia S, Zhang XM,
Mayans S, et al. Adoptive transfer of immunomodulatory M2 macrophages prevents type 1 diabetes in NOD mice. Diabetes (2012) 61:2881–92.
doi:10.2337/db11-1635
Lee J, French B, Morgan T, French SW. The
liver is populated by a broad spectrum of markers for macrophages. In alcoholic hepatitis the
macrophages are M1 and M2. Exp Mol Pathol
(2014) 96:118–25. doi:10.1016/j.yexmp.2013.09.
004
Morland B, Kaplan G. Macrophage activation
in vivo and in vitro. Exp Cell Res (1977)
108:279–88. doi:10.1016/S0014-4827(77)80035-9
McWhorter FY, Wang T, Nguyen P, Chung T,
Liu WF. Modulation of macrophage phenotype
by cell shape. Proc Natl Acad Sci U S A (2013)
110:17253–8. doi:10.1073/pnas.1308887110
Zhang XM, Lund H, Mia S, Parsa R, Harris RA. Adoptive transfer of cytokine-induced
immunomodulatory adult microglia attenuates
experimental autoimmune encephalomyelitis in
DBA/1 mice. Glia (2014) 62:804–17. doi:10.1002/
glia.22643
Mukherjee S, Rodriguez-Canales J, Hanson J,
Emmert-Buck MR, Tangrea MA, Prieto DA,
et al. Proteomic analysis of frozen tissue samples using laser capture microdissection. Methods
Mol Biol (2013) 1002:71–83. doi:10.1007/978-162703-360-2_6
Bonnet A, Bevilacqua C, Benne F, Bodin L, Cotinot
C, Liaubet L, et al. Transcriptome profiling of
sheep granulosa cells and oocytes during early
follicular development obtained by laser capture
microdissection. BMC Genomics (2011) 12:417.
doi:10.1186/1471-2164-12-417
Mia S, Warnecke A, Zhang XM, Malmström V,
Harris RA. An optimized protocol for human M2
macrophages using M-CSF and IL-4/IL-10/TGF-β

yields a dominant immunosuppressive phenotype.
Scand J Immunol (2014) 79:305–14. doi:10.1111/
sji.12162
30. Shechter R, London A, Varol C, Raposo C,
Cusimano M, Yovel G, et al. Infiltrating bloodderived macrophages are vital cells playing an antiinflammatory role in recovery from spinal cord
injury in mice. PLoS Med (2009) 6(7):e1000113.
doi:10.1371/journal.pmed.1000113
31. Yin Y, Huang X, Lynn KD, Thorpe PE.
Phosphatidylserine-targeting antibody induces
M1 macrophage polarization and promotes
myeloid-derived suppressor cell differentiation.
Cancer Immunol Res (2013) 1:256–68. doi:10.1158/
2326-6066.CIR-13-0073
32. Mills CD, Shearer J, Evans R, Caldwell MD.
Macrophage arginine metabolism and the inhibition or stimulation of cancer. J Immunol (1992)
149:2709–14.

Conflict of Interest Statement: The author declares
that the research was conducted in the absence of any
commercial or financial relationships that could be
construed as a potential conflict of interest.

Received: 09 October 2014; accepted: 13 November 2014;
published online: 03 December 2014.
Citation: Harris RA (2014) Spatial, temporal, and functional aspects of macrophages during “The Good, the
Bad, and the Ugly” phases of inflammation. Front.
Immunol. 5:612. doi: 10.3389/fimmu.2014.00612
This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology.
Copyright © 2014 Harris. This is an open-access article
distributed under the terms of the Creative Commons
Attribution License (CC BY). The use, distribution or
reproduction in other forums is permitted, provided the
original author(s) or licensor are credited and that the
original publication in this journal is cited, in accordance
with accepted academic practice. No use, distribution or
reproduction is permitted which does not comply with
these terms.

December 2014 | Volume 5 | Article 612 | 72

OPINION ARTICLE
published: 07 October 2014
doi: 10.3389/fimmu.2014.00479

“Of mice and men”: arginine metabolism in macrophages
Anita C. Thomas 1 * and Joshua T. Mattila 2
1

Bristol Heart Institute, School of Clinical Sciences, University of Bristol, Bristol, UK
Department of Microbiology and Molecular Genetics, University of Pittsburgh, Pittsburgh, PA, USA
*Correspondence: a.thomas@bristol.ac.uk
2

Edited by:
Charles Dudley Mills, BioMedical Consultants, USA
Reviewed by:
Charles Dudley Mills, BioMedical Consultants, USA
Sidney Morris, University of Pittsburgh, USA
Keywords: arginase, human macrophage, macrophage polarization, nitric oxide synthase, macrophage activation

INTRODUCTION
Macrophages are involved in inflammation
from induction to resolution. Polarization
of macrophages along the M1 (classical) or
M2 (alternative) axis occurs during inflammation and can be at least partly categorized by the route of arginine metabolism within the macrophage, balancing the
activities of the arginase and nitric oxide
synthase (NOS) enzyme families (1, 2).
Arginase activity is associated with tissue
repair responses (via ornithine production
and pro-proliferative effects). In contrast,
NOS2 generates nitric oxide (NO) species
with anti-proliferative effects that is necessary for protection against pathogens and
aberrant cells (2, 3). Other NOS enzymes
produce NO that acts in the regulation
of smooth muscle tone and other cellular
processes (4). Macrophages preferentially
expressing the arginase or NOS2 pathways
enzymes also influence T-cell activation,
proliferation, signaling, and apoptosis in
different ways (1).
While arginase and NOS enzymes can
be used to ascertain the pathway of
macrophage activation in rodents, there
has been debate as to whether they are
present in macrophages from humans and
other mammals. The arginase and NOS
enzymes are extensively conserved, and the
NOS forms found in mammals are similar to those in cnidarians, mollusks, and
other chordates (5, 6). These argininemetabolizing enzymes are present in some
human leukocytes, and there is evidence
that they are also present in macrophages
from other vertebrates, including chickens,
rabbits, cows, and primates (7–12). However, comparisons of tissue macrophages of
different species are lacking, which limits
our understanding (13). Many studies in

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humans have principally focused on blood
monocytes, leading some researchers to
question the suitability of rodents as model
of macrophage activation, as there is not
always a direct correlation with human
cells. Was Robert Koch correct when he
said “Gentlemen, never forget that mice are
not humans,” or can the differing results
between species be explained, in part, by
differences in the types of monocyte or
macrophage studied? Our purpose here is
to examine this question.

ARGININE METABOLISM IN
MAMMALIAN CELLS
Many mammalian cells, including neutrophils, granulocytes, erythrocytes, hepatocytes, cardiac myocytes, dendritic cells,
myeloid-derived suppressor cells, foam
cells, natural killer cells, endothelial cells,
and smooth muscle cells, have arginase
(12, 14–16) or NOS activity (8, 17–19),
albeit to different degrees. Macrophages
are the primary circulating cells that can
express either of these enzymes, depending on the inflammatory circumstance.
Experiments that detect NO, ornithine, or
urea production (via NOS2 or arginase)
have most often been performed on
rodent macrophages. Macrophages from
some mouse strains (e.g., the M1-biased
C57BL/6 strain) can be stimulated by
lipopolysaccharide (LPS) to produce considerable quantities of NO. Macrophages
from others strains (e.g., M2-skewed
BALB/c mice) produce much less NO (20)
and produce more ornithine instead. Some
researchers did not detect any NO production in macrophages from humans, pigs,
and rabbits (8, 11, 14, 21–23), but others
(including ourselves) have observed NOS
or arginase activity in macrophages from

rabbits, humans, and other primates (4, 7,
10, 12, 17, 24–26).

WHY IS THERE CONTROVERSY?
One main difference between the studies from laboratories is that some use
monocyte-derived macrophages (MDM),
while others study tissue macrophages
directly. A number of groups have detected
NOS or arginase activity in human monocytes or macrophages (3, 27–29); but others have not. Why is this so? Part of the
explanation lies in the fact that in vitroderived macrophages can generate different responses from macrophages obtained
in vivo as discussed below (and shown in
Table 1). Another explanation is that many
groups use the identification of enzyme
protein rather than detection of enzyme
activity as evidence of enzyme expression.
Failure to detect the presence of a protein is not definitive evidence for absence
of expression (especially when considering potentially different detection thresholds of antibodies or the high V max of
arginase, i.e., very little enzyme is required
for ornithine production).
MACROPHAGES PRODUCED IN VITRO

Macrophages have been produced in vitro
in a number of ways. Cells from bone marrow have been isolated and “differentiated”
in culture medium containing high levels
of cytokines (such as colony stimulating
factors, CSFs) to produce bone marrowderived macrophages (BMDM) (13, 23, 44–
46). Macrophages have also been produced
by isolating and culturing monocytes from
blood, to produce MDM (10, 13, 22, 30,
37, 47, 48). Production of these in vitroderived macrophages is cheap, simple,
and reproducible, but they may not be a

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Thomas and Mattila

Arginine metabolism in human macrophages

Table 1 | The presence of arginine-metabolizing enzymes in human monocytes and macrophages varies with cell source, treatment and health
status/stress level of the individual.
Cell origin
Blood monocytes

Cell

Treatment

NOS test

ARG test

Result

Monocyte,

0, 2, 3, or 5d culture

RNA, citrulline,

RNA, urea

NOS, ARG1 and ARG2 levels vary

mono-mac
Blood monocytes

Blood monocytes

FC

Reference

Monocyte,

0, 3, or 7d culture or

mono-mac

7d M-CSF, 0.75d

genes with smaller differences

IFNγ/LPS, or IL4

discounted)

Monocyte

1d culture

Monocyte

2d culture

Gene array

RNA

(27)

between monocyte subpopulations
No difference (≤ 2-fold cut-off, therefore

(30)

RNA

↑ARG1, ↓NOS2

(28)

Urea

↑ARG1

(29)

Most donors had low but detectable

(31)

(filaria-infected)
Blood monocytes
(burns victims)
Blood monocytes

Blood monocytes

Monocyte

Mono-mac

2d microfilaria,

RNA

RNA

M-CSF, IL4, or

NOS2 and ARG1 RNA expression which

IFNγ/LPS

did not change with any treatment.

3d IFNγ and/or IL4

RNA

↓ARG1, but detectable in all conditions

(32)

Transcription

No induction of NOS2 gene

(33)

run-on assay

transcription (for that particular region of

(No M-CSF)
Monocyte/macrophage

Mono-mac

?d LPS and/or IFNγ

cell line (U937)

the promoter region)
Monocyte/macrophage

Mono-mac

cell line (U937)

1d selenomethionine Griess, RNA

Selenomethionine ↓LPS-induced NOS2

and 1d LPS and/or

expression (RNA and protein) and nitrite

Western

IFNγ
Blood monocytes,

Mono-mac,

peritoneal macrophages macrophage

(34)

production

?d culture, 2d LPS,

Griess, amino

No nitrite, ornithine, citrulline

IFNγ, or

acid HPLC

production, no arginine consumption

0d or 3d LPS or

RNA, IB, ICC,

NOS2 mRNA and protein present in

cytokine

biopterin,

monocytes, ↑peritoneal macrophages

citrulline, Griess

(↑ with LPS). Both cell types produce

(22)

TNFα/GM-CSF
Blood monocytes,

Monocyte,

peritoneal macrophages mono-mac

(35)

neopterin, nitrite/nitrate and citrulline
(low levels)
Blood monocytes (MS

Macrophage

sufferers)

6d GM-CSF 0.75d

RNA, Griess

IL4, IFNγ, LPS, or

RNA, WB,

ARG1 and NOS2 mRNA and nitrite

urea

production in MS and controls, ↑with

TNFα

(36)

M1 or M2 cytokine challenge. ARG1
protein and urea production present in
controls, ↑in MS

Blood monocytes

Macrophage

Blood monocytes

Macrophage

8d M-CSF, 5d oxLDL

RNA

No change in ARG1 levels

(10)

10d M-CSF, 1d IL4,

Urea, WB

No ARG1 after induction by IL4 or IL10

(14)

or IL10

arginine

Blood monocytes

Macrophage

14d IFNγ/LPS

Griess

No nitrite production

(37)

Alveolar macrophages

Macrophage

IFNγ

Griess, citrulline

No NO production, no effect of NOS

(21)

(volunteers)
Alveolar macrophages

inhibitor
Macrophage

?d (short), 0.8d IL4,

Urea

Untreated macrophages have ARG

or forskolin (i.e.,

activity similar to unstimulated RAW

↑cAMP)

cells. ↑ARG with IL4/forskolin but not

(38)

IL4 alone
Alveolar macrophages

Macrophage

(cancer suffers,

0.75d IFNγ/LPS or
IL-10

RNA, WB

RNA

↑ARG with IL10 stimulated cells,

(39)

↑NOS2 with IFNγ/LPS stimulated cells

volunteers)
(Continued)

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Thomas and Mattila

Arginine metabolism in human macrophages

Table 1 | Continued
Cell origin

Cell

Treatment

NOS test

Alveolar macrophages

Macrophage

None

IHC, WB, RNA,

45–49% of cells from TB patients have

diaphorase

NOS2. Smoking controls had some

(TB patients,

ARG test

volunteers)

Result

Reference
(24)

NOS2-positive macrophages,
non-smoking controls have few
NOS2-positive cells

Alveolar macrophages

Macrophage

None

IHC

Macrophages in TB granulomas stain

(TB patients)

(26)

for NOS1, NOS2 and nitrotyrosine (i.e.,
active)

Alveolar macrophages

Macrophage

None

IHC

IHC

(TB patients)

ARG1 in macrophages in TB

(12)

granulomas, few have Arg2. Some
macrophages on outer margins have
both NOS2 and ARG1, some near
center have NOS2, NOS3 and ARG1

Atherosclerotic plaque

Macrophage

None

ISH, IHC

NOS2 in macrophages and smooth

macrophages

(7)

muscle cells, co-localized with oxidized
lipoproteins and peroxynitrite (i.e.,
NOS is active)

Atherosclerotic plaque

Macrophage

None

IHC, WB

Fatty streaks: no NOS2. Advanced

macrophages

(25)

plaques: NOS2 present in
macrophages near necrotic core,
associated with ceroid accumulation
and nitrotyrosine (i.e., active)

Atherosclerotic plaque

Macrophage

None

IHC

foamy macrophages

↑ARG1 in macrophages in superficial

(10)

layers, ↓ARG1 in macrophages
surrounding lipid core

Atherosclerotic plaque

Macrophage

None

IHC, ISH

NOS2 and nitrotyrosine localized to

(17)

smooth muscle cells, macrophages
and foam cells (i.e., active)
Oral macrophages

Macrophage

None

IHC, nitrate

NOS2 present in macrophages from

(40)

gingivitis samples
Placental macrophages

Macrophage

None

Skin macrophages

Macrophage

None

IHC, HPLC

(wound)

Tumor-associated

Macrophage

None

FC

Some M2 macrophages have ARG1

(16)

IHC, WB,

NOS2 present in macrophages, some

(41)

ELISA,

have ARG2, but none have ARG1.

HPLC

Controls: no ARG2

IHC

macrophages

NOS2 present in some macrophages

(42)

(bladder)

While changes in RNA expression of arginine-metabolizing enzymes have been used to identify macrophage activation states, protein changes [such as western
blotting (WB) or immunohistochemistry (IHC)] are also useful. Nitric oxide synthase (NOS) activity can be assessed directly [e.g., production of citrulline or NO (e.g.,
Griess assay)] or by the presence of markers of NO production (such as peroxynitrite, nitrotyrosine or ceroid, a complex of oxidized lipids and proteins). Arginase
(ARG) activity can be measured as urea or ornithine production (e.g., urea assays, amino acid HPLC).
d, number of days; ?d, unspecified number of days; FC, flow cytometry; M-CSF, macrophage colony-stimulating factor; IFNγ, interferon-γ; LPS, lipopolysaccharide;
IL, interleukin; TNFα, tumor necrosis factor-α; GM-CSF, granulocyte-macrophage colony-stimulating factor; IB, immunoblot; ICC, immunocytochemistry; MS, multiple
sclerosis; oxLDL, oxidized low density lipoprotein; TB, tuberculosis; ISH, in situ hybridization; Griess, Griess assay for nitrite/nitrate production.
It should be noted that NO production below the detection levels of this relatively insensitive assay may still have functional effects (43).

full representative of tissue macrophages,
as the preparation and culture procedures may not be sufficient to induce
cell activation (4). The differences between
tissue macrophages and in vitro-derived

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macrophages are at least partly dependent
on cell source, time in culture, and the
degree of manipulation in culture. Each
research group will use different types and
sources of culture media and sera, which

vary greatly in the concentrations of factors
that influence NOS2 or arginase expression, such as transforming growth factor
β (TGFβ) (4, 20, 49). Another confounding issue is that circulating monocytes and

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Thomas and Mattila

tissue macrophages arise from different
stem cell populations (50), although some
macrophages found at sites of infection or
inflammation may derive from infiltrating monocytes (51). Together, these factors
may account for many of the differences
observed in NO and urea production in
these macrophages (8, 20).
Monocyte-derived macrophages or
BMDM from different strains of mice
can differ in their response to interferon-γ
(IFNγ), LPS, and tumor necrosis factor-α
(TNFα) (4, 8), and differences in the rodent
background can result in differences in
macrophage gene expression (13, 20, 49).
Human in vitro-derived macrophages also
show variability in their responses to LPS
(4, 22, 46). It may be that the same stimulus
is able to generate quite different responses
in genetically diverse individuals, as it does
between mouse strains (38, 49, 52). In
general, human macrophages are not as
responsive to LPS as mouse macrophages,
possibly because of the lower environmental exposure of humans to LPS. It is also
possible that human monocytes may be
more effectively stimulated to become M1activated macrophages by cytokines other
than IFNγ and LPS/TNFα (e.g., IFNα) (4,
18, 43). Human macrophages take longer
time to respond to the stimulatory factors
in vitro than mouse macrophages, and
some experiments using human MDM
may have ended before a response was
detected (48). There are other indications
that the timing and length of the exposure
of the cells to varying cytokines in vitro
are important. For example, when M1polarizing cytokines were removed from
the culture medium, NOS2 levels in mouse
BMDM were reduced and NO production
(measured as nitrite) ceased (45). In addition, whichever arginase or NOS enzyme
was induced earliest, the alternative
enzyme decreased in expression and activity, unless arginine was present in excess
(15, 45, 53). Macrophages require the local
environment to continuously give appropriate activation cues. Changes in environmental cues can stimulate macrophage
populations in vitro to express varying percentages of M1 or M2 dominant activity
(54). When activation cues are reduced or
removed, macrophages may become deactivated (e.g., M2c) or indeterminate (e.g.,
have features of M1 and M2).

Frontiers in Immunology | Inflammation

Arginine metabolism in human macrophages

MACROPHAGES OBTAINED IN VIVO

Macrophages can be identified in whole
tissues and organs or isolated in large
numbers from in vivo sources such as
the peritoneum or granulomas, and either
examined immediately or used ex vivo.
Macrophages obtained in vivo or made
from monocytes can respond differently to
the same stimulus (35, 47). In one study,
monocytes and tissue macrophages were
obtained from patients with an inflammatory disease (either rheumatoid or psoriatic arthritis). Compared with tissue
macrophages, the MDM had a blunted
response to the M2 cytokines interleukin4 (IL-4) and IL-13, at least partly due to a
reduction in some of the receptor elements
for these cytokines (47). These results suggest that the response of the macrophages
to M2 cytokines may be source specific,
but it is possible that these cytokines
alone were not sufficient to fully stimulate the MDM (38). Several lines of evidence suggest that macrophages in vivo
express functional NOS2. Blood monocytes and peritoneal macrophages obtained
from women during laparoscopic procedures contained NOS2 mRNA and protein. The macrophages had higher NOS
levels than the monocytes, and this could
be increased by treatment with LPS. The
monocytes and macrophages also produced neopterin, nitrite/nitrate, and citrulline (suggesting that the enzyme was
active). Although the production of NO
from these macrophages was low, it would
probably have been sufficient to cause
functional changes (35).
Macrophages can also be obtained from
alveolar aspirates, skin, and the placenta
(10, 16, 21, 38, 39, 55, 56). For example, sponges placed subcutaneously into
mice, rats, or rabbits attract large numbers of macrophages. The sponges can
be removed from the animal and the
macrophages were isolated and purified
(10, 55, 56). It is a little more difficult
to obtain and purify macrophages from
other tissues, such as atherosclerotic vessels
(44), but intact biopsy, surgical, or cadaveric specimens can also be investigated. It
should be noted that resident macrophages
from different tissues observed at different times (and different health states)
may not necessarily have identical properties (51, 57).

In order to perform their full range of
functions, macrophage populations exhibit
“plasticity” of phenotype (52, 58), regardless of whether they are found in vivo or
derived in vitro. As macrophages adapt or
change their functions, they can simultaneously express markers of M1 and M2 activation, including NOS2 and arginase-1 (12,
59, 60). For example, tissue macrophages
(and MDM) from Mycobacterium tuberculosis-infected cynomolgus macaques have
been observed to co-express functional
NOS and arginase enzymes (12). We suggest that macrophages display a spectrum
of activation phenotypes, and it is the relative (and not absolute) proportion of M1 or
M2 markers that we can use as a ‘handle’ to
determine the type of activation state.
EFFECT OF DISEASE AND TRAUMA ON
MACROPHAGE ACTIVATION

Blood monocytes from healthy volunteers
do not usually need to produce NOS
or arginase, so it is not surprising that
many studies have not detected NOS or
arginase in these cells (10, 14, 21, 22,
29, 30, 37). However, studies performed
on tissue or cells from people undergoing stress, trauma [e.g., burns (29)], pregnancy (16), or disease {such as infection
[e.g., tuberculosis (12, 24, 26) or filarial
infection (28)], atherosclerosis (7, 10, 17,
25), autoimmune diseases (27, 36) and
cancer (42, 61)} demonstrate that human
macrophages (and sometimes monocytes)
can produce active forms of the argininemetabolizing enzymes (Table 1).
Trauma results in a pattern of gene
expression in macrophages that is consistent with a wound-healing response, with
an initial increase in NOS followed by
decreased NOS production and activity,
elevated IL-4, IL-10, and TGFβ levels, and
increased arginase expression and activity, resulting in decreased plasma arginine
levels (28, 29, 62).
Disease, however, causes different patterns of gene expression. For example,
monocytes from multiple sclerosis sufferers not only have higher levels of arginase1 and increased urea production, but
also have increased NOS2 mRNA and
nitrite production (particularly when stimulated by M1 cytokines or LPS) (36).
Macrophages from patients with inflammatory diseases, such as tuberculosis,

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Thomas and Mattila

malaria, or rheumatoid arthritis, have
increased levels of NOS2 mRNA and active
protein (4, 8, 24, 26, 63), which may contribute to elevated plasma NO levels (64).
Atherosclerosis is another inflammatory
disease with a considerable macrophage
contribution, with oxidized low-density
lipoproteins taken up by macrophages
during their transformation into foam
cells. Plaque macrophages express NOS2
RNA and protein, as well as markers
of NOS activity (including the presence
of nitrotyrosine or ceroid) (4, 7, 17,
25). Plaque macrophages and foam cells
express arginase-1 (10), and macrophages
laser-dissected from plaque have upregulated levels of arginase-2 and NOS2 (65).
Macrophages present in some neoplastic diseases also produce active NOS2 (4,
42, 66). Reducing the local levels of arginine has been proposed as a treatment for
these diseases, by reducing inflammationtriggered immune dysfunction, tumor
escape, fibrosis, and immunosuppression
(61). Possible pharmacological interventions include treatment with arginine
degrading enzymes, NOS competitors and
inhibitors, asymmetric dimethylarginine,
NO-releasing aspirins, cyclooxygenase, and
phosphodiesterase or arginase inhibitors
(8, 61). These studies suggest that an inflammatory environment is necessary in order to
observe NOS or arginase in human monocytes and macrophages. The in vitro experiments that do not demonstrate arginase or
NOS expression may simply be lacking the
additional cues needed for expression rather
than demonstrating an inability to actually
express these factors.

CONCLUSION
The modulation of macrophages to express
NOS or arginase has clear benefits for treating disease in humans (and other species).
To do this, one needs to either determine
suitable signals to stimulate these pathways
or obtain a sufficient number of human
macrophages (e.g., by tissue culture) that
function like tissue macrophages.
Because macrophages from different
inbred strains of mice vary greatly in
their macrophage NOS and arginase balance, one would predict similar variability to be found in humans as well. In
addition, the source of the macrophages
being studied has been found to be important. Several groups have reported that

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Arginine metabolism in human macrophages

human monocytes from healthy volunteers that have been differentiated or
manipulated in vitro using current protocols tend not to have detectable levels
of arginase and NOS enzymes, whereas
MDM from diseased or stressed individuals or tissue macrophages obtained
from normal, diseased, or stressed individuals do express NOS and/or arginase.
Together these observations suggest that
the current system of differentiating
macrophages from human peripheral
monocytes in vitro needs further refinement before it can be considered to be
an accurate model of human macrophage
behavior in vivo (63). In turn, we need
to understand the differences and similarities between the different species and
the cells being studied to develop experimental models that will answer some
of the outstanding questions regarding
macrophage M1/M2 or other activation
states: What regulates macrophage activation in tissues? What mechanisms regulate macrophage plasticity and stability? How does plasticity of phenotype
affect tissue macrophages? What are the
full in vivo ramifications of the M1/M2
paradigm?
Further work is important to be sure
that our observations of the human system in vitro are real, and not due to
our cell source, measurements, or manipulations. We suggest that macrophages
obtained from mice remain useful for
investigating aspects of these questions in humans/human macrophages. So,
although mice are not men (as Robert
Koch observed), we agree with Rudolf Virchow that “Between animal and human
medicine there is no dividing line – nor
should there be. The object is different
but the experience obtained constitutes the
basis of all medicine” [Rudolph Virchow,
1821–1902].

2.

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

ACKNOWLEDGMENTS
Anita C. Thomas is supported by funding from the British Heart Foundation.
Joshua T. Mattila is supported in part by
NIH RO1A103785 and Bill and Melinda
Gates Foundation grants to JoAnne Flynn
(University of Pittsburgh).

13.

14.

REFERENCES
1. Gallina G, Dolcetti L, Serafini P, De Santo C, Marigo
I, Colombo MP, et al. Tumors induce a subset of

15.

inflammatory monocytes with immunosuppressive activity on CD8+ T cells. J Clin Invest (2006)
116:2777–90. doi:10.1172/JCI28828
Lumeng CN, Bodzin JL, Saltiel AR. Obesity induces
a phenotypic switch in adipose tissue macrophage
polarization. J Clin Invest (2007) 117:175–84. doi:
10.1172/JCI29881
Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev Immunol
(2012) 32:463–88. doi:10.1615/CritRevImmunol.
v32.i6.10
Weinberg JB. Nitric oxide production and nitric
oxide synthase type 2 expression by human
mononuclear phagocytes: a review. Mol Med
(1998) 4:557–91.
Perozich J, Hempel J, Morris J. Roles of conserved
residues in the arginase family. Biochim Biophys
Acta (1998) 1382:23–37.
Dzik JM. The ancestry and cumulative evolution
of immune reactions. Acta Biochim Pol (2010)
57:443–66.
Luoma JS, Stralin P, Marklund SL, Hiltunen
TP, Sarkioja T, Yla-Herttuala S. Expression of
extracellular SOD and iNOS in macrophages
and smooth muscle cells in human and
rabbit atherosclerotic lesions: colocalization
with epitopes characteristic of oxidized LDL
and peroxynitrite-modified proteins. Arterioscler Thromb Vasc Biol (1998) 18:157–67.
doi:10.1161/01.ATV.18.2.157
Bogdan C. Nitric oxide and the immune response.
Nat Immunol (2001) 2:907–16. doi:10.1038/
ni1001-907
Djeraba A, Musset E, van RN, Quere P. Resistance
and susceptibility to Marek’s disease: nitric oxide
synthase/arginase activity balance. Vet Microbiol
(2002) 86:229–44. doi:10.1016/S0378-1135(02)
00010-X
Thomas AC, Sala-Newby GB, Ismail Y, Johnson JL, Pasterkamp G, Newby AC. Genomics of
foam cells and nonfoamy macrophages from rabbits identifies arginase-I as a differential regulator of nitric oxide production. Arterioscler Thromb
Vasc Biol (2007) 27:571–7. doi:10.1161/01.ATV.
0000256470.23842.94
Zelnickova P, Matiasovic J, Pavlova B, Kudlackova H, Kovaru F, Faldyna M. Quantitative nitric
oxide production by rat, bovine and porcine
macrophages. Nitric Oxide (2008) 19:36–41. doi:
10.1016/j.niox.2008.04.001
Mattila JT, Ojo OO, Kepka-Lenhart D, Marino
S, Kim JH, Eum SY, et al. Microenvironments in
tuberculous granulomas are delineated by distinct
populations of macrophage subsets and expression of nitric oxide synthase and arginase isoforms. J Immunol (2013) 191:773–84. doi:10.4049/
jimmunol.1300113
Murray PJ, Wynn TA. Obstacles and opportunities for understanding macrophage polarization.
J Leukoc Biol (2011) 89:557–63. doi:10.1189/jlb.
0710409
Munder M, Mollinedo F, Calafat J, Canchado
J, Gil-Lamaignere C, Fuentes JM, et al. Arginase
I is constitutively expressed in human granulocytes and participates in fungicidal activity. Blood
(2005) 105:2549–56. doi:10.1182/blood-2004-072521
Morris SM. Arginine metabolism: boundaries of
our knowledge. J Nutr (2007) 137:1602S–9S.

October 2014 | Volume 5 | Article 479 | 77

Thomas and Mattila

16. Kropf P, Baud D, Marshall SE, Munder M, Mosley
A, Fuentes JM, et al. Arginase activity mediates
reversible T cell hyporesponsiveness in human
pregnancy. Eur J Immunol (2007) 37:935–45. doi:
10.1002/eji.200636542
17. Buttery LD, Springall DR, Chester AH, Evans TJ,
Standfield EN, Parums DV, et al. Inducible nitric
oxide synthase is present within human atherosclerotic lesions and promotes the formation
and activity of peroxynitrite. Lab Invest (1996)
75:77–85.
18. Bronte V, Zanovello P. Regulation of immune
responses by l-arginine metabolism. Nat Rev
Immunol (2005) 5:641–54. doi:10.1038/nri1668
19. Lechner MG, Megiel C, Russell SM, Bingham
B, Arger N, Woo T, et al. Functional characterization of human Cd33+ and Cd11b+ myeloidderived suppressor cell subsets induced from
peripheral blood mononuclear cells co-cultured
with a diverse set of human tumor cell lines.
J Transl Med (2011) 9:90. doi:10.1186/1479-58769-90
20. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill
AM. M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.
4049/jimmunol.164.12.6166
21. Cameron ML, Granger DL, Weinberg JB, Kozumbo
WJ, Koren HS. Human alveolar and peritoneal
macrophages mediate fungistasis independently of
l-arginine oxidation to nitrite or nitrate. Am Rev
Respir Dis (1990) 142:1313–9. doi:10.1164/ajrccm/
142.6_Pt_1.1313
22. Schneemann M, Schoedon G, Hofer S, Blau N,
Guerrero L, Schaffner A. Nitric oxide synthase
is not a constituent of the antimicrobial armature of human mononuclear phagocytes. J Infect
Dis (1993) 167:1358–63. doi:10.1093/infdis/167.6.
1358
23. Kapetanovic R, Fairbairn L, Beraldi D, Sester DP, Archibald AL, Tuggle CK, et al. Pig
bone marrow-derived macrophages resemble
human macrophages in their response to bacterial lipopolysaccharide. J Immunol (2012)
188:3382–94. doi:10.4049/jimmunol.1102649
24. Nicholson S, Bonecini-Almeida MG, Lapa e
Silva JL, Nathan C, Xie QW, Mumford R, et al.
Inducible nitric oxide synthase in pulmonary alveolar macrophages from patients with tuberculosis.
J Exp Med (1996) 183:2293–302. doi:10.1084/jem.
183.5.2293
25. Cromheeke KM, Kockx MM, De Meyer GRY,
Bosmans JM, Bult H, Beelaerts WJF, et al. Inducible
nitric oxide synthase colocalizes with signs of
lipid oxidation/peroxidation in human atherosclerotic plaques. Cardiovasc Res (1999) 43:744–54.
doi:10.1016/S0008-6363(99)00148-0
26. Choi HS, Rai PR, Chu HW, Cool C, Chan ED.
Analysis of nitric oxide synthase and nitrotyrosine expression in human pulmonary tuberculosis. Am J Respir Crit Care Med (2002) 166:178–86.
doi:10.1164/rccm.2201023
27. Rouzaut A, Subira ML, de Miguel C, Domingode-Miguel E, Gonzalez A, Santiago E, et al. Coexpression of inducible nitric oxide synthase and
arginases in different human monocyte subsets.
Apoptosis regulated by endogenous NO. Biochim
Biophys Acta (1999) 1451:319–33. doi:10.1016/
S0167-4889(99)00106-8

Frontiers in Immunology | Inflammation

Arginine metabolism in human macrophages

28. Babu S, Kumaraswami V, Nutman TB. Alternatively activated and immunoregulatory monocytes
in human filarial infections. J Infect Dis (2009)
199:1827–37. doi:10.1086/599090
29. Kobayashi M, Jeschke MG, Shigematsu K, Asai
A, Yoshida S, Herndon DN, et al. M2b monocytes predominated in peripheral blood of severely
burned patients. J Immunol (2010) 185:7174–9.
doi:10.4049/jimmunol.0903935
30. Martinez FO, Gordon S, Locati M, Mantovani A.
Transcriptional profiling of the human monocyteto-macrophage differentiation and polarization:
new molecules and patterns of gene expression. J Immunol (2006) 177:7303–11. doi:10.4049/
jimmunol.177.10.7303
31. Semnani RT, Mahapatra L, Moore V, Sanprasert
V, Nutman TB. Functional and phenotypic characteristics of alternative activation induced in
human monocytes by interleukin-4 or the parasitic nematode Brugia malayi. Infect Immun (2011)
79:3957–65. doi:10.1128/IAI.05191-11
32. Raes G, Brys L, Dahal BK, Brandt J, Grooten J,
Brombacher F, et al. Macrophage galactose-type
C-type lectins as novel markers for alternatively
activated macrophages elicited by parasitic infections and allergic airway inflammation. J Leukoc
Biol (2005) 77:321–7. doi:10.1189/jlb.0304212
33. Zhang X, Laubach VE, Alley EW, Edwards
KA, Sherman PA, Russell SW, et al. Transcriptional basis for hyporesponsiveness of the
human inducible nitric oxide synthase gene
to lipopolysaccharide/interferon-gamma. J Leukoc
Biol (1996) 59:575–85.
34. Shen Y,Yang S, Shi Z, Lin T, Zhu H, Bi F, et al. SeMet
mediates anti-inflammation in LPS-induced U937
cells targeting NF-kB signaling pathway. Inflammation (2014). doi:10.1007/s10753-014-9984-0
35. Weinberg JB, Misukonis MA, Shami PJ, Mason SN,
Sauls DL, Dittman WA, et al. Human mononuclear phagocyte inducible nitric oxide synthase
(iNOS): analysis of iNOS mRNA, iNOS protein,
biopterin, and nitric oxide production by blood
monocytes and peritoneal macrophages. Blood
(1995) 86:1184–95.
36. Christophi GP, Panos M, Hudson CA, Christophi
RL, Gruber RC, Mersich AT, et al. Macrophages of
multiple sclerosis patients display deficient SHP1 expression and enhanced inflammatory phenotype. Lab Invest (2009) 89:742–59. doi:10.1038/
labinvest.2009.32
37. Padgett EL, Pruett SB. Evaluation of nitrite production by human monocyte-derived macrophages.
Biochem Biophys Res Commun (1992) 186:775–81.
doi:10.1016/0006-291X(92)90813-Z
38. Erdely A, Kepka-Lenhart D, Clark M, ZeidlerErdely P, Poljakovic M, Calhoun WJ, et al. Inhibition of phosphodiesterase 4 amplifies cytokinedependent induction of arginase in macrophages.
Am J Physiol Lung Cell Mol Physiol (2006)
290:L534–9. doi:10.1152/ajplung.00326.2005
39. Liu CY, Wang YM, Wang CL, Feng PH, Ko HW,
Liu YH, et al. Population alterations of l-arginaseand inducible nitric oxide synthase-expressed
CD11b+/CD14(-)/CD15+/CD33+
myeloidderived suppressor cells and CD8+ T lymphocytes
in patients with advanced-stage non-small cell
lung cancer. J Cancer Res Clin Oncol (2010)
136:35–45. doi:10.1007/s00432-009-0634-0

40. Ozer L, Elgun S, Ozdemir B, Pervane B, Ozmeric
N. Arginine-nitric oxide-polyamine metabolism in
periodontal disease. J Periodontol (2011) 82:320–8.
doi:10.1902/jop.2010.100199
41. Debats IB, Wolfs TG, Gotoh T, Cleutjens JP,
Peutz-Kootstra CJ, van der Hulst RR. Role of
arginine in superficial wound healing in man.
Nitric Oxide (2009) 21:175–83. doi:10.1016/j.niox.
2009.07.006
42. Suriano F, Santini D, Perrone G,Amato M,Vincenzi
B, Tonini G, et al. Tumor associated macrophages
polarization dictates the efficacy of BCG instillation in non-muscle invasive urothelial bladder cancer. J Exp Clin Cancer Res (2013) 32:87. doi:10.
1186/1756-9966-32-87
43. Bogdan C. Response to ‘Species differences in
macrophage NO production are important’. Nat
Immunol (2002) 3:102. doi:10.1038/ni0202-102a
44. Galkina E, Kadl A, Sanders J, Varughese D, Sarembock IJ, Ley K. Lymphocyte recruitment into the
aortic wall before and during development of
atherosclerosis is partially l-selectin dependent.
J Exp Med (2006) 203:1273–82. doi:10.1084/jem.
20052205
45. Yeramian A, Martin L, Serrat N, Arpa L, Soler
C, Bertran J, et al. Arginine transport via cationic
amino acid transporter 2 plays a critical regulatory role in classical or alternative activation
of macrophages. J Immunol (2006) 176:5918–24.
doi:10.4049/jimmunol.176.10.5918
46. Shalhoub J, Falck-Hansen MA, Davies AH,
Monaco C. Innate immunity and monocytemacrophage activation in atherosclerosis. J
Inflamm (2011) 8:9. doi:10.1186/1476-9255-8-9
47. Hart PH, Bonder CS, Balogh J, Dickensheets HL,
Donnelly RP, Finlay-Jones JJ. Differential responses
of human monocytes and macrophages to IL-4 and
IL-13. J Leukoc Biol (1999) 66:575–8.
48. Geelhaar-Karsch A, Schinnerling K, Conrad K,
Friebel J, Allers K, Schneider T, et al. Evaluation
of arginine metabolism for the analysis of M1/M2
macrophage activation in human clinical specimens. Inflamm Res (2013) 62:865–9. doi:10.1007/
s00011-013-0642-z
49. Mills CD. Macrophage arginine metabolism to
ornithine/urea or nitric oxide/citrulline: a life or
death issue. Crit Rev Immunol (2001) 21:399–425.
doi:10.1615/CritRevImmunol.v21.i5.10
50. Epelman S, Lavine KJ, Randolph GJ. Origin and
functions of tissue macrophages. Immunity (2014)
41:21–35. doi:10.1016/j.immuni.2014.06.013
51. Hussell T, Bell TJ. Alveolar macrophages: plasticity in a tissue-specific context. Nat Rev Immunol
(2014) 14:81–93. doi:10.1038/nri3600
52. Ravasi T, Wells CA, Hume DA. Systems biology
of transcription control in macrophages. Bioessays
(2007) 29:1215–26. doi:10.1002/bies.20683
53. El Gayar S, Thuring-Nahler H, Pfeilschifter J,
Rollinghoff M, Bogdan C. Translational control of inducible nitric oxide synthase by IL13 and arginine availability in inflammatory
macrophages. J Immunol (2003) 171:4561–8. doi:
10.4049/jimmunol.171.9.4561
54. Khallou-Laschet J, Varthaman A, Fornasa G, Compain C, Gaston AT, Clement M, et al. Macrophage
plasticity in experimental atherosclerosis. PLoS
One (2010) 5:e8852. doi:10.1371/journal.pone.
0008852

October 2014 | Volume 5 | Article 479 | 78

Thomas and Mattila

55. Albina JE, Mills CD, Barbul A, Thirkill CE, Henry
WL, Mastrofrancesco B, et al. Arginine metabolism in wounds. Am J Physiol (1988) 254:
E459–67.
56. Daley JM, Brancato SK, Thomay AA, Reichner
JS, Albina JE. The phenotype of murine wound
macrophages. J Leukoc Biol (2010) 87:59–67. doi:
10.1189/jlb.0409236
57. Johnson JL, Newby AC. Macrophage heterogeneity in atherosclerotic plaques. Curr Opin
Lipidol (2009) 20(5):370–8. doi:10.1097/MOL.
0b013e3283309848
58. Hume DA. Plenary perspective: the complexity
of constitutive and inducible gene expression in
mononuclear phagocytes. J Leukoc Biol (2012)
92:433–44. doi:10.1189/jlb.0312166
59. Poljakovic M, Porter DW, Millecchia L, KepkaLenhart D, Beighley C, Wolfarth MG, et al. Celland isoform-specific increases in arginase expression in acute silica-induced pulmonary inflammation. J Toxicol Environ Health A (2007) 70:118–27.
doi:10.1080/15287390600755075
60. Arora S, Olszewski MA, Tsang TM, McDonald
RA, Toews GB, Huffnagle GB. Effect of cytokine
interplay on macrophage polarization during
chronic pulmonary infection with Cryptococcus
neoformans. Infect Immun (2011) 79:1915–26. doi:
10.1128/IAI.01270-10

www.frontiersin.org

Arginine metabolism in human macrophages

61. Munder M. Arginase: an emerging key player in
the mammalian immune system. Br J Pharmacol
(2009) 158:638–51. doi:10.1111/j.1476-5381.2009.
00291.x
62. Ochoa JB, Bernard AC, O’Brien WE, Griffen MM,
Maley ME, Rockich AK, et al. Arginase I expression and activity in human mononuclear cells after
injury. Ann Surg (2001) 233:393–9. doi:10.1097/
00000658-200103000-00014
63. Nathan C. Role of iNOS in human host defense.
Science (2006) 312:1874–5. doi:10.1126/science.
312.5782.1874b
64. Boutlis CS, Tjitra E, Maniboey H, Misukonis MA,
Saunders JR, Suprianto S, et al. Nitric oxide production and mononuclear cell nitric oxide synthase activity in malaria-tolerant papuan adults.
Infect Immun (2003) 71:3682–9. doi:10.1128/IAI.
71.7.3682-3689.2003
65. Tuomisto TT, Korkeela A, Rutanen J, Viita H,
Brasen JH, Riekkinen MS, et al. Gene expression
in macrophage-rich inflammatory cell infiltrates
in human atherosclerotic lesions as studied by
laser microdissection and DNA array: overexpression of HMG-CoA reductase, colony stimulating factor receptors, CD11A/CD18 integrins,
and interleukin receptors. Arterioscler Thromb
Vasc Biol (2003) 23:2235–40. doi:10.1161/01.ATV.
0000102551.91154.96

66. Pantano F, Berti P, Guida FM, Perrone G, Vincenzi B, Amato MM, et al. The role of macrophages
polarization in predicting prognosis of radically
resected gastric cancer patients. J Cell Mol Med
(2013) 17:1415–21. doi:10.1111/jcmm.12109

Conflict of Interest Statement: The authors declare
that the research was conducted in the absence of any
commercial or financial relationships that could be
construed as a potential conflict of interest.
Received: 01 August 2014; accepted: 19 September 2014;
published online: 07 October 2014.
Citation: Thomas AC and Mattila JT (2014) “Of mice
and men”: arginine metabolism in macrophages. Front.
Immunol. 5:479. doi: 10.3389/fimmu.2014.00479
This article was submitted to Inflammation, a section of
the journal Frontiers in Immunology.
Copyright © 2014 Thomas and Mattila. This is an openaccess article distributed under the terms of the Creative
Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted,
provided the original author(s) or licensor are credited
and that the original publication in this journal is cited,
in accordance with accepted academic practice. No use,
distribution or reproduction is permitted which does not
comply with these terms.

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REVIEW ARTICLE
published: 04 November 2014
doi: 10.3389/fimmu.2014.00510

Rethinking regenerative medicine: a macrophage-centered
approach
Bryan N. Brown 1,2 † , Brian M. Sicari 1,3 † and Stephen F. Badylak 1,3 *
1
2
3

McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA, USA
Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA, USA
Department of Surgery, University of Pittsburgh, Pittsburgh, PA, USA

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Fulvio D’Acquisto, Queen Mary
University of London, UK
Manuela Mengozzi, Brighton and
Sussex Medical School, UK
*Correspondence:
Stephen F. Badylak , McGowan
Institute for Regenerative Medicine,
Suite 300, 450 Technology Drive,
Pittsburgh, PA 15219, USA
e-mail: badylaks@upmc.edu
†

Bryan N. Brown and Brian M. Sicari
have contributed equally to this work.

Regenerative medicine, a multi-disciplinary approach that seeks to restore form and function to damaged or diseased tissues and organs, has evolved significantly during the past
decade. By adapting and integrating fundamental knowledge from cell biology, polymer science, and engineering, coupled with an increasing understanding of the mechanisms which
underlie the pathogenesis of specific diseases, regenerative medicine has the potential for
innovative and transformative therapies for heretofore unmet medical needs. However, the
translation of novel technologies from the benchtop to animal models and clinical settings is
non-trivial and requires an understanding of the mechanisms by which the host will respond
to these novel therapeutic approaches. The role of the innate immune system, especially
the role of macrophages, in the host response to regenerative medicine based strategies
has recently received considerable attention. Macrophage phenotype and function have
been suggested as critical and determinant factors in downstream outcomes. The constructive and regulatory, and in fact essential, role of macrophages in positive outcomes
represents a significant departure from the classical paradigms of host–biomaterial interactions, which typically consider activation of the host immune system as a detrimental event.
It appears desirable that emerging regenerative medicine approaches should not only
accommodate but also promote the involvement of the immune system to facilitate positive outcomes. Herein, we describe the current understanding of macrophage phenotype
as it pertains to regenerative medicine and suggest that improvement of our understanding of context-dependent macrophage polarization will lead to concurrent improvement in
outcomes.
Keywords: regenerative medicine, biomaterials, host response, foreign-body reaction, stem cells, macrophages

INTRODUCTION
The macrophage has long been known to play an important role in
the tissue remodeling response which occurs following injury. In
brief, macrophages arrive at the site of tissue injury 24–48 h postinjury, serve as phagocytes clearing the wound bed and initiating
the processes that lead to the default outcome of scar tissue formation (1, 2). However, only recently it has been recognized that
macrophages can have positive impacts upon tissue remodeling
following injury (3–9). While the specific mechanisms by which
macrophages direct tissue remodeling responses remain a subject
of ongoing research, it has been suggested that a transition from a
pro-inflammatory (M1) phenotype to a more regulatory or antiinflammatory M2 phenotype is a key aspect of tissue remodeling
which promotes functional outcomes as opposed to scar tissue
formation.
A correlation of macrophage phenotype with functional
recovery in wound healing has been suggested for more than
two decades (9). With the introduction and general acceptance of the M1/M2 phenotypic dichotomy (10), correlation of
macrophage polarization states and functional recovery has now
been reported in several other tissues and organ systems and

Frontiers in Immunology | Inflammation

represents an area of increasing interest for those in the area
of wound healing and regeneration. The central dogma of this
macrophage-centered approach is that treatments which facilitate an efficient and timely switch from a pro-inflammatory
to an anti-inflammatory and regulatory phenotype, will logically promote functional tissue remodeling over scar tissue
formation.
The M1/M2 paradigm has been widely studied in the context
of disease pathogenesis, particularly cancer, for more than two
decades (11–13). The participation of M1 and M2 macrophages
in a diverse set of diseases including atherosclerosis, endometriosis, and pulmonary fibrosis is also now recognized (2, 12, 14–18).
In addition, there is evidence for the importance of macrophages
in tissue and organ development and in processes such as limb
regeneration in the axolotl (19, 20). Loss of macrophages during
these processes leads to defects in development or retardation of
the regenerative process in the axolotl. Similar findings have been
reported in other regenerative species including zebrafish, where
ablation of macrophages results in defects in fin regeneration following injury (21). This ability to promote a regenerative response
is lost in higher order species with increasing complexity of the

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Brown et al.

immune system, having been replaced with a default mechanism
of “rapid resolution” (i.e., scarring). While the mechanisms which
underlie the loss of regenerative potential remain largely unknown,
a better understanding of the role of the innate immune system in
the regenerative process of lower organisms may provide targets
for regeneration strategies in humans (22).
Regenerative medicine approaches to tissue reconstruction or
organ replacement seek to restore the form and function of lost,
damaged, or diseased tissues. These approaches logically rely upon
our understanding of wound healing, development, and regeneration as guideposts for design. These approaches may incorporate one or more biomaterials, biologically active molecules,
and/or cell sources. Recent advances in these areas have enabled
highly innovative and promising therapies, but translation of such
strategies, without exception, requires in depth investigation and
understanding of the host response following delivery.
The purpose of the present review is (1) to provide rationale
for a macrophage centric approach to tissue reconstruction; and
(2) to give an overview of the current state-of-the-understanding
of the implications associated with host macrophage responses in
regenerative medicine.
It should be noted that the description of macrophages as having either an M1 or M2 phenotype is a simplification of the in vivo
reality. Though it is now clear that M1 and M2 macrophages
each play distinct roles in tissue remodeling following injury, the
inflammatory process which occurs following injury is dynamic
both spatially and temporally and macrophages may express transitional phenotypes. Logically, these cells will also express functions such as phagocytosis, antigen presentation, and effector
molecule production to differing degrees during the inflammation
and remodeling process. For the purposes of simplicity and general
discussion, and as the M1/M2 terminology are used ubiquitously
throughout the literature, we describe macrophage phenotype as
M1 and M2 in the below examples with further discussion of
the spectrum of possible phenotypes and their potential roles in
regenerative medicine thereafter.

A MACROPHAGE CENTRIC APPROACH
There is evidence for both pathogenic and protective roles of
macrophages in many biologic processes (12, 23). It is well understood that uncontrolled inflammation can be a detrimental process
(e.g., inflammatory bowel disease, rheumatoid arthritis). However, an organized and well regulated macrophage response has
been shown to be a determinant of tissue remodeling following
injury, with the potential for positive outcomes and functional
recovery. The key role of macrophages in functional recovery
following injury suggests that methods which are capable of modulating the macrophage response in a controlled, reproducible,
and well-defined manner, may also meet with improved outcomes in regenerative medicine applications. Below, we review
the role of macrophages in the response to tissue injury and the
subsequent remodeling process in three different tissue environments as a baseline from which to understand the potential role of
macrophages in regenerative medicine approaches to tissue reconstruction and to provide the rationale for a macrophage centric
approach.

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Rethinking regenerative medicine: a macrophage-centered approach

THE M1/M2 PARADIGM DURING THE SKELETAL MUSCLE INJURY
RESPONSE

The role of the M1/M2 paradigm during the skeletal muscle injury
response is relatively well characterized. Skeletal muscle tissue possesses inherent regenerative capacity following acute injury such as
exercise-induced trauma. The capacity of muscle tissue to regenerate relies heavily upon a population of normally quiescent muscle
specific progenitor cells, referred to as “satellite cells,” and their
interactions with inflammatory cells that infiltrate the injured
muscle microenvironment (24, 25). An efficient skeletal muscle
injury response which successfully restores the injured muscle tissue requires satellite cell progression through a structured process
of activation including proliferation and subsequent maturation
into committed myoblasts, myoblast alignment, and finally, fusion
and differentiation into new contractile skeletal muscle myotubes
(26–29). This carefully regulated process of satellite cell differentiation is controlled, in large part, by the activity of an orchestrated
heterogeneous inflammatory response consisting predominantly
of M1 and M2 polarized macrophages (30, 31).
Following acute skeletal muscle injury, one of the earliest events
is the infiltration of the damaged tissue by inflammatory cells.
Neutrophils comprise the initial wave of cells to enter the damaged
muscle tissue and reach elevated levels as soon as 2 h post-injury
and maximum numbers after 6–24 h (30, 32, 33). Neutrophils
phagocytose host necrotic cellular or bacterial debris and propagate a pro-inflammatory response through the release of reactive
oxygen species and T-helper (Th)1 associated pro-inflammatory
cytokines, which recruit monocytes and macrophages. However,
neutrophil numbers decline through apoptosis and the neutrophil response is generally resolved by 3–4 days post-injury (30).
Monocyte-derived macrophages recruited to the damaged tissue
shortly after neutrophil infiltration represent the predominant
immunologic participant in the skeletal muscle injury response
thereafter.
Monocytes originate in the bone marrow and express
chemokine receptors and adhesion molecules which facilitate their
extravasation and migration from the blood to the injured skeletal muscle tissue site where they differentiate into macrophages
(34, 35). Tissue resident macrophages normally found within
the skeletal muscle microenvironment are also likely to play
a role. However, the relative contributions of tissue resident
macrophages as compared to circulating macrophages to the
tissue remodeling process remain unknown. Immediately after
injury, infiltrating macrophages become polarized toward a proinflammatory or M1 phenotype. The mechanisms behind this
M1 activation remain only partially understood and include
macrophage exposure to pro-inflammatory cytokines (i.e., IFNγ, TNF-α) and/or necrotic cellular or bacterial debris (30, 35–37).
M1 macrophages within the injured muscle microenvironment
phagocytose necrotic muscle debris and participate in a transient pro-inflammatory response, reaching elevated levels at 24 h
post-injury and maximum numbers after 2 days (30, 35). In addition to producing large amounts of pro-inflammatory cytokines
(i.e., TNF-α, IL-1β, IL-12), M1 macrophages process and present
antigen and express high levels of iNOS which facilitates NO
production (38, 39).

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After 2 days, macrophages participating in the remodeling of injured skeletal muscle show a transition from the
pro-inflammatory M1 to the immunoregulatory and antiinflammatory M2 phenotype. The mechanisms behind this M1
to M2 phenotypic switch remain only partially understood but
include exposure of M1 macrophages to increased IL-10 concentrations from skeletal muscle at 48 h post-injury (40, 41);
M1 macrophage mediated phagocytosis of apoptotic, as opposed
to necrotic, cells (40, 42); and exposure of M1 macrophages to
degradation products from extracellular matrix (ECM) (43). M2
macrophages reach peak numbers within areas of injured muscle at 4 days post-injury and remain a predominant cell-type
present in the remodeling muscle microenvironment for several
days (44, 45). M2 macrophages facilitate resolution of inflammation through the release of anti-inflammatory cytokines (i.e.,
IL-10, IL-13), which deactivate pro-inflammatory cell phenotypes
and promote tissue remodeling and repair (35, 44, 45).
This transition of the initial response dominated by M1
macrophages to a more M2 dominated population following acute
muscle injury facilitates skeletal muscle remodeling and is required
for efficient and compete functional restoration. Specifically, the
pro-inflammatory products of M1 macrophages promote the activation and expansion of quiescent muscle satellite cells within the
tissue injury site (35, 38, 46, 47). For example, TNF-α produced
in large quantities by M1 macrophages represents a well-accepted
mitogen for satellite cell-derived skeletal muscle myoblasts (31,
48). Following satellite cell and myoblast expansion, paracrine
signals from M2 macrophages facilitate the alignment, fusion,
and differentiation, of these skeletal muscle progenitor cells. For
example, IL-10, an immunomodulatory cytokine produced by M2
macrophages, is myogenic for skeletal muscle progenitor cells (4,
31).
The participation of a heterogeneous population macrophages
following muscle injury is highly regulated. For example, the perturbation or prolongation of either the M1 or M2 macrophage
population during the skeletal muscle injury response results
in impaired skeletal muscle regeneration. Depletion of phagocytic leukocytes, including macrophages, prior to toxin induced
skeletal muscle injury blocks the removal of cellular debris and
impairs regeneration (31). Depletion of macrophages at the time
of injury prevents the participation of M1 macrophages in the
early response and therefore shows their importance to muscle
regeneration (31, 49). Furthermore, immediately following injury,
skeletal muscle shows decreased activity of muscle specific transcription factors in TNF-α knockout animals when compared to
their wild-type counterparts (50, 51), suggesting that TNF-α from
M1 macrophages promotes the early or proliferative stage of myogenesis. However, prolonging TNF-α activity beyond the early
proliferative stage of myogenesis has deleterious effects. While
TNF-α promotes skeletal muscle precursor cell mitogenesis, it also
inhibits myogenesis (52–54).
The participation of M2 macrophages is also required for
the skeletal muscle injury response. As stated above, these
immunomodulatory cells drive the late or differentiation stage
of myogenesis. For example, when macrophages were deleted
after 2 days post-injury, a time point consistent with the transition to an M2 macrophage response, myoblast differentiation, and

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Rethinking regenerative medicine: a macrophage-centered approach

subsequent regeneration was impaired (55). Furthermore, transgenic animals unable to mount a M2 macrophage response show
an accumulation of proliferative myoblasts and a lack of myogenic
differentiation following injury (7). These studies show the importance of a present, yet regulated, M1 to M2 phenotypic transition
of macrophages for efficient skeletal muscle regeneration.
THE M1/M2 PARADIGM DURING CUTANEOUS WOUND HEALING

Adult mammalian cutaneous wound healing is another highly
regulated process that follows a sequence of events comprising
the following three interdependent and overlapping phases: (1)
the inflammatory phase; (2) the granulation tissue formation
and wound contraction phase; and (3) the matrix deposition
and tissue remodeling phase (56, 57). Multiple studies have now
demonstrated distinct macrophage phenotypes associated with
each of these phases and with remodeling outcomes following
injury.
The onset of wound healing, designated the inflammatory
phase, can be further sub-divided into an early and late inflammatory phase. Immediately following injury, hemostasis provides a
provisional matrix for cell migration. During the early inflammatory phase, which occurs at 1–4 days post-injury, neutrophils and
monocyte-derived macrophages respond to pro-inflammatory
signals released from the wound microenvironment including
growth factors, cytokines, damage associated molecular patterns
(DAMPS), and pathogen associated molecular patterns (PAMPS)
(58). These pro-inflammatory effector molecules along with the
presence of necrotic cellular and bacterial debris facilitate the
polarization of infiltrating macrophages toward the M1 phenotype (59). M1 macrophages associated with the early inflammatory
phase are highly phagocytic and participate in the inflammatory phase by producing large quantities of pro-inflammatory
cytokines (i.e., TNF-α), proteases, and ROS with the ultimate
goal of pathogen control and removal of necrotic cell and tissue
debris (57, 59).
The late inflammatory phase, which occurs at 5–7 days postinjury, is marked by an accumulation of apoptotic as opposed to
necrotic cells, which upon phagocytosis facilitate the polarization
of macrophages toward the immunomodulatory M2 phenotype
(59). During the granulation tissue formation and wound contraction phase of cutaneous wound healing at 7–10 days post-injury,
paracrine effector molecules, including cytokines such as IL-10
and growth factors such as VEGF, PDGF-β, and TGF-β, produced
by M2 macrophages recruit fibroblasts into the wound site and
promote myofibroblast differentiation (6, 60). M2 macrophages
continue to release anti-inflammatory and pro-angiogenic factors, which facilitate the resolution of inflammation, recruitment
of endothelial cells, and deposition of new ECM (61, 62). Activated
myofibroblasts bridge the wound gap and develop contractile
forces to facilitate wound contraction. Growth factors produced by
M2 macrophages and myofibroblasts synergistically promote the
proliferation and migration of keratinocytes to facilitate wound
re-epithelialization (60, 61).
The matrix deposition and tissue remodeling phase, which
occurs after 10 days post-injury, is marked by a decrease in
macrophage numbers populating the wound site, along with an
overall decrease in total cellularity. Granulation tissue formation

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reaches a plateau and tissue present within the wound site is
partially remodeled into fibrotic scar tissue at this time (56, 57).
The above stages of cutaneous wound healing are carefully
regulated by the activity of responding macrophages. Similar to
the M1/M2 macrophage paradigm associated with the skeletal
muscle injury response, cutaneous wound healing is dependent
upon a heterogeneous macrophage population and an M1 to
M2 phenotypic transition. The pro-inflammatory activity of M1
macrophages during the early inflammatory phase is required for
efficient pathogen control. Conversely, M2 macrophage activity
during the late inflammatory phase is required for the resolution of inflammation and the recruitment of cells, which facilitate
granulation tissue formation and wound re-epithelialization. Perturbation of the M1 macrophage phenotype during the early
inflammatory phase, either by conditional depletion or due to
impaired recruitment, results in delayed granulation tissue formation and wound closure (63, 64). Similarly, prolonging the M1
macrophage phenotype (i.e., preventing the phenotypic transition to M2) through the exogenous addition of TNF-α during
the late inflammatory phase also resulted in poor wound remodeling outcomes (65). Depletion of M2 macrophages during the
late inflammatory phase results in prolonged inflammation and
impaired wound repair (63, 66). These M2 macrophage depleted
cutaneous wounds resemble chronic wounds typically associated
with the pathogenesis of chronic venous ulcers (CVU) and diabetes. In fact, studies have shown that failure of cutaneous wound
macrophages to undergo the M1 to M2 phenotypic transition represents a hallmark of these chronic inflammatory diseases (65,
67, 68). Taken together, these studies show the importance of
functional macrophage heterogeneity and the extent to which
immunomodulatory effects of M2 macrophages are critical for
efficient wound healing and tissue remodeling.
THE M1/M2 PARADIGM DURING THE CNS INJURY RESPONSE

The role of M1 and M2 macrophages following injury in the central nervous system (CNS) is more ambiguous when compared to
other tissues, and is made more complex by the presence of the
blood–brain barrier. However, similarities to the macrophage heterogeneity associated with the injury response in other tissues are
increasingly being reported.
Central nervous system resident macrophages, referred to as
microglia, have long been considered the primary responders
to injury in the CNS with little to no role having been recognized for circulating cells until recently. Generally, microglia are
recruited to, and form a dense barrier around, the lesion site
immediately following spinal cord injury (69, 70). These activated
microglia produce large quantities of cytotoxic factors and proinflammatory cytokines including IL-1β, IL-6, and TNF-α. This
pro-inflammatory response facilitates pathogen control and debris
clearance, and also the recruitment of neutrophils and bloodderived monocytes and macrophages (5, 71–73); however, this
response is also commonly cited as a driver of poor remodeling outcomes following injury in the CNS. As with other tissue
injury responses, functional remodeling following CNS injury
involves a transition from a pro-inflammatory to an immunoregulatory and homeostatic response. It remains unknown if microglia
show M1 to M2 phenotypic plasticity similar to that observed to

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monocyte-derived macrophages. However, recent studies suggest
that CNS microglia drive an early pro-inflammatory response, but
infiltrating macrophages from the circulation may facilitate the
M2-like tissue remodeling response (74). Specifically, recruited
blood-derived macrophages, showing an anti-inflammatory phenotype consistent with M2 polarization, do not directly enter the
lesion center, but are found around the lesion site at 3 days postinjury (74, 75). These immunoregulatory macrophages have been
shown to arrive at the site of injury by specifically trafficking
through a remote blood–cerebrospinal-fluid (CSF) barrier, the
brain ventricular choroid plexus (CP) (8). Once at the injury
site, these M2 macrophages produce IL-10 for mitigation of the
pro-inflammatory response and contribute to repair mechanisms
including remyelination (8, 75, 76).
The participation of M2 polarized macrophages in the CNS
injury response is essential to the repair process. For example,
the endogenous partial recovery which can be observed following spinal cord injury is abrogated when M2-like macrophages are
depleted using antibodies or conditional ablation (74, 75). Consistent with this notion, blockage of CP mediated macrophage
trafficking inhibits M2 macrophage recruitment and subsequently
impaired recovery following injury (76). These studies show
the importance of a heterogeneous macrophage response to
CNS injury and, specifically, that M2 macrophages contribute to
processes beyond inflammation.
The above studies support an emerging dogma of effective
recovery from tissue injury in which initial responses consisting predominantly of M1 macrophages and secondary or later
stages consisting predominantly of M2 macrophages drive functional remodeling outcomes. Furthermore, it is appears that M2
macrophages contribute to more than immunomodulation during the response which follows tissue injury. Several organ systems,
in addition to the above examples of muscle, skin, and CNS tissue have now been shown to undergo similar responses following
injury and are also characterized by heterogeneous and temporally
shifting macrophage phenotypes.

A DEPARTURE FROM THE “CLASSICAL PARADIGM”
The observation of dichotomies in macrophage phenotype in disease pathogenesis as well as tissue remodeling following injury
represents a departure from the classical understanding of the
macrophage as a primarily phagocytic and pro-inflammatory cell.
The foreign-body reaction (FBR) has been well studied over the
last three decades (77). Logically, this response is an extension
of normal wound healing as the implantation of a biomaterial
necessarily requires the creation of a surgical injury. The seminal works in this area by Anderson (77, 78) and others describe
the host response to implanted materials as occurring in stages
including injury (implantation), protein adsorption, acute inflammation, chronic inflammation, FBR, granulation tissue formation,
and encapsulation. These processes are well recognized to be
dominated by mononuclear cells, and macrophages in particular.
During a FBR persistent inflammatory stimuli, such as the
presence of a non-degradable biomaterial, lead to chronic inflammation and the formation of multinucleated foreign-body giant
cells (79). Multinucleate giant cells are formed by the fusion of persistent pro-inflammatory macrophages, consistent with the M1

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phenotype, located at the surface of the biomaterial and further exacerbate the deleterious inflammatory response through
a process known as “frustrated phagocytosis” (80, 81). Failure to
resolve the inflammatory response results in a FBR, leading to
the deposition of disorganized fibrous tissue consistent with scaring and encapsulation of the implant (82, 83). This dense fibrous
scar isolates the implant and prevents its integration with the
surrounding host tissue.
The interpretation of the FBR as a negative occurrence in this
context led to the development of materials with a focus on “inertness” and “biocompatibility” (84–86). This focus upon the host
response to biomaterials resulted in an associated emphasis upon
the material characteristics which determine the host response and
downstream outcomes. However, these same characteristics may
not be ideal in the setting of regenerative medicine where the focus
is upon the restoration of function through the development of
new host tissues rather than through the provision of a simple
mechanical substitute. These concepts do not imply that medical
devices such as hip implants and surgical mesh are not effective
for their intended functions, but rather that the intended use, and
therefore, the design characteristics and functional requirements,
of these materials are incompatible with the goals of regenerative
medicine.
The emergence of regenerative medicine and the need and
desire for therapies which restore endogenous tissue function has
led to a significant increase in our understanding of the role of stem
cells in tissue repair as well as innovation and development of new
biomaterials as stand alone therapies and/or delivery systems for
cells or biologic factors. These materials are most often degradable in nature, and include engineered biologic cues or inherent
bioactivity when derived from natural or tissue based sources. As
such, the host response to these materials will be significantly different and more complex than the response to mono-component,
synthetic or metallic implants. Further complexity is seen when
materials are used in combination with cells or other factors.
It is now recognized that certain materials when used alone
or in concert with a cellular component can provide an inductive
template for constructive and functional tissue remodeling. That
is, the provision of a bioactive material and/or cells leads to the
formation of new, site-appropriate tissue. One example of such
materials is biologic scaffolds composed of ECM (87, 88). These
materials are derived through the decellularization of source tissues and organs and are widely utilized in regenerative medicine
approaches to tissue reconstruction (89, 90). By the nature of the
source (i.e., intact tissue), the materials that result from efficient
decellularization can be thought of as degradable reservoirs of tissue specific structural and functional components. These materials
have been shown to be effective templates for constructive remodeling in both pre-clinical and clinical applications, and in several
body systems (87, 88). However, it should be noted that reports of
the effectiveness of ECM based scaffold materials are variable and
highly dependent on the methods of scaffold production.
Despite the distinct differences in long-term outcomes which
have been observed with various ECM based scaffold materials,
all ECM implants have been shown to elicit a histologically similar cellular response in the first week to month post-implantation
(3, 91). This response is characterized by an early infiltrate of

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neutrophils followed by a dense infiltrate of mononuclear cells.
Under the classical paradigm, such a response would commonly
be associated with progression to a FBR with negative implications
for functional tissue remodeling outcomes. However, the response
typically proceeds down one of three distinct pathways: (1) a classic
FBR with encapsulation and no signs of constructive remodeling;
(2) chronic inflammation and degradation or integration of the
material with little to no constructive remodeling; or (3) reduction of the inflammatory infiltrate with subsequent constructive
remodeling (3, 91).
Based upon these disparate outcomes, it was hypothesized
that, though the early host response to the materials was histologically similar (i.e., characterized by a dense infiltration of
mononuclear cells in the site of implantation), differences in the
early macrophage phenotype to certain ECM scaffold materials
might exist and that these differences may be related to downstream remodeling outcomes. Indeed, this hypothesis was shown
to be correct with those ECM scaffolds which elicited constructive
remodeling outcomes being associated with a timely transition
from an M1 to an M2 macrophage phenotype (3, 92, 93). These
studies have provided the impetus for investigation of macrophage
phenotype in a number of regenerative medicine applications
using biomaterials and cell-based therapies. The results of these
investigations now clearly show a correlation between macrophage
phenotype and successful outcomes associated with multiple
regenerative medicine strategies. A review of selected studies which
demonstrate this phenomenon are described below with a focus
upon multiple strategies (materials, cells, and bioactive factors)
which show associations between macrophage phenotype and
remodeling outcomes.
THE M1/M2 PARADIGM IN TISSUE ENGINEERING AND REGENERATIVE
MEDICINE

An endogenous host injury response, consisting of immunomodulation, including the participation of M2 type macrophages represents a necessary component of efficient and functional tissue
repair. It is therefore logical that regenerative medicine strategies
aimed at activating or augmenting endogenous repair mechanisms
should utilize a similar strategy. Regenerative medicine strategies
aimed at promoting M2 macrophage activation have included celltherapy and the implementation of synthetic and biologic scaffold
materials, among others.
CELLULAR THERAPY

Cellular therapy is generically defined as the transplantation or
delivery of exogenous cells to sites of injured or missing tissues. Stem and/or progenitor cells are often used in regenerative medicine applications because of their multi-lineage differentiation potential and well-recognized resistance to oxidative
stress (94, 95).
The cell source is most commonly autologous due to immune
rejection considerations, although many studies are investigating the use of allogeneic sources. Cellular therapy based strategies aimed at promoting tissue remodeling have been used to
treat injured tissues including the myocardium, the spinal cord,
and skeletal muscle, among others. Despite moderate pre-clinical
and clinical success, cell-therapy is associated with limitations

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including failure of the exogenous cells to engraft within host
tissue (96–101). It is now increasingly recognized that therapeutic outcomes associated with cellular therapy are largely a
result of paracrine effects exerted by the transplanted or delivered cells upon the injured host tissue microenvironment rather
than direct differentiation of the transplanted cells into new
tissues (102–107). These paracrine effects include modulation
of macrophage polarization and beneficial remodeling events
facilitated by a transition to the M2 macrophage phenotype
(108–110).
Co-culture experiments have shown that the secreted products of different stem/progenitor cells directly promote an M2
macrophage phenotype (111–113). A large number of pre-clinical
studies also support these results. For example, following spinal
cord injury, transplanted bone marrow-derived mesenchymal
stem cells (MSC) modulate the host inflammatory microenvironment by promoting an M1 to M2 transition, which ultimately leads to a permissive environment for axonal extension
and functional recovery (114). Furthermore, following traumatic
brain injury, intravenous (IV) delivery of multipotent progenitor cells promotes the polarization of microglia to an M2-like
phenotype (115). Several studies, using cells of multiple origins (i.e., autologous and allogeneic bone marrow-derived MSCs,
adipose derived MSCs, and umbilical derived MSCs, among others), suggest that the therapeutic effects associated with exogenous cell delivery for the treatment of myocardial infarction are
a result of enhanced macrophage polarization switching (116–
118). Cellular therapy mediated M2 macrophage polarization has
been used to promote tissue remodeling and repair in several
anatomic locations and disease states including kidney ischemiareperfusion injury and asthma associated alveolar inflammation,
among others (119–121).
SCAFFOLD MATERIALS

Regenerative medicine strategies aimed at promoting tissue reconstruction or replacement often employ the use of surgically
implantable synthetic or biologic materials designed to serve as cellular support scaffolds. As described above, implantation of these
materials following injury alters the default injury response. For
example, following surgical placement, synthetic and/or biologic
scaffold materials are able to affect the phenotype of infiltrating
inflammatory cells, host progenitor cell activity, as well as fibrosis
and fibrous capsule development (78, 122). These effects depend
on the scaffold composition, degradability, cellularity, porosity,
and implantation site among others (78).
SYNTHETIC SCAFFOLD MATERIALS

As stated above, the surgical placement of non-degradable synthetic scaffold materials is commonly associated with a FBR consisting of persistent M1 macrophage activity and an increased
deposition of scar tissue (3, 78, 91, 123–125). Recently, strategies aimed at modulating material properties to reduce the persistent pro-inflammatory M1 macrophage response to synthetic
biomaterials have been examined. These strategies have included
alterations in scaffold surface chemistry and structural characteristics. However, some of the studies examining these strategies
are associated with conflicting results. For example, one study

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suggests that synthetic scaffold materials composed of fibers with
smaller diameters are associated with more M2-like macrophage
activation when compared to their larger diameter counterparts
(126). In contrast, another study showed that larger fiber diameter
enhanced M2 macrophage polarization (127).
A recent series of studies has demonstrated the effects of material pore size upon integration of the material as well as macrophage
phenotype (13, 128–131). In these studies, materials were produced with tight distributions of pore sizes. Results showed that
materials possessing pores of roughly 30–40 µm were shown to
integrate with reduced encapsulation and higher vascularity when
implanted into dermis or cardiac tissues (13, 131). These changes
in outcome were also associated with shifts in macrophage phenotype (128, 131). However, interestingly, the shifts in phenotype
were observed to be spatially distinct with cells outside of the pore
templated implants having an increased M2 phenotype as compared to non-porous implants, and the cells within the implant
having a predominantly M1 phenotype (128). These studies suggest that manipulation of the structural and surface characteristics
of synthetic scaffold materials can affect macrophage phenotype. Specifically, some of these manipulations appear to alter
the macrophage response and are also associated with improved
outcomes.
Another manner in which biomaterials can be tailored to promote shifts in macrophage phenotype is through the use of biologically active molecules such as growth factors and cytokines.
Examples of these approaches are numerous and are commonly
employed in regenerative medicine with resulting improvements
in remodeling outcomes. A recent study investigated the effects
of incorporation of either M1 (IFN-γ) or an M2 (IL-4) promoting cytokines within a polysulfone tube upon nerve growth
across a gap defect when the tubes were used as guidance conduits
(132). The results of the study demonstrated that polarization of
macrophages toward a more M2 phenotype was associated with
increased Schwann cell infiltration and neurite outgrowth. These
effects were further examined in vitro, with results suggesting that
macrophage derived factors were at least in part the cause of the
observed chemotaxis of Schwann cells.
BIOLOGIC SCAFFOLD MATERIALS

The biologically derived scaffold materials used in regenerative
medicine applications are sourced from a variety of natural sources
including mammalian tissues as well as plant, insect, and bacterial
sources. These materials offer the inherent advantage of the native
ligand landscape and bioactivity resulting from their source material. This inherent bioactivity also leads to added complexity in the
host response to these materials. Among these materials, scaffolds
derived from mammalian tissues represent the most commonly
used materials in pre-clinical and clinical regenerative medicine
applications (133–135).
Biologic scaffold materials composed of mammalian ECM have
been used to promote constructive tissue remodeling in a variety
of clinical applications including hernia repair, rotator cuff reconstruction, esophageal preservation, and skeletal muscle replacement, among others (88, 136, 137). ECM bioscaffolds are derived
through the decellularization of mammalian tissue (89, 90, 138).
The most common tissue sources are xeno- or allogeneic in nature

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and include decellularized dermis, small intestine, bladder, and
pericardium among others.
It is now well recognized that the ability of these materials to
promote constructive remodeling is tied to their ability to modulate the host macrophage response (3, 92, 93, 135). Multiple
studies have shown that ECM based scaffold materials which are
properly prepared facilitate a transition from an M1 to an M2
phenotype around 7–14 days post-implantation (3, 92, 93). The
exact mechanisms by which these materials facilitate this response
remain largely unknown; however, a number of key aspects have
been identified. The materials must be adequately decellularized
to remove potentially immunogenic cellular constituents and the
material must be able to degrade (3, 92, 93, 139). In the presence
of excess cellular material or if the material has been chemically
crosslinked to prevent degradation, an extended M1 type immune
response with no transition to an M2 response is observed and is
associated with poor remodeling outcomes or encapsulation.
The necessity of degradation for the transition to an M2 phenotype suggests that breakdown products of the ECM scaffold
material may possess immunomodulatory activity. Studies have
shown that ECM bioscaffolds can be solubilized and the degradation products formed into a hydrogel under physiologic conditions (140, 141). This hydrogel ECM, when used as a coating for
polypropylene surgical mesh, can facilitate a transition from the
default M1 and FBR type response to a more M2 type response
with a reduction in the FBR and encapsulation (142). These results,
as well as other recent in vitro studies, further demonstrate the
inherent immunomodulatory nature of ECM based biomaterials
as well as their ability to improve remodeling outcomes (124, 142).

WORDS OF CAUTION
The above examples clearly illustrate an emerging paradigm in
regenerative medicine. That is, strategies which are able to modulate the host response from an M1 to an M2 macrophage
response are associated with better outcomes. However, these
results should be interpreted with caution. Macrophage phenotypes have been described in many ways (143). “M1” and “M2”
(with M2 macrophages including subsets M2a, M2b, and M2c)
represents the common terminology used to describe these cells.
Macrophage phenotypes have also been described as a spectrum
between M1 and M2 with any individual cell being capable of
expressing multiple aspects of either phenotype at any given time.
Given this phenotypic heterogeneity, and the transient nature
of the remodeling process following injury, further study of
biomaterials-macrophage and stem cell-macrophage interactions
are warranted, as is more thorough definition of the resultant phenotypes and their unique functions. It is unlikely that macrophages
which result from interactions with biomaterials, particularly
those with inherent naturally occurring ligand landscapes, or stem
cells will possess phenotypes which precisely resemble the canonical IFN-γ and LPS (M1) or IL-4, IL-13 (M2a), IC and TLR/IL1-R
ligand (M2b), or IL-10 (M2c) activated macrophages.
Adding further complexity to the definition of macrophage
phenotypes in regenerative medicine applications is the variability
in tissue resident macrophage populations. For example, microglia
are the resident macrophages of the brain and derive from the
embryonic yolk sac during development and persist in the brain

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Rethinking regenerative medicine: a macrophage-centered approach

thereafter, presumably through a process of local replication (144–
146). As is described above, these cells have been demonstrated in
a number of studies to have phenotypes which are distinct from
circulating monocyte-derived macrophages and are known to play
distinct roles in a number of CNS disease processes. Other tissue
resident macrophage populations also exist, each with a distinct
and tissue specific phenotype (147). While a full description of
tissue resident macrophages in all body systems and their distinct
phenotypic characteristics is beyond the scope of this review, it
is important to understand how these differences and the relative
contributions of local versus circulating cells will affect outcomes.
At present, studies investigating macrophage phenotype following exposure to biomaterial implants have largely relied upon
single surface or gene expression markers as indicators of M1 and
M2 polarization. It is now well established that macrophages possess highly complex and plastic phenotypes and that the use of
multiple phenotypic markers is essential. Further, a better understanding of the functional implications of these phenotypes is
needed to create a mechanistic understanding of the ways in
which macrophages may direct tissue remodeling outcomes following biomaterial implantation or stem cell delivery. With this
understanding, next generation therapies can be developed to target and modulate specific macrophage phenotypes with desirable
characteristics for the given application.
It should further be recognized that baseline macrophage polarization states may be affected by patient characteristics. As is
mentioned above, there is now increasing evidence for changes
in macrophage phenotype and response to activating stimuli with
age and disease both acute and chronic (12, 23, 148, 149). Commonly employed pharmacologic interventions may also affect the
response. Also, it is logical that the tissue microenvironment which
results following an injury may also be different than that which
is experimentally and sterilely created in an animal model. Thus,
there is a need to investigate macrophage response to regenerative
medicine strategies in animal models which, at least in part, can
mimic aspects of these complex situations.

CONCLUSION
Macrophage polarization has been clearly shown to be an important determinant of success in regenerative medicine strategies
for tissue reconstruction. Macrophages can promote both positive
and negative outcomes, which are dependent upon the context in
which they are encountered, their phenotype, and function. However, at present, there remains much to be investigated and defined
regarding macrophage phenotypes associated with biomaterials,
stem cells, and regenerative medicine. Thus, context specific definitions and identification of beneficial phenotypes are needed.
Similarly, the unique functions of these cells must also be clearly
defined in order to better understand their true role in the remodeling process. Indeed, a focus upon macrophage function during
the process of constructive remodeling may prove more useful
than further characterization of complex phenotypic markers.
Moving studies from correlative to causative and expanding the
number of outcome metrics, both phenotypic and functional, will
assist in defining both biomaterials and stem cell associated phenotypes and also provides targets for next generation regenerative
medicine therapies, which seek to modulate macrophages as a

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means of promoting functional tissue recovery – a macrophage
centric approach. It is increasingly clear that those strategies that
adopt such an approach to regenerative medicine will meet with
improved success.

REFERENCES
1. Adamson R. Role of macrophages in normal wound healing: an overview.
J Wound Care (2009) 18(8):349–51. doi:10.12968/jowc.2009.18.8.43636
2. Koh TJ, DiPietro LA. Inflammation and wound healing: the role of the macrophage. Expert Rev Mol Med (2011) 13:e23. doi:10.1017/S1462399411001943
3. Brown BN, Londono R, Tottey S, Zhang L, Kukla KA, Wolf MT, et al.
Macrophage phenotype as a predictor of constructive remodeling following
the implantation of biologically derived surgical mesh materials. Acta Biomater
(2012) 8(3):978–87. doi:10.1016/j.actbio.2011.11.031
4. Deng B, Wehling-Henricks M, Villalta SA, Wang Y, Tidball JG. IL-10 triggers
changes in macrophage phenotype that promote muscle growth and regeneration. J Immunol (2012) 189(7):3669–80. doi:10.4049/jimmunol.1103180
5. Longbrake EE, Lai W, Ankeny DP, Popovich PG. Characterization and modeling of monocyte-derived macrophages after spinal cord injury. J Neurochem
(2007) 102(4):1083–94. doi:10.1111/j.1471-4159.2007.04617.x
6. Novak ML, Koh TJ. Macrophage phenotypes during tissue repair. J Leukoc Biol
(2013) 93(6):875–81. doi:10.1189/jlb.1012512
7. Ruffell D, Mourkioti F, Gambardella A, Kirstetter P, Lopez RG, Rosenthal
N, et al. A CREB-C/EBPbeta cascade induces M2 macrophage-specific gene
expression and promotes muscle injury repair. Proc Natl Acad Sci U S A (2009)
106(41):17475–80. doi:10.1073/pnas.0908641106
8. Shechter R, Miller O, Yovel G, Rosenzweig N, London A, Ruckh J, et al. Recruitment of beneficial M2 macrophages to injured spinal cord is orchestrated by
remote brain choroid plexus. Immunity (2013) 38(3):555–69. doi:10.1016/j.
immuni.2013.02.012
9. Albina JE, Mills CD, Henry WL Jr, Caldwell MD. Temporal expression of different pathways of 1-arginine metabolism in healing wounds. J Immunol (1990)
144(10):3877–80.
10. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
11. Mantovani A, Sozzani S, Locati M, Allavena P, Sica A. Macrophage polarization: tumor-associated macrophages as a paradigm for polarized M2 mononuclear phagocytes. Trends Immunol (2002) 23(11):549–55. doi:10.1016/S14714906(02)02302-5
12. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
13. Mills CD, Shearer J, Evans R, Caldwell MD. Macrophage arginine metabolism and the inhibition or stimulation of cancer. J Immunol (1992)
149(8):2709–14.
14. Brown BN, Ratner BD, Goodman SB, Amar S, Badylak SF. Macrophage
polarization: an opportunity for improved outcomes in biomaterials and
regenerative medicine. Biomaterials (2012) 33(15):3792–802. doi:10.1016/j.
biomaterials.2012.02.034
15. Capobianco A, Rovere-Querini P. Endometriosis, a disease of the macrophage.
Front Immunol (2013) 4:9. doi:10.3389/fimmu.2013.00009
16. Pechkovsky DV, Prasse A, Kollert F, Engel KM, Dentler J, Luttmann W,
et al. Alternatively activated alveolar macrophages in pulmonary fibrosismediator production and intracellular signal transduction. Clin Immunol
(2010) 137(1):89–101. doi:10.1016/j.clim.2010.06.017
17. Smith KA, Pearson CB, Hachey AM, Xia DL, Wachtman LM. Alternative activation of macrophages in rhesus macaques (Macaca mulatta) with endometriosis.
Comp Med (2012) 62(4):303–10.
18. Stoger JL, Gijbels MJ, van der Velden S, Manca M, van der Loos CM, Biessen EA,
et al. Distribution of macrophage polarization markers in human atherosclerosis. Atherosclerosis (2012) 225(2):461–8. doi:10.1016/j.atherosclerosis.2012.
09.013
19. Pollard JW. Trophic macrophages in development and disease. Nat Rev
Immunol (2009) 9(4):259–70. doi:10.1038/nri2528
20. Godwin JW, Pinto AR, Rosenthal NA. Macrophages are required for adult salamander limb regeneration. Proc Natl Acad Sci U S A (2013) 110(23):9415–20.
doi:10.1073/pnas.1300290110

www.frontiersin.org

Rethinking regenerative medicine: a macrophage-centered approach

21. Li L, Yan B, Shi YQ, Zhang WQ, Wen ZL. Live imaging reveals differing roles
of macrophages and neutrophils during zebrafish tail fin regeneration. J Biol
Chem (2012) 287(30):25353–60. doi:10.1074/jbc.M112.349126
22. Godwin JW, Rosenthal N. Scar-free wound healing and regeneration in
amphibians: immunological influences on regenerative success. Differentiation
(2014) 87(1–2):66–75. doi:10.1016/j.diff.2014.02.002
23. Wynn TA, Chawla A, Pollard JW. Macrophage biology in development, homeostasis and disease. Nature (2013) 496(7446):445–55. doi:10.1038/nature12034
24. Mauro A. Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol (1961)
9:493–5. doi:10.1083/jcb.9.2.493
25. Muir AR, Kanji AH, Allbrook D. The structure of the satellite cells in skeletal
muscle. J Anat (1965) 99(Pt 3):435–44.
26. Wozniak AC, Kong J, Bock E, Pilipowicz O, Anderson JE. Signaling satellite-cell
activation in skeletal muscle: markers, models, stretch, and potential alternate
pathways. Muscle Nerve (2005) 31(3):283–300. doi:10.1002/mus.20263
27. Zammit PS, Partridge TA, Yablonka-Reuveni Z. The skeletal muscle satellite
cell: the stem cell that came in from the cold. J Histochem Cytochem (2006)
54(11):1177–91. doi:10.1369/jhc.6R6995.2006
28. Hawke TJ, Garry DJ. Myogenic satellite cells: physiology to molecular biology.
J Appl Physiol (2001) 91(2):534–51.
29. Charge SB, Rudnicki MA. Cellular and molecular regulation of muscle regeneration. Physiol Rev (2004) 84(1):209–38. doi:10.1152/physrev.00019.2003
30. Tidball JG, Villalta SA. Regulatory interactions between muscle and the
immune system during muscle regeneration. Am J Physiol Regul Integr Comp
Physiol (2010) 298(5):R1173–87. doi:10.1152/ajpregu.00735.2009
31. Arnold L, Henry A, Poron F, Baba-Amer Y, van Rooijen N, Plonquet A, et al.
Inflammatory monocytes recruited after skeletal muscle injury switch into
antiinflammatory macrophages to support myogenesis. J Exp Med (2007)
204(5):1057–69. doi:10.1084/jem.20070075
32. Bondesen BA, Mills ST, Pavlath GK. The COX-2 pathway regulates growth of
atrophied muscle via multiple mechanisms. Am J Physiol Cell Physiol (2006)
290(6):C1651–9. doi:10.1152/ajpcell.00518.2005
33. Contreras-Shannon V, Ochoa O, Reyes-Reyna SM, Sun D, Michalek JE, Kuziel
WA, et al. Fat accumulation with altered inflammation and regeneration in
skeletal muscle of CCR2-/- mice following ischemic injury. Am J Physiol Cell
Physiol (2007) 292(2):C953–67. doi:10.1152/ajpcell.00154.2006
34. Swirski FK, Nahrendorf M, Etzrodt M, Wildgruber M, Cortez-Retamozo V,
Panizzi P, et al. Identification of splenic reservoir monocytes and their deployment to inflammatory sites. Science (2009) 325(5940):612–6. doi:10.1126/
science.1175202
35. Tidball JG. Inflammatory processes in muscle injury and repair. Am J Physiol
Regul Integr Comp Physiol (2005) 288(2):R345–53. doi:10.1152/ajpregu.00454.
2004
36. St Pierre BA, Tidball JG. Differential response of macrophage subpopulations
to soleus muscle reloading after rat hindlimb suspension. J Appl Physiol (1994)
77(1):290–7.
37. McLennan IS. Degenerating and regenerating skeletal muscles contain several
subpopulations of macrophages with distinct spatial and temporal distributions. J Anat (1996) 188(Pt 1):17–28.
38. Villalta SA, Nguyen HX, Deng B, Gotoh T, Tidball JG. Shifts in macrophage
phenotypes and macrophage competition for arginine metabolism affect the
severity of muscle pathology in muscular dystrophy. Hum Mol Genet (2009)
18(3):482–96. doi:10.1093/hmg/ddn376
39. Gordon S, Taylor PR. Monocyte and macrophage heterogeneity. Nat Rev
Immunol (2005) 5(12):953–64. doi:10.1038/nri1733
40. Cohen HB, Mosser DM. Extrinsic and intrinsic control of macrophage
inflammatory responses. J Leukoc Biol (2013) 94(5):913–9. doi:10.1189/jlb.
0413236
41. Liu D, Sartor MA, Nader GA, Gutmann L, Treutelaar MK, Pistilli EE, et al.
Skeletal muscle gene expression in response to resistance exercise: sex specific
regulation. BMC Genomics (2010) 11:659. doi:10.1186/1471-2164-11-659
42. Fadok VA, Bratton DL, Konowal A, Freed PW, Westcott JY, Henson PM.
Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production through autocrine/paracrine mechanisms involving
TGF-beta, PGE2, and PAF. J Clin Invest (1998) 101(4):890–8.
43. Sicari BM, Dziki JL, Siu BF, Medberry CJ, Dearth CL, Badylak SF. The promotion of a constructive macrophage phenotype by solubilized extracellular
matrix. Biomaterials (2014) 35(30):8605–12. doi:10.1016/j.biomaterials.2014.
06.060

November 2014 | Volume 5 | Article 510 | 87

Brown et al.

44. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism
and functions. Immunity (2010) 32(5):593–604. doi:10.1016/j.immuni.2010.
05.007
45. Gordon S. Alternative activation of macrophages. Nat Rev Immunol (2003)
3(1):23–35. doi:10.1038/nri978
46. Collins RA, Grounds MD. The role of tumor necrosis factor-alpha (TNFalpha) in skeletal muscle regeneration. Studies in TNF-alpha(-/-) and TNFalpha(-/-)/LT-alpha(-/-) mice. J Histochem Cytochem (2001) 49(8):989–1001.
doi:10.1177/002215540104900807
47. Zador E, Mendler L, Takacs V, de Bleecker J, Wuytack F. Regenerating soleus and
extensor digitorum longus muscles of the rat show elevated levels of TNF-alpha
and its receptors, TNFR-60 and TNFR-80. Muscle Nerve (2001) 24(8):1058–67.
doi:10.1002/mus.110
48. Li YP. TNF-alpha is a mitogen in skeletal muscle. Am J Physiol Cell Physiol
(2003) 285(2):C370–6. doi:10.1152/ajpcell.00453.2002
49. Teixeira CF, Zamuner SR, Zuliani JP, Fernandes CM, Cruz-Hofling MA, Fernandes I, et al. Neutrophils do not contribute to local tissue damage, but play
a key role in skeletal muscle regeneration, in mice injected with Bothrops asper
snake venom. Muscle Nerve (2003) 28(4):449–59. doi:10.1002/mus.10453
50. Chen SE, Gerken E, Zhang Y, Zhan M, Mohan RK, Li AS, et al. Role of TNF{alpha} signaling in regeneration of cardiotoxin-injured muscle. Am J Physiol
Cell Physiol (2005) 289(5):C1179–87. doi:10.1152/ajpcell.00062.2005
51. Warren GL, Hulderman T, Jensen N, McKinstry M, Mishra M, Luster MI, et al.
Physiological role of tumor necrosis factor alpha in traumatic muscle injury.
FASEB J (2002) 16(12):1630–2. doi:10.1096/fj.02-0187fje
52. Langen RC, Schols AM, Kelders MC, Wouters EF, Janssen-Heininger YM.
Inflammatory cytokines inhibit myogenic differentiation through activation of
nuclear factor-kappaB. FASEB J (2001) 15(7):1169–80. doi:10.1096/fj.00-0463
53. Langen RC, Van Der Velden JL, Schols AM, Kelders MC, Wouters EF, JanssenHeininger YM. Tumor necrosis factor-alpha inhibits myogenic differentiation through MyoD protein destabilization. FASEB J (2004) 18(2):227–37.
doi:10.1096/fj.03-0251com
54. Langen RC, Schols AM, Kelders MC, van der Velden JL, Wouters EF, JanssenHeininger YM. Muscle wasting and impaired muscle regeneration in a murine
model of chronic pulmonary inflammation. Am J Respir Cell Mol Biol (2006)
35(6):689–96. doi:10.1165/rcmb.2006-0103OC
55. Tidball JG, Wehling-Henricks M. Macrophages promote muscle membrane
repair and muscle fibre growth and regeneration during modified muscle loading in mice in vivo. J Physiol (2007) 578(Pt 1):327–36. doi:10.1113/jphysiol.
2006.118265
56. Singer AJ, Clark RA. Cutaneous wound healing. N Engl J Med (1999)
341(10):738–46. doi:10.1056/NEJM199909023411006
57. Gurtner GC, Werner S, Barrandon Y, Longaker MT. Wound repair and regeneration. Nature (2008) 453(7193):314–21. doi:10.1038/nature07039
58. Huebener P, Schwabe RF. Regulation of wound healing and organ fibrosis by toll-like receptors. Biochim Biophys Acta (2013) 1832(7):1005–17.
doi:10.1016/j.bbadis.2012.11.017
59. Galli SJ, Borregaard N, Wynn TA. Phenotypic and functional plasticity of cells
of innate immunity: macrophages, mast cells and neutrophils. Nat Immunol
(2011) 12(11):1035–44. doi:10.1038/ni.2109
60. Sindrilaru A, Scharffetter-Kochanek K. Disclosure of the culprits:
macrophages-versatile regulators of wound healing. Adv Wound Care (New
Rochelle) (2013) 2(7):357–68. doi:10.1089/wound.2012.0407
61. Hinz B, Phan SH, Thannickal VJ, Prunotto M, Desmouliere A, Varga J, et al.
Recent developments in myofibroblast biology: paradigms for connective tissue remodeling. Am J Pathol (2012) 180(4):1340–55. doi:10.1016/j.ajpath.2012.
02.004
62. Stout RD. Editorial: macrophage functional phenotypes: no alternatives in dermal wound healing? J Leukoc Biol (2010) 87(1):19–21. doi:10.1189/jlb.0509311
63. Lucas T, Waisman A, Ranjan R, Roes J, Krieg T, Muller W, et al. Differential roles of macrophages in diverse phases of skin repair. J Immunol (2010)
184(7):3964–77. doi:10.4049/jimmunol.0903356
64. Brancato SK, Albina JE. Wound macrophages as key regulators of repair: origin, phenotype, and function. Am J Pathol (2011) 178(1):19–25. doi:10.1016/
j.ajpath.2010.08.003
65. Sindrilaru A, Peters T, Wieschalka S, Baican C, Baican A, Peter H, et al. An unrestrained proinflammatory M1 macrophage population induced by iron impairs
wound healing in humans and mice. J Clin Invest (2011) 121(3):985–97.
doi:10.1172/JCI44490

Frontiers in Immunology | Inflammation

Rethinking regenerative medicine: a macrophage-centered approach

66. Mirza R, DiPietro LA, Koh TJ. Selective and specific macrophage ablation is
detrimental to wound healing in mice. Am J Pathol (2009) 175(6):2454–62.
doi:10.2353/ajpath.2009.090248
67. Mirza R, Koh TJ. Dysregulation of monocyte/macrophage phenotype in
wounds of diabetic mice. Cytokine (2011) 56(2):256–64. doi:10.1016/j.cyto.
2011.06.016
68. Khanna S, Biswas S, Shang Y, Collard E, Azad A, Kauh C, et al. Macrophage dysfunction impairs resolution of inflammation in the wounds of diabetic mice.
PLoS One (2010) 5(3):e9539. doi:10.1371/journal.pone.0009539
69. Davalos D, Grutzendler J, Yang G, Kim JV, Zuo Y, Jung S, et al. ATP mediates
rapid microglial response to local brain injury in vivo. Nat Neurosci (2005)
8(6):752–8. doi:10.1038/nn1472
70. Hines DJ, Hines RM, Mulligan SJ, Macvicar BA. Microglia processes block the
spread of damage in the brain and require functional chloride channels. Glia
(2009) 57(15):1610–8. doi:10.1002/glia.20874
71. Pineau I, Lacroix S. Proinflammatory cytokine synthesis in the injured
mouse spinal cord: multiphasic expression pattern and identification of the
cell types involved. J Comp Neurol (2007) 500(2):267–85. doi:10.1002/cne.
21149
72. Yang L, Blumbergs PC, Jones NR, Manavis J, Sarvestani GT, Ghabriel MN. Early
expression and cellular localization of proinflammatory cytokines interleukin1beta, interleukin-6, and tumor necrosis factor-alpha in human traumatic spinal cord injury. Spine (2004) 29(9):966–71. doi:10.1097/00007632200405010-00004
73. Yang L, Jones NR, Blumbergs PC, Van Den Heuvel C, Moore EJ, Manavis J,
et al. Severity-dependent expression of pro-inflammatory cytokines in traumatic spinal cord injury in the rat. J Clin Neurosci (2005) 12(3):276–84.
doi:10.1016/j.jocn.2004.06.011
74. Schwartz M. “Tissue-repairing” blood-derived macrophages are essential for
healing of the injured spinal cord: from skin-activated macrophages to
infiltrating blood-derived cells? Brain Behav Immun (2010) 24(7):1054–7.
doi:10.1016/j.bbi.2010.01.010
75. Shechter R, London A, Varol C, Raposo C, Cusimano M, Yovel G, et al.
Infiltrating blood-derived macrophages are vital cells playing an antiinflammatory role in recovery from spinal cord injury in mice. PLoS Med
(2009) 6(7):e1000113. doi:10.1371/journal.pmed.1000113
76. Miron VE, Boyd A, Zhao JW, Yuen TJ, Ruckh JM, Shadrach JL, et al. M2
microglia and macrophages drive oligodendrocyte differentiation during CNS
remyelination. Nat Neurosci (2013) 16(9):1211–8. doi:10.1038/nn.3469
77. Anderson JM. Inflammatory response to implants. ASAIO Trans (1988)
34(2):101–7. doi:10.1097/00002480-198804000-00005
78. Anderson JM, Rodriguez A, Chang DT. Foreign body reaction to biomaterials.
Semin Immunol (2008) 20(2):86–100. doi:10.1016/j.smim.2007.11.004
79. Dadsetan M, Jones JA, Hiltner A, Anderson JM. Surface chemistry mediates
adhesive structure, cytoskeletal organization, and fusion of macrophages. J Biomed Mater Res A (2004) 71(3):439–48. doi:10.1002/jbm.a.30165
80. Henson PM. The immunologic release of constituents from neutrophil leukocytes. II. Mechanisms of release during phagocytosis, and adherence to nonphagocytosable surfaces. J Immunol (1971) 107(6):1547–57.
81. Hernandez-Pando R, Bornstein QL, Aguilar Leon D, Orozco EH, Madrigal VK,
Martinez Cordero E. Inflammatory cytokine production by immunological
and foreign body multinucleated giant cells. Immunology (2000) 100(3):352–8.
doi:10.1046/j.1365-2567.2000.00025.x
82. Kovacs EJ. Fibrogenic cytokines: the role of immune mediators in the development of scar tissue. Immunol Today (1991) 12(1):17–23. doi:10.1016/01675699(91)90107-5
83. Williams GT, Williams WJ. Granulomatous inflammation – a review. J Clin
Pathol (1983) 36(7):723–33. doi:10.1136/jcp.36.7.723
84. Ratner BD. The biocompatibility manifesto: biocompatibility for the twentyfirst century. J Cardiovasc Transl Res (2011) 4(5):523–7. doi:10.1007/s12265011-9287-x
85. Williams DF. On the mechanisms of biocompatibility. Biomaterials (2008)
29(20):2941–53. doi:10.1016/j.biomaterials.2008.04.023
86. Brown BN, Badylak SF. Expanded applications, shifting paradigms and an
improved understanding of host-biomaterial interactions. Acta Biomater
(2013) 9(2):4948–55. doi:10.1016/j.actbio.2012.10.025
87. Badylak SF, Brown BN, Gilbert TW, Daly KA, Huber A, Turner NJ. Biologic
scaffolds for constructive tissue remodeling. Biomaterials (2011) 32(1):316–9.
doi:10.1016/j.biomaterials.2010.09.018

November 2014 | Volume 5 | Article 510 | 88

Brown et al.

88. Brown BN, Badylak SF. Extracellular matrix as an inductive scaffold for functional tissue reconstruction. Transl Res (2014) 163(4):268–85. doi:10.1016/j.
trsl.2013.11.003
89. Crapo PM, Gilbert TW, Badylak SF. An overview of tissue and whole organ
decellularization processes. Biomaterials (2011) 32(12):3233–43. doi:10.1016/
j.biomaterials.2011.01.057
90. Gilbert TW, Sellaro TL, Badylak SF. Decellularization of tissues and organs.
Biomaterials (2006) 27(19):3675–83. doi:10.1016/j.biomaterials.2006.02.014
91. Valentin JE, Badylak JS, McCabe GP, Badylak SF. Extracellular matrix bioscaffolds for orthopaedic applications. A comparative histologic study. J Bone Joint
Surg Am (2006) 88(12):2673–86. doi:10.2106/JBJS.E.01008
92. Badylak SF, Valentin JE, Ravindra AK, McCabe GP, Stewart-Akers AM.
Macrophage phenotype as a determinant of biologic scaffold remodeling. Tissue Eng Part A (2008) 14(11):1835–42. doi:10.1089/ten.tea.2007.0264
93. Brown BN, Valentin JE, Stewart-Akers AM, McCabe GP, Badylak SF.
Macrophage phenotype and remodeling outcomes in response to biologic scaffolds with and without a cellular component. Biomaterials (2009)
30(8):1482–91. doi:10.1016/j.biomaterials.2008.11.040
94. Rafii S, Lyden D. Therapeutic stem and progenitor cell transplantation
for organ vascularization and regeneration. Nat Med (2003) 9(6):702–12.
doi:10.1038/nm0603-702
95. Urish KL, Vella JB, Okada M, Deasy BM, Tobita K, Keller BB, et al. Antioxidant levels represent a major determinant in the regenerative capacity of
muscle stem cells. Mol Biol Cell (2009) 20(1):509–20. doi:10.1091/mbc.E0803-0274
96. Guerette B, Asselin I, Skuk D, Entman M, Tremblay JP. Control of inflammatory damage by anti-LFA-1: increase success of myoblast transplantation. Cell
Transplant (1997) 6(2):101–7. doi:10.1016/S0963-6897(96)00230-8
97. Huard J, Acsadi G, Jani A, Massie B, Karpati G. Gene transfer into skeletal muscles by isogenic myoblasts. Hum Gene Ther (1994) 5(8):949–58.
doi:10.1089/hum.1994.5.8-949
98. Fan Y, Maley M, Beilharz M, Grounds M. Rapid death of injected myoblasts
in myoblast transfer therapy. Muscle Nerve (1996) 19(7):853–60. doi:10.1002/
(SICI)1097-4598(199607)19:7<853::AID-MUS7>3.0.CO;2-8
99. Camargo FD, Chambers SM, Drew E, McNagny KM, Goodell MA. Hematopoietic stem cells do not engraft with absolute efficiencies. Blood (2006)
107(2):501–7. doi:10.1182/blood-2005-02-0655
100. Glimm H, Oh IH, Eaves CJ. Human hematopoietic stem cells stimulated to
proliferate in vitro lose engraftment potential during their S/G(2)/M transit
and do not reenter G(0). Blood (2000) 96(13):4185–93.
101. Sicari BM, Dearth CL, Badylak SF. Tissue engineering and regenerative medicine approaches to enhance the functional response to skeletal muscle injury.
Anat Rec (2014) 297(1):51–64. doi:10.1002/ar.22794
102. Gnecchi M, He H, Liang OD, Melo LG, Morello F, Mu H, et al. Paracrine action
accounts for marked protection of ischemic heart by Akt-modified mesenchymal stem cells. Nat Med (2005) 11(4):367–8. doi:10.1038/nm0405-367
103. Murry CE, Reinecke H, Pabon LM. Regeneration gaps: observations on stem
cells and cardiac repair. J Am Coll Cardiol (2006) 47(9):1777–85. doi:10.1016/
j.jacc.2006.02.002
104. Gharaibeh B, Lavasani M, Cummins JH, Huard J. Terminal differentiation is not
a major determinant for the success of stem cell therapy – cross-talk between
muscle-derived stem cells and host cells. Stem Cell Res Ther (2011) 2(4):31.
doi:10.1186/scrt72
105. Perez-Ilzarbe M, Agbulut O, Pelacho B, Ciorba C, San Jose-Eneriz E, Desnos
M, et al. Characterization of the paracrine effects of human skeletal myoblasts
transplanted in infarcted myocardium. Eur J Heart Fail (2008) 10(11):1065–72.
doi:10.1016/j.ejheart.2008.08.002
106. Gnecchi M, Zhang Z, Ni A, Dzau VJ. Paracrine mechanisms in adult stem
cell signaling and therapy. Circ Res (2008) 103(11):1204–19. doi:10.1161/
CIRCRESAHA.108.176826
107. Arthur A, Zannettino A, Gronthos S. The therapeutic applications of multipotential mesenchymal/stromal stem cells in skeletal tissue repair. J Cell Physiol
(2009) 218(2):237–45. doi:10.1002/jcp.21592
108. Le Blanc K, Ringden O. Immunomodulation by mesenchymal stem cells and
clinical experience. J Intern Med (2007) 262(5):509–25. doi:10.1111/j.13652796.2007.01844.x
109. Le Blanc K. Mesenchymal stromal cells: tissue repair and immune modulation.
Cytotherapy (2006) 8(6):559–61. doi:10.1080/14653240601045399

www.frontiersin.org

Rethinking regenerative medicine: a macrophage-centered approach

110. Le Blanc K. Immunomodulatory effects of fetal and adult mesenchymal stem
cells. Cytotherapy (2003) 5(6):485–9. doi:10.1080/14653240310003611
111. Kim J, Hematti P. Mesenchymal stem cell-educated macrophages: a novel type
of alternatively activated macrophages. Exp Hematol (2009) 37(12):1445–53.
doi:10.1016/j.exphem.2009.09.004
112. Eggenhofer E, Hoogduijn MJ. Mesenchymal stem cell-educated macrophages.
Trans Res (2012) 1(1):12. doi:10.1186/2047-1440-1-12
113. Gao S, Mao F, Zhang B, Zhang L, Zhang X, Wang M, et al. Mouse bone
marrow-derived mesenchymal stem cells induce macrophage M2 polarization through the nuclear factor-kappaB and signal transducer and activator of transcription 3 pathways. Exp Biol Med (2014) 239(3):366–75.
doi:10.1177/1535370213518169
114. Nakajima H, Uchida K, Guerrero AR, Watanabe S, Sugita D, Takeura N, et al.
Transplantation of mesenchymal stem cells promotes an alternative pathway of
macrophage activation and functional recovery after spinal cord injury. J Neurotrauma (2012) 29(8):1614–25. doi:10.1089/neu.2011.2109
115. Walker PA, Bedi SS, Shah SK, Jimenez F, Xue H, Hamilton JA, et al. Intravenous
multipotent adult progenitor cell therapy after traumatic brain injury: modulation of the resident microglia population. J Neuroinflammation (2012) 9:228.
doi:10.1186/1742-2094-9-228
116. Dayan V, Yannarelli G, Billia F, Filomeno P, Wang XH, Davies JE, et al. Mesenchymal stromal cells mediate a switch to alternatively activated monocytes/macrophages after acute myocardial infarction. Basic Res Cardiol (2011)
106(6):1299–310. doi:10.1007/s00395-011-0221-9
117. Yannarelli G, Dayan V, Pacienza N, Lee CJ, Medin J, Keating A. Human umbilical cord perivascular cells exhibit enhanced cardiomyocyte reprogramming and
cardiac function after experimental acute myocardial infarction. Cell Transplant
(2013) 22(9):1651–66. doi:10.3727/096368912X657675
118. Adutler-Lieber S, Ben-Mordechai T, Naftali-Shani N, Asher E, Loberman D,
Raanani E, et al. Human macrophage regulation via interaction with cardiac
adipose tissue-derived mesenchymal stromal cells. J Cardiovasc Pharmacol Ther
(2013) 18(1):78–86. doi:10.1177/1074248412453875
119. Wise AF, Williams TM, Kiewiet MB, Payne NL, Siatskas C, Samuel CS, et al.
Human mesenchymal stem cells alter macrophage phenotype and promote
regeneration via homing to the kidney following ischemia-reperfusion injury.
Am J Physiol Renal Physiol (2014) 306(10):F1222–35. doi:10.1152/ajprenal.
00675.2013
120. Geng Y, Zhang L, Fu B, Zhang J, Hong Q, Hu J, et al. Mesenchymal
stem cells ameliorate rhabdomyolysis-induced acute kidney injury via the activation of M2 macrophages. Stem Cell Res Ther (2014) 5(3):80. doi:10.1186/
scrt469
121. Song X, Xie S, Lu K, Wang C. Mesenchymal stem cells alleviate experimental asthma by inducing polarization of alveolar macrophages. Inflammation
(2014). doi:10.1007/s10753-014-9954-6
122. Brodbeck WG, Anderson JM. Giant cell formation and function. Curr Opin
Hematol (2009) 16(1):53–7. doi:10.1097/MOH.0b013e32831ac52e
123. Wolf MT, Dearth CL, Ranallo CA, LoPresti ST, Carey LE, Daly KA, et al.
Macrophage polarization in response to ECM coated polypropylene mesh.
Biomaterials (2014) 35(25):6838–49. doi:10.1016/j.biomaterials.2014.04.115
124. Faulk DM, Londono R, Wolf MT, Ranallo CA, Carruthers CA, Wildemann
JD, et al. ECM hydrogel coating mitigates the chronic inflammatory response
to polypropylene mesh. Biomaterials (2014) 35(30):8585–95. doi:10.1016/j.
biomaterials.2014.06.057
125. Wolf MT, Vodovotz Y, Tottey S, Brown BN, Badylak SF. Predicting in vivo
responses to biomaterials via combined in vitro and in silico analysis. Tissue
Eng Part C Methods (2014). doi:10.1089/ten.tec.2014.0167
126. Saino E, Focarete ML, Gualandi C, Emanuele E, Cornaglia AI, Imbriani M, et al.
Effect of electrospun fiber diameter and alignment on macrophage activation
and secretion of proinflammatory cytokines and chemokines. Biomacromolecules (2011) 12(5):1900–11. doi:10.1021/bm200248h
127. Garg K, Pullen NA, Oskeritzian CA, Ryan JJ, Bowlin GL. Macrophage functional polarization (M1/M2) in response to varying fiber and pore dimensions
of electrospun scaffolds. Biomaterials (2013) 34(18):4439–51. doi:10.1016/j.
biomaterials.2013.02.065
128. Sussman EM, Halpin MC, Muster J, Moon RT, Ratner BD. Porous implants
modulate healing and induce shifts in local macrophage polarization in the
foreign body reaction. Ann Biomed Eng (2014) 42(7):1508–16. doi:10.1007/
s10439-013-0933-0

November 2014 | Volume 5 | Article 510 | 89

Brown et al.

129. Bota PC, Collie AM, Puolakkainen P, Vernon RB, Sage EH, Ratner BD, et al.
Biomaterial topography alters healing in vivo and monocyte/macrophage activation in vitro. J Biomed Mater Res A (2010) 95(2):649–57. doi:10.1002/jbm.a.
32893
130. Fukano Y, Usui ML, Underwood RA, Isenhath S, Marshall AJ, Hauch KD,
et al. Epidermal and dermal integration into sphere-templated porous poly(2hydroxyethyl methacrylate) implants in mice. J Biomed Mater Res A (2010)
94(4):1172–86. doi:10.1002/jbm.a.32798
131. Madden LR, Mortisen DJ, Sussman EM, Dupras SK, Fugate JA, Cuy JL, et al.
Proangiogenic scaffolds as functional templates for cardiac tissue engineering. Proc Natl Acad Sci U S A (2010) 107(34):15211–6. doi:10.1073/pnas.
1006442107
132. Mokarram N, Merchant A, Mukhatyar V, Patel G, Bellamkonda RV. Effect of
modulating macrophage phenotype on peripheral nerve repair. Biomaterials
(2012) 33(34):8793–801. doi:10.1016/j.biomaterials.2012.08.050
133. Badylak SF. Regenerative medicine and developmental biology: the role of the
extracellular matrix. Anat Rec B New Anat (2005) 287(1):36–41. doi:10.1002/
ar.b.20081
134. Badylak SF. The extracellular matrix as a biologic scaffold material. Biomaterials
(2007) 28(25):3587–93. doi:10.1016/j.biomaterials.2007.04.043
135. Badylak SF. Decellularized allogeneic and xenogeneic tissue as a bioscaffold for
regenerative medicine: factors that influence the host response. Ann Biomed
Eng (2014) 42(7):1517–27. doi:10.1007/s10439-013-0963-7
136. Badylak SF, Vorp DA, Spievack AR, Simmons-Byrd A, Hanke J, Freytes DO,
et al. Esophageal reconstruction with ECM and muscle tissue in a dog model.
J Surg Res (2005) 128(1):87–97. doi:10.1016/j.jss.2005.03.002
137. Sicari BM, Rubin JP, Dearth CL, Wolf MT, Ambrosio F, Boninger M, et al.
An acellular biologic scaffold promotes skeletal muscle formation in mice and
humans with volumetric muscle loss. Sci Transl Med (2014) 6(234):234ra58.
doi:10.1126/scitranslmed.3008085
138. Gilbert TW. Strategies for tissue and organ decellularization. J Cell Biochem
(2012) 113(7):2217–22. doi:10.1002/jcb.24130
139. Keane TJ, Londono R, Turner NJ, Badylak SF. Consequences of ineffective
decellularization of biologic scaffolds on the host response. Biomaterials (2012)
33(6):1771–81. doi:10.1016/j.biomaterials.2011.10.054
140. Freytes DO, Martin J, Velankar SS, Lee AS, Badylak SF. Preparation and rheological characterization of a gel form of the porcine urinary bladder matrix.
Biomaterials (2008) 29(11):1630–7. doi:10.1016/j.biomaterials.2007.12.014
141. Wolf MT, Daly KA, Brennan-Pierce EP, Johnson SA, Carruthers CA, D’Amore
A, et al. A hydrogel derived from decellularized dermal extracellular matrix.
Biomaterials (2012) 33(29):7028–38. doi:10.1016/j.biomaterials.2012.06.051

Frontiers in Immunology | Inflammation

Rethinking regenerative medicine: a macrophage-centered approach

142. Wolf MT, Carruthers CA, Dearth CL, Crapo PM, Huber A, Burnsed OA,
et al. Polypropylene surgical mesh coated with extracellular matrix mitigates
the host foreign body response. J Biomed Mater Res A (2014) 102A(1):234–46.
doi:10.1002/jbm.a.34671
143. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental
guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
144. Ginhoux F, Lim S, Hoeffel G, Low D, Huber T. Origin and differentiation of
microglia. Front Cell Neurosci (2013) 7:45. doi:10.3389/fncel.2013.00045
145. Prinz M, Tay TL, Wolf Y, Jung S. Microglia: unique and common features with other tissue macrophages. Acta Neuropathol (2014) 128(3):319–31.
doi:10.1007/s00401-014-1267-1
146. Prinz M, Priller J. Microglia and brain macrophages in the molecular age:
from origin to neuropsychiatric disease. Nat Rev Neurosci (2014) 15(5):300–12.
doi:10.1038/nrn3722
147. Davies LC, Jenkins SJ, Allen JE, Taylor PR. Tissue-resident macrophages. Nat
Immunol (2013) 14(10):986–95. doi:10.1038/ni.2705
148. Akira S, Misawa T, Satoh T, Saitoh T. Macrophages control innate inflammation. Diabetes Obes Metab (2013) 15(Suppl 3):10–8. doi:10.1111/dom.12151
149. Mahbub S, Deburghgraeve CR, Kovacs EJ. Advanced age impairs macrophage
polarization. J Interferon Cytokine Res (2012) 32(1):18–26. doi:10.1089/jir.
2011.0058

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 28 August 2014; accepted: 01 October 2014; published online: 04 November
2014.
Citation: Brown BN, Sicari BM and Badylak SF (2014) Rethinking regenerative medicine: a macrophage-centered approach. Front. Immunol. 5:510. doi:
10.3389/fimmu.2014.00510
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Brown, Sicari and Badylak. This is an open-access article distributed
under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s)
or licensor are credited and that the original publication in this journal is cited, in
accordance with accepted academic practice. No use, distribution or reproduction is
permitted which does not comply with these terms.

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ORIGINAL RESEARCH ARTICLE
published: 08 September 2014
doi: 10.3389/fimmu.2014.00430

Hepatic localization of macrophage phenotypes during
fibrogenesis and resolution of fibrosis in mice and humans
Leonie Beljaars 1 *, Marlies Schippers 1 , Catharina Reker-Smit 1 , Fernando O. Martinez 2 , Laura Helming 3 ,
Klaas Poelstra 1 and Barbro N. Melgert 1
1
2
3

Department of Pharmacokinetics, Toxicology and Targeting, University of Groningen, Groningen, Netherlands
Botnar Research Centre, Nuffield Department of Orthopaedics, Rheumatology and Musculoskeletal Sciences, University of Oxford, Oxford, UK
Institute for Medical Microbiology, Immunology and Hygiene, Technische Universität München, Munich, Germany

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Janos G. Filep, University of
Montreal, Canada
Stephen F. Badylak, University of
Pittsburgh, USA
*Correspondence:
Leonie Beljaars, Department of
Pharmacokinetics, Toxicology and
Targeting, Pharmacy, University of
Groningen, A. Deusinglaan 1,
Groningen 9713 AV, Netherlands
e-mail: e.beljaars@rug.nl

Macrophages have been found to both promote liver fibrosis and contribute to its resolution by acquiring different phenotypes based on signals from the micro-environment. The
best-characterized phenotypes in the macrophage spectrum are labeled M1 (classically
activated) and M2 (alternatively activated). Until now the in situ localization of these phenotypes in diseased livers is poorly described. In this study, we therefore aimed to localize
and quantify M1- and M2-dominant macrophages in diseased mouse and human livers.The
scarred collagen-rich areas in cirrhotic human livers and in CCl4-damaged mouse livers contained many macrophages. Though total numbers of macrophages were higher in fibrotic
livers, the number of parenchymal CD68-positive macrophages was significantly lower as
compared to normal. Scar-associated macrophages were further characterized as either
M1-dominant (IRF-5 and interleukin-12) or M2-dominant (CD206, transglutaminase-2, and
YM-1) and significantly higher numbers of both of these were detected in diseased livers
as compared to healthy human and mouse livers. Interestingly, in mouse, livers undergoing resolution of fibrosis, the total number of CD68+ macrophages was significantly lower
compared to their fibrotic counterparts. M2-dominant (YM-1) macrophages were almost
completely gone in livers undergoing resolution, while numbers of M1-dominant (IRF-5)
macrophages were almost unchanged and the proteolytic activity (MMP9) increased. In
conclusion, this study shows the distribution of macrophage subsets in livers of both
human and murine origin. The presence of M1- and M2-dominant macrophages side by
side in fibrotic lesions suggests that both are involved in fibrotic responses, while the persistence of M1-dominant macrophages during resolution may indicate their importance
in regression of fibrosis. This study emphasizes that immunohistochemical detection of
M1/M2-dominant macrophages provides valuable information in addition to widely used
flow cytometry and gene analysis.
Keywords: cirrhosis, fibrosis, resolution, M1, M2, IRF-5, IL-12, TGM-2

INTRODUCTION
Chronic injury of the liver leads to induction of fibrogenic
processes that ultimately can progress to cirrhosis, a state in
which excessive extracellular matrix deposition hampers normal liver functions. Hepatic stellate cells (HSC) are regarded
as the principal cells that are involved in scar tissue deposition
(1, 2). Recent studies indicate that the role of Kupffer cells has
been underestimated in fibrogenesis and consequently hepatic
macrophages have gained more attention recently (3–7). Kupffer cells are well-known producers of reactive oxygen species,
cytokines and chemokines, that perpetuate hepatic inflammatory
responses, and of matrix-degrading enzymes. In addition, these
macrophages can phagocytose micro-organisms, apoptotic cells,
and cellular debris generated during tissue injury and remodeling.
Duffield et al. (8) clearly showed that Kupffer cells exert different, even opposing roles during various stages of liver fibrosis.
They showed that macrophage activities during the injury phase

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were predominantly associated with promotion of matrix deposition and HSC activities, while during recovery macrophages
were associated with enhanced resolution and higher production
of matrix metalloproteinases (MMPs) (8, 9). These diverse roles
may indicate that activated macrophages differentiate into diverse
phenotypes during various stages of liver disease.
Activated macrophages are described to polarize into different phenotypes depending on signals they receive from their
environment. Many types can be distinguished, and the mostused, but rather simplified, classification system discerns classically activated macrophages (also called M1) and alternatively
activated macrophages (also called M2) (10–12). In fact, these
phenotypes represent their dominant appearance in a wide spectrum of overlapping activation types. Other M2-like transitional
phenotypes have been described as well, but to date these have
been difficult to distinguish from M2 macrophages in situ in
tissues due to lack of phenotype-specific markers (6, 13–15).

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In general, M1-dominant macrophages have enhanced microbicidal and tumoricidal capacity and secrete high levels of proinflammatory cytokines like interleukin-12 (IL-12). M1-dominant
macrophages can also inhibit fibrotic activities of fibroblasts by
releasing antifibrogenic or fibrolytic factors such as MMPs (16,
17). M2-dominant macrophages, activated by interleukin-4 and
interleukin-13, are associated with increased fibrogenesis, tissue
remodeling, and angiogenesis (17–19). In vitro, Song et al. (17)
showed that the M2-dominant macrophages produce profibrogenic factors like platelet-derived growth factor-BB (PDGFBB)
and transforming growth factor-β (TGFbeta) and that these M2dominant cells increase collagen production and proliferation of
fibroblasts. Although M2 macrophages are predominantly considered to be pro-fibrotic, they are also associated with antifibrotic properties, which may be explained by the different and
overlapping M2 phenotypes that exist (5, 11). For instance, M2
macrophages can also aid resolution of fibrosis by phagocytizing
apoptotic cells and matrix components via mannose and scavenger receptors (20–22). In addition, Pesce et al. (23) showed that
arginase-1 expressing M2 cells were related to suppression rather
than induction of fibrosis.
Thus far, most of the knowledge generated about the different
macrophage subsets is derived from in vitro studies, from flow
cytometry analyses of isolated liver macrophages (6), and from
gene analysis of liver homogenates (24). Although these techniques
generate useful quantitative information, histological detection
of macrophages gives unique and additional information with
regard to their tissue localization without selection due to isolation limitations or with minor risk of missing changes because
other cells express the same markers, such as observed in tissue
homogenates (25).
How the different phenotypes are distributed in diseased
liver tissue is still largely unexplored. Therefore, we aimed to
illustrate, using immunohistochemical techniques, how different
macrophage phenotypes are distributed in situ during fibrogenic
responses and resolution of fibrosis using the general M1 and M2
classification as a starting point. Of the markers commonly used,
we chose IL-12 and IRF-5 as markers for the M1-dominant subtype
(26). Inducible nitric oxide synthase (iNOS), another commonly
used M1 marker, was not chosen because its dominant expression in hepatocytes would make distinguishing neighboring iNOS
expressing macrophages difficult (27, 28). To detect M2 polarization, we used upregulation of the mannose receptor (MRC1; also
known as CD206), transglutaminase-2 (TGM-2), and chitinaselike secretory protein YM-1 (mouse only) (29–32). TGM-2 was
recently identified as a new human and murine M2 marker (33).
The commonly used M2 marker arginase could not be used for
reasons similar to iNOS (27).

Localization of liver macrophage subsets

performed according to strict governmental and international
guidelines on animal experimentation.
ANIMAL MODELS

Chronic liver injury (fibrosis) model

Mice received twice-weekly intraperitoneal injections of CCl4 for
4 or 8 weeks. The dose of CCl4 was gradually increased (diluted
in olive oil; week 1: 0.5 ml/kg, week 2: 0.8 ml/kg, week 3–8:
1 ml/kg). Mice were sacrificed after 4 or 8 weeks reflecting early
and advanced fibrosis, respectively.
Resolution model

Mice received CCl4 for 4 weeks (with increasing CCl4 doses as
described in the previous section). After 4 weeks, CCl4 administration was stopped and the mice were allowed to recover for a
week after which they were sacrificed (n = 6 per group).
HUMAN LIVERS

Residual human liver tissue samples were obtained from the
Department of Hepato-Pancreato-Biliary Surgery and Liver Transplantation [University Medical Center Groningen (UMCG), the
Netherlands]. At the UMCG, all patients eligible for organ transplantation are asked to sign a general consent form for the
use of left-over body material (after diagnostic procedures) for
research purposes. The experimental protocols were approved by
the Medical Ethical Committee of the UMCG (Groningen) and
the anonymized tissue samples were used according to Dutch
guidelines. Normal human liver tissue (n = 7) was obtained from
residual liver tissue from donor livers discarded for transplantation because of technical reasons. Cirrhotic human liver tissue
(n = 6) was obtained from patients undergoing liver transplantation. Indications for transplantation were a.o. primary sclerosing
cholangitis (PSC), primary biliary cirrhosis (PBC), congenital cirrhosis, and Wilson’s cirrhosis. Although all human liver material is
anonymized, some available patient characteristics are listed below
(see Table 1).
TISSUE PROCESSING

Tissue specimens from at least three different mouse liver lobes
were snap frozen in isopentane (−80°C) for immunohistochemical analysis, or in liquid nitrogen for western blot analysis.
A wedge (10–60 g) of freshly obtained human liver was cut,
perfused with cold University of Wisconsin organ storage solution (DuPont Critical Care, Waukegan, IL, USA) immediately after
resection, and pieces were snap frozen in isopentane (−80°C).

Table 1 | Available patient characteristics of the used human livers.

MATERIALS AND METHODS

Patient characteristics

ANIMALS

Male mice (BALB/c, ±25 g) were obtained from Harlan (Zeist, The
Netherlands) and housed in a temperature-controlled room with
12 h light/dark regimen. The animal experiments were approved
by the Institutional Animal Care and Use Committee (IACUC)
of the University of Groningen (The Netherlands) and were

Frontiers in Immunology | Molecular Innate Immunity

Age (years)
Gender

Normal livers

Cirrhotic livers

N =7

N =6

41 (10–57)

49 (35–66)

N = 4: F

N = 3: F

N = 2: M

N = 1: M

N = 1: not known

N = 2: not known

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Beljaars et al.

FIGURE 1 | Expression and localization of macrophages in normal
and fibrotic mouse livers (8 weeks CCL4). Immunohistochemical
analysis shows increased extracellular matrix deposition [collagen type I
(A,B)] and presence of macrophages [CD68 (C–F)] in fibrotic (B,D,F) as
compared to healthy livers (A,C,E). Note the increased size of certain
CD68-positive macrophages in the fibrotic areas (F) in the CCL4 livers

IMMUNOHISTOCHEMICAL ANALYSIS

Acetone-fixed cryostat sections (4 µm) were stained according
to standard immunohistochemical procedures with 3-amino-9ethyl-carbazole to detect expression of relevant markers (32).
Sections were incubated with the primary antibody for 1 h. Primary antibodies to detect fibrotic extracellular matrix (polyclonal
goat anti-collagen type I from Southern Biotech), macrophages
[mouse anti-human CD68 (DAKO), monoclonal rat anti-mouse
CD68 (AbD Serotec, Düsseldorf, Germany), and polyclonal rabbit
anti-human CD68 (Santa Cruz Biotechnology)], M1 macrophages
[polyclonal rabbit anti-human and mouse IRF-5 (Protein Tech,
Manchester, UK), goat polyclonal anti-human IL-12 p40 antibody
(ThermoScientific), and goat polyclonal anti-human and mouse
MMP9 (Santa Cruz)], and M2 macrophages [polyclonal goat antimouse chitinase 3-like/ECF-L (YM-1; R&D), rabbit anti-human
TGM-2 (AbD Serotec) and CD206 (rat anti-mouse CD206 and
mouse anti-human CD206) both from BioLegends (ITK Diagnostics, Uithoorn, The Netherlands)] were used. Staining of CD68
was quantified by image analysis with Cell∧ D analysis program
(Olympus, Zoetermeer, The Netherlands).
To detect co-localization, we used double-staining techniques
with peroxidase and AEC (red) and alkaline phosphatase and
Naphtol AS-MX phosphate/Fast Blue BB (blue) (34). Double
stainings for IRF-5, IL-12, and CD206 were visualized with
NovaRed (red) and BCIP/NBT (blue) from Vector Laboratories.
WESTERN BLOT ANALYSIS

Tissues samples were homogenized on ice in cold RIPA buffer
[50 mM Tris-HCl, 150 mM NaCl, 0.1% SDS, 0.1% Igepal in 0.5%
sodium deoxycholate with one tablet of protease inhibitor cocktail and one tablet of phosphatase inhibitor (Roche Diagnostics, Mannheim, Germany)] and lysates were centrifuged for 1 h
(13,000 rpm, 4°C). The supernatants were stored at −80°C. Total
protein (100 µg) from each sample was applied on SDS-PAGE
(10%), transferred to polyvinylidene difluoride membranes, and

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Localization of liver macrophage subsets

[arrow heads in insert (C,F)]. (G,H) Image analysis of CD68 staining.
While the total area of CD68+ cells was increased in fibrotic livers, a
significantly lower CD68-stained area was found in the parenchyma (p) of
fibrotic livers as compared to normal. Magnifications: 40× (A–D), 100×
(E,F), and 200× (inserts). f, Fibrotic matrix; p, liver parenchyma.
N = 6/group. **p < 0.01.

incubated overnight at 4°C with the indicated primary antibodies. After washing and incubation with secondary horseradish
peroxidase-coupled antibodies, the protein bands were visualized with ECL (Perkin-Elmer, Groningen, The Netherlands) and
quantified by G-Box (Syngene, Cambridge, UK).
In order to quantify the marker of interest, we corrected
the expression with the expression of the housekeeping protein
GADPH (for human samples) or β-actin (for mouse samples).
STATISTICAL ANALYSIS

Results are expressed as means ± SEM. All data were analyzed with
the Mann–Whitney U test (Graph Pad software). Differences were
considered significant at p < 0.05.

RESULTS
LOCALIZATION OF CD68-POSITIVE MACROPHAGES IN HUMAN AND
MOUSE FIBROTIC LIVERS

After chronic CCl4 damage (8 weeks CCL4), pericentral necrosis
led to wound-healing responses with influx of myofibroblasts and
increased collagen deposition (Figures 1A,B) in mouse livers. As
compared to normal livers, a higher number of CD68-positive cells
was found in fibrotic livers, and these macrophages predominantly
concentrated in scars during advanced fibrosis (Figures 1C,D,G).
Furthermore, several scar-associated macrophages differed in
appearance from macrophages in normal livers, even some resembled giant cells (Figures 1D,F). Remarkably, significantly less staining for CD68+ cells was found in the parenchymal areas of fibrotic
livers as compared to normal (Figures 1E–H).
Collagen deposition was also greatly increased in end-stage
human cirrhotic livers (Figures 2A,B), and a similar hepatic
distribution of macrophages as in mice was found (Figure 2).
Macrophages (CD68+ ) were prominently present in cirrhotic scars
irrespective of the origin of cirrhosis (Figures 2D–F). The total
number of CD68+ cells was somewhat, though not significantly,
higher in cirrhotic than normal livers (Figure 2G). Again, a trend

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Localization of liver macrophage subsets

FIGURE 2 | Expression and localization of macrophages in normal and
cirrhotic human livers. Immunohistochemical analysis shows enhanced
deposition of the extracellular matrix protein collagen type I (A,B) and
presence of macrophages [CD68 (C–F)] in normal (C) cirrhotic livers of various
origins [(D) PBC (E) PSC and (F) congenital cirrhosis]. Note the abundant

presence of macrophages in the collagenous fibrotic bands (F). (G,H) Image
analysis of CD68 staining in human livers. Reduced CD68 staining was found
in the parenchymal area (p) of human cirrhotic livers as compared to normal.
Magnifications: 40× (A,B) and 100× (C–F). f, fibrotic matrix; p, liver
parenchyma. N = 5 cirrhotic livers, N = 6 normal livers.

FIGURE 3 | Localization of IL-12 (M1) in cirrhotic human livers.
Immunohistochemical staining for IL-12p40 in cirrhotic human livers (B),
while no staining was observed in normal livers (A). (C) Co-localization of

IL-12 and CD68. Arrows indicate co-localization, f, fibrotic matrix.
Magnifications: 100× (A,B) and 400× (C). N = 5 cirrhotic livers, N = 6
normal livers.

toward less staining for CD68 was found in the parenchymal areas
of cirrhotic livers as compared to healthy livers (Figure 2H).
M1-DOMINANT MACROPHAGES IN MOUSE AND HUMAN LIVERS

Interleukin-12, a major cytokine produced by classically activated
macrophages, was used as an immunohistochemical marker to
detect the M1-dominant subset in the liver (Figure 3). In human
livers, higher numbers of IL-12-positive cells were detected in cirrhotic human livers (Figure 3B) as opposed to barely detectable
IL-12 staining in healthy livers (Figure 3A). This increased expression was confirmed by western blot analysis (Figure 4B). IL-12
positive cells were found solely in the cirrhotic collagen bands.
The staining for IL-12 co-localized completely with CD68, but
only a minor fraction of the CD68 population was positive for
IL-12 (Figure 3C).
IRF-5 was also used to identify M1 macrophages in human and
mouse livers (Figure 5). IRF-5 staining co-localized completely
with CD68 in livers of both species (Figure 5A). Similar to IL12, only a subset of the total number of macrophages expressed
IRF-5. To prove the phenotype-specificity of this M1 marker, we
performed double-immunostainings of IRF-5 and the M2 marker
CD206 in human livers and found little to no co-localization

Frontiers in Immunology | Molecular Innate Immunity

(Figure 5B). Microscopic analysis showed that IRF-5 staining was
almost absent in normal mouse and human livers (Figures 5C,E).
In advanced fibrotic mouse livers, the staining was present in cells
residing in the scarred areas (Figure 5D). Similarly, in human livers
IRF-5 staining was also predominantly found in cells of the septa
(Figure 5F). Western blot analysis of liver homogenates revealed
a significantly higher expression of IRF-5 in diseased mouse and
human livers as compared to healthy livers (Figures 4A,B).
M2-DOMINANT MACROPHAGES IN MOUSE AND HUMAN LIVERS

Subsequently, we studied the hepatic distribution of alternatively
activated macrophages (Figures 6–8). CD206 is a well-known
marker for both mouse and human M2-dominant macrophages.
CD206/CD68 double-positive cells were present in fibrotic livers
and were predominantly found in scars (Figure 6A). In addition
to this, CD206 staining was present in liver parenchyma and this
staining most likely reflected expression of CD206 on sinusoidal
endothelial cells [identified with CD31 (Figure 6F)]. The pronounced endothelial staining of CD206 complicates interpretation
of analyzes of whole tissue homogenates (like western blot and
mRNA expression analyses) used in the macrophage field. Microscopic evaluation of sections stained for both CD206 and CD68

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Localization of liver macrophage subsets

FIGURE 4 | Western blot analysis of the expressions of collagen type I, IRF-5 (M1), IL-12 (M1), and YM-1 (M2) in normal and fibrotic mouse (A) and
human (B) livers. *p < 0.05. N = 5 cirrhotic human livers, N = 6 normal human livers, N = 4 normal mouse livers, N = 6 fibrotic mouse livers.

indicated that double-positive cells were more frequent in fibrotic
liver than in normal livers, whereas western blot analysis revealed
reduced expression in both human and mouse fibrotic livers (data
not shown).
YM-1 was used as another M2 marker for mouse livers
(Figure 7). Expression of YM-1 is restricted to mice and can
therefore not be used for human liver tissue (29). YM-1 colocalized with CD68 and with CD206 (Figures 7A,B). All cells that
expressed YM-1 were positive for CD68 and CD206, but not all
CD68-positive cells stained positive for YM-1. The hepatic expression of YM-1 was clearly higher after chronic CCl4 damage as
demonstrated by immunohistochemical staining (Figures 7C–F)
and western blot analysis (Figure 4A). YM-1 was present in the
fibrotic collagenous bands of the CCL4-damaged livers.
The recently described M2 marker TGM-2 (33) was also
used to identify M2-dominant macrophages in human livers
(Figure 8). Immunohistochemical staining for TGM-2 resulted
in staining of the parenchymal area of normal livers, mostly

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staining hepatocytes, but in cirrhotic livers additional strong positive cells were found in septa (Figures 8A,B). TGM-2 staining
present in scars co-localized with CD68 (Figure 8C) and with
CD206 (Figure 8D) confirming presence of TGM-2 in hepatic M2dominant macrophages that accumulate in these areas. As with
iNOS, arginase-1, and CD206, quantitative evaluation of TGM-2
was confounded by its high expression in hepatocytes. We could
not detect differences between normal and cirrhotic livers (data
not shown).
M1- AND M2-DOMINANT MACROPHAGES IN A MOUSE MODEL OF
RESOLUTION

Cessation of fibrosis-inducing agents induces reversal of the
fibrotic process (35). This is also apparent in our mouse model
with lower hepatic collagen type I in livers of mice in which
CCl4 administration was stopped versus their fibrotic equivalents
(Figures 9A–C). Since macrophages are important during resolution (8, 9), we studied the localization and numbers of macrophage

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Localization of liver macrophage subsets

FIGURE 5 | Localization of IRF-5 (M1) in mouse and human livers.
(A) Co-localization of IRF-5 and CD68 in mouse and human livers.
(B) Double-staining for IRF-5 (blue staining) and CD206 (red staining) showed
no co-localization. (C,D) Immunohistochemical staining of IRF-5 in normal

mice livers (C) and in livers after chronic CCl4 damage (D). (E,F) IRF-5 staining
of normal (E) and cirrhotic (F) human livers. f, fibrotic matrix. Magnifications:
100× (C–F) and 400× (A,B). N = 5 cirrhotic human livers, N = 6 normal
human livers, N = 4 normal mouse livers, N = 6 fibrotic mouse livers.

FIGURE 6 | Immunohistochemical staining for CD206 (MCR-1; mannose
receptor) in human (A–C) and mouse (D,E) normal (B,D) and cirrhotic
(C,E) livers. (A) Co-localization of CD206 (blue staining) and CD68 (red
staining). Arrows indicate co-localization, asterisks indicate endothelial

staining of CD206. (F) Immunohistochemical staining for CD31 in fibrotic
mouse livers illustrating staining of sinusoidal endothelial cells.
Magnifications: 100× (B–F) and 400× (A). N = 5 cirrhotic human livers, N = 6
normal human livers, N = 4 normal mouse livers, N = 6 fibrotic mouse livers.

phenotypes in these two groups of mice. Expression of CD68 was
significantly lower in livers undergoing resolution as compared to
their fibrotic counterparts (Figures 9D–H).

Frontiers in Immunology | Molecular Innate Immunity

We detected a slightly reduced expression of IRF-5 with both
immunohistochemical and western blot analysis (Figure 10).
However, a clear difference in the number of M2-dominant

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FIGURE 8 | Localization TGM-2 (M2) in human livers.
(A,B) Immunohistochemical localization of TGM-2 in normal (A) and
cirrhotic (B) human livers. Note the presence of the strongly stained cells in
the fibrotic matrix (F). (C) Co-localization of TGM-2 (blue staining) and CD68
(red staining). (D) Co-localization of TGM-2 (blue staining) and CD206 (red
staining). Arrows indicate co-localization. Magnifications: 100× (A,B) and
400× (C,D). N = 5 cirrhotic human livers, N = 6 normal human livers.

FIGURE 7 | Localization of YM-1 (M2) in mouse livers. (A) Co-localization
of YM-1 (red staining) and CD68 (blue staining). (B) Co-localization of YM-1
(red staining) and CD206 (blue staining). (C–F) Immunohistochemical
localization of YM-1 in livers of normal mice (C,E) and in advanced fibrosis
(D,F). Magnifications: 40× (C,D), 200× (E,F), and 400× (A,B). N = 4
normal mouse livers, N = 6 fibrotic mouse livers.

macrophages was found (Figure 11). YM-1 staining was abundantly present in fibrotic livers, but in livers undergoing resolution
this M2 marker was almost completely gone. Western blot analysis revealed a reduction of 81 ± 8% in YM-1 expression during
resolution (Figure 11E).
Since the number of IRF-5+ (M1-dominant) macrophages was
almost unchanged in fibrotic livers compared to livers undergoing resolution, we measured MMP expression as a functional read
out of the presence of these macrophages. M1-dominant cells are
known to express MMP9 (17, 21, 36, 37) and western blot analysis of these livers revealed significantly higher expression of MM9
92 kDa, which is known as pro-MMP9, and its processed form
(67 kDa MMP9) in the livers undergoing resolution (Figure 10F).

DISCUSSION
Recently, activation of macrophages into different phenotypes has
been subject of study in various diseases including in liver diseases.
Almost all knowledge obtained thus far is derived from in vitro
studies or from FACS or PCR analyses of tissues. These in vitro
studies have been essential to discover markers to distinguish the
various macrophages phenotypes and to identify the specific activities of these subsets. How these in vitro-generated phenotypes
relate to macrophages in situ is largely unexplored. In this study,
results were obtained from the CCL4 mouse model at several time

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points in disease progression (reflecting early and advanced fibrosis) and resolution. Although we are aware that more time points
in this mouse model can support broader conclusions, our outcomes with regard to the presence and localizations of the various
macrophage phenotypes are first steps toward understanding the
dynamics of macrophage phenotypes in relation to localization. A
major advantage of our studies is the verification of mouse data
in samples of human liver disease. The fact that we find similar
distributions of macrophage phenotypes in end-stage disease of
a number of different etiologies may point at converging disease
mechanisms irrespective of cause.
We used many commonly used markers M1- and M2-dominant
phenotypes and found that not all of them can be used reliably
for liver tissue. With the ones that can be used, we demonstrated
that M1- and M2-dominant subsets are localized side by side in
scars of human and mouse cirrhotic livers. Although M1 and
M2 markers can be expected to be present on the same cell,
based on the theory of overlapping spectra of macrophage subsets (13–15), with our markers (IRF-5 and CD206), we found
little to no co-localization. We showed that IL-12 and IRF5 are useful immunohistochemical markers for M1-dominant
macrophages in liver tissue (both mouse and human) and YM1 for M2-dominant macrophages in murine liver tissue. CD206
and TGM-2 can be useful for immunohistochemistry of M2dominant macrophages in human liver tissue, but are much
less specific and therefore are hard to quantify. Furthermore,
in fibrotic livers undergoing resolution we found that M2dominant macrophages (YM-1 positive cells) disappeared, while
M1-dominant macrophages (IRF-5 positive cells) persisted in the
scarred areas producing MMPs.
Interleukin-12 and IRF-5 were used to identify the classically
activated macrophages in fibrotic livers. Krausgruber et al. (26)

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FIGURE 9 | Immunohistochemical and western blot analysis of the
hepatic expressions of collagen type I (A–C) and macrophages [CD68
(D–H)] in fibrosis [4 weeks of CCl4 in mice (A,D,G)] and in livers

undergoing resolution [after cessation of CCl4 administration, resolution
(B,E,H)]. **p < 0.01, # p < 0.1. Magnifications: 40× (A–E) and 200× (G,H).
N = 6/group.

FIGURE 10 | Expressions of IRF-5 and MMP9 (M1) in fibrotic mouse livers
[4 weeks CCl4 (A,B)] and in fibrotic livers undergoing resolution [after
cessation of 4 weeks of CCl4 administration (C,D)]. Immunohistochemical
pictures demonstrate an overview [(A,C) magnification 40×] and close up
[(B,D) magnification 200×]. (E) Western blot quantification of hepatic IRF-5

expression in fibrosis versus resolution group, and (F) western blot
quantification of MMP9 expression in fibrosis versus resolution group.
A 92 kDa pro-form and a 67 kDa processed form of MMP9 is significantly
increasingly expressed in livers undergoing resolution. *p < 0.05, **p < 0.01.
N = 6/group.

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FIGURE 11 | Expressions of YM-1 (M2) in fibrotic mouse livers [4 weeks
CCl4 (A,B)] and in fibrotic livers undergoing resolution [after cessation of
CCl4 administration (C,D)]. Immunohistochemical pictures demonstrate an

showed high expression of IRF-5 in human M1 macrophages
in culture, while M2 and non-activated macrophages did not
express IRF-5. We now show that IRF-5 can be used to identify a subset of macrophages in vivo in human and mouse livers.
Our study clearly demonstrates that M1-dominant macrophages
(CD68/IRF-5+ cells) are significantly increased in diseased livers as compared to normal. IRF-5+ cells are located in fibrous
septa in advanced fibrosis. These localizations may correspond
with reported in vitro M1 activities such as production of proinflammatory cytokines and chemokines (5, 11, 17). The observation that M1-dominant macrophages are still present in livers
undergoing resolution might be related to their ability to produce MMPs (12, 16, 17, 37). Classical activation of macrophages
in vitro resulted in higher expression of MMP7 and MMP9 and
both may be necessary in the collagenous scars for removal of
collagen fibers. Indeed, in our studies we detected higher MMP9
expression in fibrotic livers undergoing resolution. In addition,
Fallowfield et al. (9) demonstrated higher hepatic MMP13 expression by scar-associated macrophages in CCL4-damaged livers and
it was found that resolution of CCl4-induced fibrosis was retarded
in MMP13-deficient mice. However, macrophage phenotypes in
these scars were not further characterized. We now show with
our localization studies that during fibrogenesis scar-associated
macrophages are both of M1- and M2-dominant phenotype, while
during resolution the scar-associated macrophages are predominantly M1 cells. It therefore appears that M1 macrophages may
also be responsible for the MMP13 production that is necessary
for resolution. Co-localization studies with IRF-5 and MMP13
may provide additional insights.
To identify M2-dominant macrophages, we started with
the well-known marker CD206 (mannose receptor, MCR-1)
(11). While in many organs M2-dominant macrophages specifically express CD206, in livers CD206 expression is found in
macrophages as well as in sinusoidal endothelial cells, making
quantitative interpretations difficult. In addition, we therefore
used the well-known M2-selective marker YM-1, which does not
have this disadvantage. However, this marker is only present in
rodents (29) and cannot be used for human tissues. TGM-2 is

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Localization of liver macrophage subsets

overview [(A,C) magnification 40×] and close up [(B,D) magnification 200×].
(E) Western blot quantification of YM-1 expression in fibrosis versus
resolution. **p < 0.01. N = 6/group.

a novel marker for M2 macrophages recently described by us
in lungs (33). The advantage of TGM-2 is that this marker is
conserved in mice and humans. TGM-2 is a multifunctional
enzyme involved in transamidation and cross-linking of proteins. It is also linked to apoptosis, cellular differentiation, and
matrix stabilization (38–40). In liver, Popov et al. (41) showed
that TGM-2 is enhanced in mice with CCl4-induced fibrosis, but
they found no relationship between TGM-2 and stabilization of
fibrotic matrix. However, TGM-2 expression was not related to
macrophage activities. Although the hepatic expression is not limited to macrophages, as can be seen in Figure 8, TGM-2 staining
in the scar-associated macrophages in cirrhotic livers is much
stronger than in other hepatic cells. Therefore, this marker can be
used for immunohistochemical stainings but quantification using
western blot or PCR will not yield useful results. To summarize,
using a combination of the markers CD206, YM-1, and TGM-2,
we are able to show that M2-dominant macrophages are present in
scar tissue during hepatic fibrogenesis. We now show that TGM-2
is co-expressed in CD68+ and in CD206+ cells in fibrotic septa in
human and mouse livers, confirming its presence in M2-dominant
hepatic macrophages.
This study clearly shows the presence of M1- and M2-dominant
macrophages side by side in fibrotic lesions in human and mouse
livers, indicating that apparently both are necessary in fibrotic
responses. At least two questions remain: (1) where do these
macrophages come from, meaning are they derived from incoming monocytes and are thus bone marrow-derived or do they
develop from tissue-resident Kupffer cells that are embryonic in
origin (42). Unfortunately, our study cannot answer this question, as there are no markers discovered yet that can reliably
distinguish bone marrow-derived from embryonic macrophages.
Previous studies showed that monocytes do infiltrate the liver
during fibrogenesis and resolution and also that Kupffer cells do
proliferate during injury (43, 44). Understanding the dynamics
of all these different macrophages during fibrogenesis/resolution
and their interactions is a subject of intense research interest.
(2) How these macrophage phenotypes interact with each other
and with other resident cells to enhance or dissolve fibrosis. Song

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Beljaars et al.

et al. (17) showed that M2-dominant macrophages increased the
proliferation index and collagen synthesis of co-cultivated WI-38
fibroblasts, while M1-dominant macrophages markedly reduced
collagen production by these cells. Most in vitro studies suggest
that M2 activation results in enhanced fibrogenesis, while M1 activation inhibits fibrogenesis through antifibrogenic or fibrolytic
factors. Just recently, Lopez-Navarrette (18) showed the importance of M2-dominant macrophages in promoting fibrogenesis in
a CCl4-induced model of liver fibrosis in which Kupffer cells were
stimulated to polarize to an M2-dominant phenotype after hepatic
inoculation of Taenia crassiceps larvae. Our results also suggest a more pro-fibrotic character of M2-dominant macrophages,
because M2 markers were present in fibrotic lesions in human and
mouse livers, but were nearly absent in the livers during resolution
of fibrosis.
Recently, Ramachandran et al. (6) suggested a restorative role
for macrophages during resolution of fibrosis after cessation of
CCL4 intoxications using flow cytometry. The persistence of
M1-dominant macrophages during resolution in our studies indicates that this restorative phenotype may have a more M1 bias.
M1-dominant macrophages have been reported to be major producers of various MMPs and MMP-producing macrophages were
previously reported to be present during liver regeneration in mice
(3, 9, 16). However, M2-dominant macrophages can also express
MMPs [most notably MMP12 (45)] and were found to be important cells for efferocytosis and phagocytosis of matrix debris (16,
21, 46–48). These characteristics of M2-dominant cells may also
be necessary during the resolution phase. The reason we do not
see M2-dominant macrophages anymore in our resolution model
may be caused by the fact that the resolution is ongoing (based
on reduced hepatic collagen deposition) and these functions of
M2-dominant macrophages may have less important. Studying
macrophage phenotype localizations at more time points during
resolution may shed more light on the specific dynamics of the
macrophage phenotypes during resolution.
In conclusion, using a set of established as well as recently identified markers we now clearly show local accumulation of both M1and M2-dominant macrophages in fibrotic septa of mouse and
human end-stage cirrhotic livers. This provides a basis for further
exploring the different activities of these various macrophage phenotypes during liver fibrosis and resolution of fibrosis. The observation that during liver remodeling M1-dominant macrophages
may persist and M2-dominant macrophages may disappear indicates that different combinations of M1 versus M2-dominant
macrophages may play a key role in fibrogenesis and resolution.
Manipulation of their balance may therefore be of therapeutic
value.

ACKNOWLEDGMENTS
We gratefully acknowledge the surgeons of the Department
of Hepato-Pancreato-Biliary Surgery and Liver Transplantation
(University Medical Center Groningen, Groningen, The Netherlands) for providing us with healthy and cirrhotic human liver
tissue. In addition, we thank several students of Pharmacy (J. Lodder, K. Feenstra, L. Keyzer, E. Alons, and N. Aughsteen) for their
practical contributions to this paper.

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Localization of liver macrophage subsets

REFERENCES
1. Pinzani M, Rombouts K, Colagrande S. Fibrosis in chronic liver diseases: diagnosis and management. J Hepatol (2005) 42(Suppl 1):S22–36. doi:10.1016/j.
jhep.2004.12.008
2. Friedman SL. Evolving challenges in hepatic fibrosis. Nat Rev Gastroenterol
Hepatol (2010) 7(8):425–36. doi:10.1038/nrgastro.2010.97
3. Ramachandran P, Iredale JP. Macrophages: central regulators of hepatic fibrogenesis and fibrosis resolution. J Hepatol (2012) 56(6):1417–9. doi:10.1016/j.jhep.
2011.10.026
4. Heymann F, Trautwein C, Tacke F. Monocytes and macrophages as cellular targets in liver fibrosis. Inflamm Allergy Drug Targets (2009) 8(4):307–18.
doi:10.2174/187152809789352230
5. Wynn TA, Barron L. Macrophages: master regulators of inflammation and fibrosis. Semin Liver Dis (2010) 30(3):245–57. doi:10.1055/s-0030-1255354
6. Ramachandran P, Pellicoro A, Vernon MA, Boulter L, Aucott RL, Ali A, et al.
Differential Ly-6C expression identifies the recruited macrophage phenotype,
which orchestrates the regression of murine liver fibrosis. Proc Natl Acad Sci U
S A (2012) 109(46):E3186–95. doi:10.1073/pnas.1119964109
7. Sica A, Invernizzi P, Mantovani A. Macrophage plasticity and polarization in liver
homeostasis and pathology. Hepatology (2014) 59(5):2034–42. doi:10.1002/hep.
26754
8. Duffield JS, Forbes SJ, Constandinou CM, Clay S, Partolina M, Vuthoori S, et al.
Selective depletion of macrophages reveals distinct, opposing roles during liver
injury and repair. J Clin Invest (2005) 115(1):56–65. doi:10.1172/JCI22675
9. Fallowfield JA, Mizuno M, Kendall TJ, Constandinou CM, Benyon RC, Duffield
JS, et al. Scar-associated macrophages are a major source of hepatic matrix
metalloproteinase-13 and facilitate the resolution of murine hepatic fibrosis.
J Immunol (2007) 178(8):5288–95. doi:10.4049/jimmunol.178.8.5288
10. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism and
functions. Immunity (2010) 32(5):593–604. doi:10.1016/j.immuni.2010.05.007
11. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine
system in diverse forms of macrophage activation and polarization. Trends
Immunol (2004) 25(12):677–86. doi:10.1016/j.it.2004.09.015
12. Martinez FO, Sica A, Mantovani A, Locati M. Macrophage activation and polarization. Front Biosci (2008) 13:453–61. doi:10.2741/2692
13. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
14. Mantovani A, Biswas SK, Galdiero MR, Sica A, Locati M. Macrophage plasticity
and polarization in tissue repair and remodelling. J Pathol (2013) 229(2):176–85.
doi:10.1002/path.4133
15. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity.
J Innate Immun (2014). doi:10.1159/000364945
16. Huang WC, Sala-Newby GB, Susana A, Johnson JL, Newby AC. Classical
macrophage activation up-regulates several matrix metalloproteinases through
mitogen activated protein kinases and nuclear factor-kappaB. PLoS One (2012)
7(8):e42507. doi:10.1371/journal.pone.0042507
17. Song E, Ouyang N, Horbelt M, Antus B, Wang M, Exton MS. Influence of alternatively and classically activated macrophages on fibrogenic activities of human
fibroblasts. Cell Immunol (2000) 204(1):19–28. doi:10.1006/cimm.2000.1687
18. Lopez-Navarrete G, Ramos-Martinez E, Suarez-Alvarez K, Aguirre-Garcia J,
Ledezma-Soto Y, Leon-Cabrera S, et al. Th2-associated alternative Kupffer cell
activation promotes liver fibrosis without inducing local inflammation. Int J Biol
Sci (2011) 7(9):1273–86. doi:10.7150/ijbs.7.1273
19. Gibbons M, MacKinnon A, Ramachandran P, Dhaliwal K, Duffin R, PhythianAdams A, et al. Ly6Chi monocytes direct alternatively activated profibrotic
macrophage regulation of lung fibrosis. Am J Respir Crit Care Med (2011)
184(5):569–81. doi:10.1164/rccm.201010-1719OC
20. Wynn TA, Ramalingam TR. Mechanisms of fibrosis: therapeutic translation for
fibrotic disease. Nat Med (2012) 18(7):1028–40. doi:10.1038/nm.2807
21. Boorsma CE, Draijer C, Melgert BN. Macrophage heterogeneity in respiratory
diseases. Mediators Inflamm (2013) 769214:19. doi:10.1155/2013/769214
22. Lopez-Guisa JM, Cai X, Collins SJ, Yamaguchi I, Okamura DM, Bugge TH,
et al. Mannose receptor 2 attenuates renal fibrosis. J Am Soc Nephrol (2012)
23(2):236–51. doi:10.1681/ASN.2011030310
23. Pesce JT, Ramalingam TR, Mentink-Kane MM, Wilson MS, El Kasmi KC, Smith
AM, et al. Arginase-1-expressing macrophages suppress Th2 cytokine-driven
inflammation and fibrosis. PLoS Pathog (2009) 5(4):e1000371. doi:10.1371/
journal.ppat.1000371

September 2014 | Volume 5 | Article 430 | 100

Beljaars et al.

24. Louvet A, Teixeira-Clerc F, Chobert MN, Deveaux V, Pavoine C, Zimmer A,
et al. Cannabinoid CB2 receptors protect against alcoholic liver disease by regulating Kupffer cell polarization in mice. Hepatology (2011) 54(4):1217–26.
doi:10.1002/hep.24524
25. Randolph GJ. Proliferating macrophages prevail in atherosclerosis. Nat Med
(2013) 19(9):1094–5. doi:10.1038/nm.3316
26. Krausgruber T, Blazek K, Smallie T, Alzabin S, Lockstone H, Sahgal N, et al. IRF5
promotes inflammatory macrophage polarization and TH1-TH17 responses.
Nat Immunol (2011) 12(3):231–8. doi:10.1038/ni.1990
27. Uhlen M, Oksvold P, Fagerberg L, Lundberg E, Jonasson K, Forsberg M,
et al. Towards a knowledge-based human protein atlas. Nat Biotechnol (2010)
28(12):1248–50. doi:10.1038/nbt1210-1248
28. Vos TA, Van Goor H, Tuyt L, De Jager-Krikken A, Leuvenink R, Kuipers F, et al.
Expression of inducible nitric oxide synthase in endotoxemic rat hepatocytes is
dependent on the cellular glutathione status. Hepatology (1999) 29(2):421–6.
doi:10.1002/hep.510290231
29. Raes G, Van den Bergh R, De Baetselier P, Ghassabeh GH, Scotton C, Locati M,
et al. Arginase-1 and Ym1 are markers for murine, but not human, alternatively
activated myeloid cells. J Immunol (2005) 174(11):6561. doi:10.4049/jimmunol.
174.11.6561
30. Raes G, De Baetselier P, Noel W, Beschin A, Brombacher F, Hassanzadeh Gh
G. Differential expression of FIZZ1 and Ym1 in alternatively versus classically
activated macrophages. J Leukoc Biol (2002) 71(4):597–602.
31. Van Gorp H, Delputte PL, Nauwynck HJ. Scavenger receptor CD163, a Jackof-all-trades and potential target for cell-directed therapy. Mol Immunol (2010)
47(7–8):1650–60. doi:10.1016/j.molimm.2010.02.008
32. Melgert BN, ten Hacken NH, Rutgers B, Timens W, Postma DS, Hylkema MN.
More alternative activation of macrophages in lungs of asthmatic patients.
J Allergy Clin Immunol (2011) 127(3):831–3. doi:10.1016/j.jaci.2010.10.045
33. Martinez FO, Helming L, Milde R, Varin A, Melgert BN, Draijer C, et al. Genetic
programs expressed in resting and IL-4 alternatively activated mouse and
human macrophages: similarities and differences. Blood (2013) 121(9):e57–69.
doi:10.1182/blood-2012-06-436212
34. Beljaars L, Molema G, Weert B, Bonnema H, Olinga P, Groothuis GM, et al.
Albumin modified with mannose 6-phosphate: a potential carrier for selective
delivery of antifibrotic drugs to rat and human hepatic stellate cells. Hepatology
(1999) 29(5):1486–93. doi:10.1002/hep.510290526
35. Iredale JP. Models of liver fibrosis: exploring the dynamic nature of inflammation and repair in a solid organ. J Clin Invest (2007) 117(3):539–48.
doi:10.1172/JCI30542
36. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol (2011) 11(11):723–37. doi:10.1038/nri3073
37. Hanania R, Sun HS, Xu K, Pustylnik S, Jeganathan S, Harrison RE. Classically
activated macrophages use stable microtubules for matrix metalloproteinase9 (MMP-9) secretion. J Biol Chem (2012) 287(11):8468–83. doi:10.1074/jbc.
M111.290676
38. Belkin AM. Extracellular TG2: emerging functions and regulation. FEBS J (2011)
278(24):4704–16. doi:10.1111/j.1742-4658.2011.08346.x
39. Fesus L, Piacentini M. Transglutaminase 2: an enigmatic enzyme with
diverse functions. Trends Biochem Sci (2002) 27(10):534–9. doi:10.1016/S09680004(02)02182-5

www.frontiersin.org

Localization of liver macrophage subsets

40. Fesus L, Szondy Z. Transglutaminase 2 in the balance of cell death and survival.
FEBS Lett (2005) 579(15):3297–302. doi:10.1016/j.febslet.2005.03.063
41. Popov Y, Sverdlov DY, Sharma AK, Bhaskar KR, Li S, Freitag TL, et al. Tissue
transglutaminase does not affect fibrotic matrix stability or regression of liver
fibrosis in mice. Gastroenterology (2011) 140(5):1642–52. doi:10.1053/j.gastro.
2011.01.040
42. Guilliams M, Ginhoux F, Jakubzick C, Naik SH, Onai N, Schraml BU, et al.
Dendritic cells, monocytes and macrophages: a unified nomenclature based on
ontogeny. Nat Rev Immunol (2014) 14(8):571–8. doi:10.1038/nri3712
43. Liaskou E, Zimmermann HW, Li KK, Oo YH, Suresh S, Stamataki Z,
et al. Monocyte subsets in human liver disease show distinct phenotypic and
functional characteristics. Hepatology (2013) 57(1):385–98. doi:10.1002/hep.
26016
44. Tacke F, Zimmermann HW. Macrophage heterogeneity in liver injury and fibrosis. J Hepatol (2014) 60(5):1090–6. doi:10.1016/j.jhep.2013.12.025
45. Nelson MP, Christmann BS, Dunaway CW, Morris A, Steele C. Experimental pneumocystis lung infection promotes M2a alveolar macrophage-derived
MMP12 production. Am J Physiol Lung Cell Mol Physiol (2012) 303(5):L469–75.
doi:10.1152/ajplung.00158.2012
46. Madala SK, Pesce JT, Ramalingam TR, Wilson MS, Minnicozzi S, Cheever
AW, et al. Matrix metalloproteinase 12-deficiency augments extracellular
matrix degrading metalloproteinases and attenuates IL-13-dependent fibrosis.
J Immunol (2010) 184(7):3955–63. doi:10.4049/jimmunol.0903008
47. Dalli J, Serhan CN. Specific lipid mediator signatures of human phagocytes:
microparticles stimulate macrophage efferocytosis and pro-resolving mediators.
Blood (2012) 120(15):e60–72. doi:10.1182/blood-2012-04-423525
48. Morimoto K, Janssen WJ, Terada M. Defective efferocytosis by alveolar
macrophages in IPF patients. Respir Med (2012) 106(12):1800–3. doi:10.1016/j.
rmed.2012.08.020

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.

Received: 10 July 2014; paper pending published: 02 August 2014; accepted: 22 August
2014; published online: 08 September 2014.
Citation: Beljaars L, Schippers M, Reker-Smit C, Martinez FO, Helming L, Poelstra K
and Melgert BN (2014) Hepatic localization of macrophage phenotypes during fibrogenesis and resolution of fibrosis in mice and humans. Front. Immunol. 5:430. doi:
10.3389/fimmu.2014.00430
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
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September 2014 | Volume 5 | Article 430 | 101

REVIEW ARTICLE
published: 07 October 2014
doi: 10.3389/fimmu.2014.00489

Functional relationship between tumor-associated
macrophages and macrophage colony-stimulating factor
as contributors to cancer progression
Damya Laoui 1,2 , Eva Van Overmeire 1,2 , Patrick De Baetselier 1,2 , Jo A. Van Ginderachter 1,2 and Geert Raes 1,2 *
1
2

Myeloid Cell Immunology Laboratory, VIB, Brussels, Belgium
Unit of Cellular and Molecular Immunology, Vrije Universiteit Brussel, Brussels, Belgium

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Lidija Klampfer, Montefiore Medical
Center, USA; Albert Einstein Cancer
Center, USA
Robert Adam Harris, Karolinska
Institutet, Sweden
*Correspondence:
Geert Raes, Unit of Cellular and
Molecular Immunology, VIB Myeloid
Cell Immunology Laboratory, Vrije
Universiteit Brussel, Pleinlaan 2,
Brussels 1050, Germany
e-mail: geert.raes@vub.ac.be

The current review article describes the functional relationship between tumor-associated
macrophages (TAM) as key cellular contributors to cancer malignancy on the one hand and
macrophage-colony-stimulating factor (M-CSF or CSF-1) as an important molecular contributor on the other. We recapitulate the available data on expression of M-CSF and the M-CSF
receptor (M-CSFR) in human tumor tissue as constituents of a stromal macrophage signature and on the limits of the predictive and prognostic value of plasma M-CSF levels. After
providing an update on current insights into the nature of TAM heterogeneity at the level of
M1/M2 phenotype andTAM subsets, we give an overview of experimental evidence, based
on genetic, antibody-mediated, and pharmacological disruption of M-CSF/M-CSFR signaling, for the extent to which M-CSFR signaling can not only determine theTAM quantity, but
can also contribute to shaping the phenotype and heterogeneity of TAM and other related
tumor-infiltrating myeloid cells (TIM). Finally, we review the accumulating information on
the – sometimes conflicting – effects blocking M-CSFR signaling may have on various
aspects of cancer progression such as tumor growth, invasion, angiogenesis, metastasis,
and resistance to therapy and we thereby discuss in how far these different effects actually
reflect a contribution of TAM.
Keywords: M-CSF, CSF-1, M-CSFR, CSF-1R, tumor-associated macrophages, M1, M2, cancer progression

INTRODUCTION
CANCER MALIGNANCY

Ca ncer is a complex multi-step process, in which normal cells
acquire a certain growth advantage via a process analogous to Darwinian evolution. These cellular changes can occur under many
different circumstances, which contributes to the heterogeneity
and variability of the occurrence, development, and outcome of
neoplastic disease (1). The traits required for malignant growth
include self-sufficiency from external growth signals, insensitivity to negative growth signals, resistance to apoptosis, limitless
replicative potential, sustained angiogenesis, acquisition of tissue
invasiveness, and metastasis. Recently, genetic instability, altered
Abbreviations: DAMP, damage-associated molecular pattern; EGF, epidermal
growth factor; Fizz1, found in inflammatory zone 1; GAST, gastrointestinal stromal tumor; G-CSF, granulocyte-colony-stimulating factor; GM-CSF, granulocytemacrophage colony-stimulating factor; HUVEC, human umbilical vein endothelial cell; mAb, monoclonal antibody; M-CSF, macrophage-colony-stimulating factor; M-CSFR, M-CSF receptor; Mgl2, macrophage galactose-type C-type lectin-2;
MMP9, matrix metalloproteinase 9; MMR, macrophage mannose receptor; MOMDSC, monocytic myeloid-derived suppressor cells; PAMP, pathogen-associated
molecular pattern; PMN-MDSC, polymorphonuclear myeloid-derived suppressor
cells; PyMT, polyomavirus middle T oncogene; RNI, reactive nitrogen intermediates;
ROS, reactive oxygen species; RT2, RIP1-Tag2; SR-A, scavenger receptor-A; TAM,
tumor-associated macrophages; TEM, Tie2-expressing monocytes/macrophages;
TGF-β, transforming growth factor-β; TIL, tumor-infiltrating lymphocytes; TIM,
tumor-infiltrating myeloid cells; TMEM, tumor microenvironment of metastasis;
VEGF, vascular endothelial growth factor.

Frontiers in Immunology | Molecular Innate Immunity

energy metabolism, the capacity to evade elimination by the
immune system, including active immune suppression, as well as
smoldering, non-resolving inflammation, leading to accumulation
of random genetic alterations in cancer cells due to inflammatory
mediators, have been established as additional hallmarks of cancer (2–6). In this regard, tumors consist not only of neoplastic
cells, but should be considered as organ-like structures in which
a complex bidirectional interplay exists between transformed and
non-transformed cells. The malignant potential of transformed
cells requires an apt support structure from the stroma, which can
consist of fibroblasts, adipocytes, blood, and lymph vessels, but
may also be considerably infiltrated by a wide range of immune
cells, such as tumor-associated macrophages (TAM) (7).
PRO- AND ANTI-TUMORAL ROLES OF TAM

Tumor-associated macrophages are the predominant leukocytes
infiltrating solid tumors and can represent up to 50% of the
tumor mass. The clinical significance of these cells is illustrated
by the significant link between TAM number and density and a
poor prognosis in 80% of the reported studies. The main exception to this general trend seems to be colorectal cancer, for which a
high TAM density is significantly associated with enhanced overall
survival (8–10).
Tumor-associated macrophages stimulated by TLR ligands,
agonistic anti-CD40, or IFN-γ were shown to have important
anti-tumoral activities, provided that cancer cell phagocytosis is

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not inhibited by CD47 expression on the malignant cells, which
is a “don’t-eat me signal” (11, 12). In addition, pro-inflammatory
macrophages are able to eliminate cancer cells via the production
of reactive oxygen species (ROS) and reactive nitrogen intermediates (RNI) and secrete chemokines that recruit and prime T cells
toward an anti-tumor phenotype in some cancer types, resulting
in retarded tumor growth or tumor regression (13–17).
Whereas TAM can exert anti-tumoral activities, the ambiguous role of macrophages in tumor progression is reflected in the
finding that TAM can also actively contribute to each stage of cancer development and progression (Figure 1A). They can promote
cancer cell proliferation, invasion, metastasis, and angiogenesis by
releasing cytokines, growth factors, extracellular matrix-degrading
enzymes, and angiogenic factors including vascular endothelial
growth factor (VEGF), prokineticin (Bv8), and matrix metalloproteinase 9 (MMP9). TAM also inhibit cytotoxic T-cell activity by the
secretion of suppressive cytokines such as IL-10 and transforming
growth factor-β (TGF-β), high levels of arginase activity, and the
production of ROS or RNI (18–22). Finally, TAM contribute to
tumor relapse following tumor irradiation and anti-angiogenic
therapy (23).
It seems unlikely that the diverse anti-tumoral and tumorpromoting activities of TAM are performed by a single cell type,
and the existence of distinct TAM subpopulations, linked to different intratumoral microenvironments, has been predicted (10, 24).
Depending on the cancer type, the stage of tumor progression and
location within the tumor tissue, molecularly and functionally distinct TAM subpopulations coexist in tumors (25–27). This TAM
heterogeneity likely reflects the inherent plasticity of macrophages
in response to (micro-)environmental triggers.
MACROPHAGE PLASTICITY

Macrophages have a remarkable plasticity and are found in all
tissues, where they display great anatomical and functional diversity. They are implicated in a spectrum of roles required for tissue
homeostasis, ranging from host defense against infectious agents,
to tissue development, wound healing, and immune regulation.
Accordingly, macrophages are able to adopt diverse phenotypes
or activation states in response to environmental cues, such as
cytokines, pathogen-associated molecular patterns (PAMP), tissue damage-associated molecular patterns (DAMP), and other
immune stimuli (7, 28, 29).
Macrophage activation is conventionally categorized on a linear scale, in which the two opposing phenotypes are referred to
as the classical (M1) versus alternative (M2) macrophage activation state, originally mirroring the Th1 versus Th2 nomenclature
(30–34). M1 macrophage activation is driven by exposure to IFNγ and TLR ligands. These macrophages secrete pro-inflammatory
cytokines (such as IL-12, IL-1, IL-6, TNF, ROS, RNI), promote Th1
responses, exert cytotoxic activities, and are involved in defense
against bacterial infections and intracellular pathogens. The M2
activation state refers to macrophages that are not M1 activated
and comprise various activation states, induced by a wide array
of different stimuli, leading to different macrophage classification
systems by different authors. These stimuli include Th2 cytokines
(such as IL-4 and IL-13), anti-inflammatory cytokines (such as IL10 and TGF-β), hormones (such as glucocorticoids), and immune

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TAM and M-CSF in cancer progression

complexes. Consequently, non-M1 macrophages have very diverse
functions, ranging from parasite control to immune suppression,
wound repair, tissue remodeling, and angiogenesis. Features of
these non-M1 macrophages are the low secretion levels of proinflammatory cytokines, high expression of macrophage mannose
receptor (MMR) and scavenger receptor-A (SR-A), and an arginine metabolism shifted toward the production of ornithine and
polyamines by arginase (35–39). Although the M1/M2 classification has proven useful, any form of classification underscores
the complexity of the in vivo situation, in which numerous stimuli
interact to define the final differentiated state and mixed functional
profile of macrophages (40–42). In this context, new nomenclature and experimental guidelines for dealing with macrophage
activation and polarization have very recently been proposed (43).
M-CSF AS DRIVER OF BOTH DIFFERENTIATION AND PHENOTYPIC
POLARIZATION OF MACROPHAGES

The myelopoietic growth factors macrophage-colony-stimulating
factor (M-CSF, also known as CSF-1), granulocyte-macrophagecolony-stimulating factor (GM-CSF) and IL-34 are major
cytokines in controlling the proliferation, differentiation, and
functional regulation of monocytes, macrophages, and dendritic
cells [reviewed in Ref. (44)]. M-CSF and IL-34 are produced by
a variety of stromal and epithelial cell types and signal through
the M-CSF receptor (M-CSFR, CSF-1R, or CD115), a type III
receptor tyrosine kinase (45), encoded by the Csf1r/c-fms protooncogene (46, 47), that seems to be mainly restricted to cells of the
mononuclear phagocyte lineage (48).
Especially, M-CSF instructs the myeloid fate in single
hematopoietic stem cells, by inducing the myeloid master regulator transcription factor PU.1 (49). Embryonic yolk sac-derived
precursors and fetal liver monocytes have been found to give rise to
many tissue-resident macrophages that seed all tissues prenatally
and are maintained via self-renewal throughout adult life (50). The
importance of M-CSF for establishing and maintaining the tissueresident macrophage pool is illustrated by the M-CSF-deficient
osteopetrotic (op/op) mouse, which not only suffers from congenital osteopetrosis due to a severe deficiency of osteoclasts, but
also features severe defects in many tissue-resident macrophage
populations (51). Besides having effects on macrophage precursor
differentiation, M-CSF is known to stimulate macrophage survival (52) and self-renewal during steady-state and inflammation
(53). However, macrophage populations in distinct tissues are differentially affected by the M-CSF deficiency. For example, skin
Langerhans cells and brain microglia were seemingly normal in
op/op mice, but were largely absent from M-CSFR-deficient mice,
a finding which has been explained by the trophic role of IL-34,
whose production is restricted to keratinocytes and neurons under
steady-state (54, 55).
In addition to a role in resident tissue macrophage differentiation and maintenance, M-CSFR signaling has also been
assigned an important role in polarization of macrophage activation, flowing from the observation of significant differences
in the transcriptomes of the macrophage populations primarily generated with the use of M-CSF or GM-CSF. M-CSF-driven
macrophage differentiation leads to the expression of a substantial part of the M2 transcriptome, including expression of MMR

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FIGURE 1 | Scheme of the possible effects of TAM and of M-CSFR
blockade on cancer progression. (A) Possible tumor-promoting effects of
TAM. TAM can promote cancer progression and reduce the efficacy of
radiotherapy, chemotherapy, immunotherapy, and anti-angiogenic therapy by a
combination of different mechanisms. TAM can contribute to enhanced
cancer cell numbers by (Aa) inhibiting anti-tumor immune responses and via
(Ab) stimulation/maintenance of cancer cell proliferation. TAM can also exert
pro-angiogenic activities by enhancing (Ac) angiogenic switching and (Ad)
endothelial cell proliferation. Finally, TAM can contribute to cancer malignancy
by facilitating (Ae) cancer cell invasion and (Af) seeding, extravasation,
survival, and subsequent proliferation of cancer cells at metastatic sites.
(B) Possible effects of M-CSFR signaling blockade on cancer progression.

and SR-A, while GM-CSF rather induces a pro-inflammatory
M1-type of activation (49, 56–58). As such, blocking M-CSFR
signaling in myometrial macrophages stimulated the occurrence
of an M1-like MHC-IIhigh population at the expense of M2like MHC-IIlow macrophages in the pregnant mouse uterus (59).

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TAM and M-CSF in cancer progression

Depending on the tumor type/model and the blocking agents used to impede
M-CSFR signaling, M-CSFR blockade has in most cases been reported to
attenuate cancer progression and/or synergistically enhance the effect of
chemo-, radio-, and/or immunotherapy via various effects, including (Ba)
promotion of tumor-infiltrating lymphocytes (TIL) recruitment and/or
activation, (Bb) enhanced phagocytosis/killing of cancer cells, (Bc) a delayed
angiogenic switch, (Bd) reduced density of proliferating endothelial cells, (Be)
inhibition of both TAM and cancer cell migration and invasion, (Bf) reduced
metastasis. In some cases, (Bg) reduction of tumor weight and primary tumor
growth has been reported. A number of studies have attributed these effects
to (i) ablation of TAM numbers and/or (ii) phenotypic reprograming of TAM
from tumor promoting (often M2-like) TAM to anti-tumor (often M1-like) TAM.

The same study also demonstrated an important role for M-CSF
in mediating monocyte extravasation to the tissue, via M-CSFdependent upregulation of the chemokine CCL2, adding further
evidence to the notion that M-CSF affects macrophage dynamics
at multiple levels.

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Since high M-CSF levels are frequently found in tumor-bearing
hosts, the M-CSFR signaling could also play a role in shaping the
TAM pool and regulating their activation state.

ASSOCIATION OF M-CSF AND M-CSFR LEVELS WITH
HUMAN CANCER PROGRESSION
M-CSF, M-CSFR, AND/OR M-CSF RESPONSE SIGNATURE EXPRESSION
IN TUMOR TISSUE

Various studies have documented analyses in which attempts were
made to correlate clinical cancer patient parameters such as disease
staging and survival with protein and/or mRNA expression levels
of M-CSF, M-CSFR, and/or M-CSF response genes. The latter were
thereby in turn considered to correlate with the presence of high
levels of TAM and thus to represent a macrophage signature.
High M-CSF expression levels, as detected via IHC on tissue
sections, have been reported to associate with higher histological tumor grading and in many cases also with more frequent
metastases and poor prognosis in various cancer types, including breast cancer (60), serous and mucinous ovarian epithelial tumors (61), endometrioid carcinomas (62), and papillary
renal cell carcinoma (63). In gynecological and non-gynecological
leiomyosarcoma, expression of individual markers such as MCSF was found to show at least a trend for correlation with
poor outcome, but only the co-expression of M-CSF and three
M-CSF-response genes (CTSL1, FCGR3a, and CD163) was independently associated with a worse survival in a multivariate
analysis (64).
Studying the expression of M-CSFR via IHC in a large cohort of
clinical breast cancer specimens using tissue microarrays revealed
that M-CSFR expression was strongly associated with nodal status
(65). In fact, in multivariate analysis, M-CSFR was not independent of nodal status as a predictor of survival. The study also
revealed that M-CSFR expression was associated with decreased
overall survival in non-metastatic breast cancer patients, but not
in node-positive patients (65). Of note, in a recent manuscript, low
levels of the M-CSFR gene were reported to predict worse overall
survival based on online survival analysis tools allowing an evaluation of the prognostic value of genes in breast cancer patients using
microarray data (66). In another recent study, a high number of
tumor stromal cells – but not the cancer cells themselves – expressing M-CSFR was found to be an independent prognostic marker
for lower event free survival and lower overall survival in classical
Hodgkin lymphoma (67).
In line with variable results on association of M-CSFR expression with overall survival among different cancer types and patient
groups, one report of a study using gene microarray and tissue
microarray analyses for evaluating the prognostic value of an MCSF response signature in breast cancer patients, mentioned a
complex relationship of the signature with survival. Indeed, when
patients were substratified in subsets, the M-CSF response signature was associated with poor prognosis among low-grade tumors
and showed a trend for an association with improved prognosis among estrogen-receptor-negative tumors and among tumors
with a TP53 mutation gene-expression signature (68). This variability in the association of M-CSF/macrophage signatures with
clinical parameters points to the need to properly identify patient
groups in which an M-CSF/macrophage signature correlates with

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TAM and M-CSF in cancer progression

worse prognosis and which are thus most likely to benefit from
M-CSF/macrophage-targeted therapies.
It should also be remarked that the presence of an MCSF/macrophage signature is not a uniform feature in all cancer
patients. In fact, gene microarray and tissue microarray analyses
revealed M-CSF and M-CSF response signature genes to be present
in 17–25% of breast cancers (68) and in about 27% of myoinvasive
endometrioid carcinomas (62). Yet, in the latter case, concordance between the expression of the M-CSF signature in primary
endometrioid carcinomas and in their corresponding lymph node
metastases was reported. Moreover, in case of breast carcinoma,
expression of the M-CSF signature was not only detected in some
patients in case of invasive ductal carcinoma, but was already
detected at the stage of ductal carcinoma in situ. Also in that case,
a correlation was found between the presence of the M-CSF signature in ductal carcinoma in situ and in invasive ductal carcinoma
within the same patient (69). This conservation of the expression
of the M-CSF signatures upon disease progression is promising
when considering targeting of the M-CSF pathway as a therapeutic
option for invasive and/or metastatic disease, and suggests that the
presence of a M-CSF/macrophage signature in the primary tumor
may be useful for patient stratification to identify those patients
who are most likely to respond to M-CSF/macrophage-targeted
therapies.
CIRCULATING M-CSF

In some cases, M-CSF is produced to such high levels in cancer
patients that it can be detected systemically. Overall trends from
studies in patients with newly diagnosed breast tumors indicate
that circulating plasma M-CSF levels are not higher in patients
with localized tumors than in controls, but are elevated in patients
with regionally advanced disease and distant metastases (70, 71).
Median M-CSF levels were also reported to be dramatically higher
in patients with newly diagnosed tumors of the head and neck,
in men with prostate cancer metastatic to bone and women with
advanced metastatic breast cancer than those seen in patients with
newly diagnosed breast tumors (70).
Prospective studies of the prognostic value of serum M-CSF
levels have yielded conflicting results. One study on 471 women
with pre-invasive and invasive breast carcinoma reported no significant association between pre-treatment plasma levels of MCSF and overall/relapse free survival at a median follow up of
5.6 years. In this study, patients were classified into three groups
based on the level of initial M-CSF, using median and twice median
plasma values as cut-off points (70). In contrast, a recent study
of 572 women with early breast cancer, that had not undergone
local or systemic anti-cancer treatment prior to serum collection,
revealed significantly poorer outcome at a median follow-up of
5.2 years in patients with above-median M-CSF concentrations as
compared to those with below-median M-CSF concentrations.
In this study population, log M-CSF serum concentrations at
study enrollment were predictive of poor survival in both univariate analysis, as well as multivariate analysis adjusted for age,
tumor size, nodal status, and tumor grade (71). In a retrospective, nested case–control study of breast cancer risk in 726 breast
cancer patients and 734 matched controls with no cancer history, the association of circulating M-CSF levels with the risk of

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developing breast cancer was found to vary by menopausal status. High M-CSF levels were associated with a reduced risk of
premenopausal breast cancer, whereas they were associated with
an increased risk of postmenopausal breast cancer (72). Interestingly, in the aforementioned prospective study, the reported poorer
outcome in patients with above-median M-CSF concentrations
was confined to postmenopausal women, while no such effect was
observed in premenopausal women with early breast cancer (71).
Overall, although the practical use of serum M-CSF levels as
prognostic factor for cancer risk and/or outcome may be complicated by a high heterogeneity among patient groups and difficulties
in determining optimal cut-off levels for plasma M-CSF, these
results do suggest that, at least in some patient groups, M-CSF
and M-CSF-dependent macrophages may be directly involved in
tumor progression and malignant behavior and thus constitute
interesting therapeutic targets.

TAM PHENOTYPIC AND SUBPOPULATION HETEROGENEITY
Originally, TAM were characterized as M2-like cells, proficient in
inducing trophic functions like tumor angiogenesis, invasion, proliferation, and expressing the anti-inflammatory cytokine IL-10.
These cells were also reported to express M2-specific markers like
arginase-1, macrophage galactose-type C-type lectin-2 (Mgl2),
found in inflammatory zone 1 (Fizz1), Ym1, TGF-β, SR-A, and
MMR (73–76). However, some studies of chronic inflammationinduced cancer indicate the presence of TAM with an inflammatory M1-like phenotype, releasing inflammatory cytokines like
IL-12, TNF, IL-6, and IL-1, or with overlapping M1 and M2
characteristics (77–79).
A dynamic switch in the phenotype of TAM during tumor
progression might account for the mixed activation state of TAM
subsets found in different established tumors. Indeed, in some
models, tumor progression is associated with a switch from M1like to M2-like TAM (80, 81). Hence, M2-like TAM can be linked
to tumor promotion and their presence is indicative of poor
prognosis (82, 83). Accordingly, a high M1/M2 TAM ratio has
been associated with extended survival in many cancer types (84).
Moreover, inhibition of monocyte differentiation to M2-like TAM
through inhibition of NF-κB signaling, results in an M1-like phenotype and reduced tumor growth (85). Hence, a picture emerges
whereby M2-like TAM are pro-tumoral, and M1-like TAM exert
anti-tumoral activities.
Accumulating evidence suggests that different TAM activation states found within the same tumor may reflect responses
to divergent local microenvionmental signals (86). As previously mentioned, tumors are complex organoid structures containing peritumoral stroma, perivascular regions, and hypoxic
regions, which can all be populated by TAM, albeit with a different molecular profile and exerting specialized functions (86–
89). Different studies, using state-of-the-art microscopy, clearly
illustrated the existence of at least two distinct microenvironments in the same tumor, which were both infiltrated by TAM
subsets. TAM residing in avascular regions are sessile, have a
high phagocytic capacity, and express high levels of many prototypical M2 markers such as MMR. In contrast, perivascular
TAM are migratory, are not able to ingest dextran, have a less
pronounced M2-profile and produce epidermal growth factor

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TAM and M-CSF in cancer progression

(EGF), which attracts M-CSF-producing cancer cells, resulting
in migration and intravasation of cancer cells (90–94). In line
with these findings, differentially activated macrophages within
the same tumors, residing in distinctively oxygenated tumor
regions, could be discriminated based on the expression of MHCII. MHC-IIhigh TAM are excluded from hypoxic avascular areas
and more M1 oriented, while hypoxic MHC-IIlow TAM express
higher levels of M2-associated markers and are more angiogenic
(25–27, 95). However, increasing the oxygenation of neoplastic
lesions by vessel normalization in PHD2-haplodeficient mice was
recently found not to alter the expression of the most prominent M2 markers, such as MMR, IL-4Rα, and Arginase-1. Rather,
reduced hypoxia down-regulated a subset of genes and proteins
involved in glycolysis, angiogenesis, and metastasis, thereby lowering their angiogenic functions, specifically and solely in the
hypoxic MHC-IIlow TAM subset (27). Hence, hypoxia is not the
main driver of TAM differentiation, but M2-like TAM preferentially home to hypoxic areas where the pro-tumoral activities
of these cells are promoted. The importance of the intratumoral TAM location in shaping the phenotype of TAM subpopulations was further confirmed by a study showing that
Neuropilin-1 deficiency in macrophages prohibits their migration to hypoxic tumor areas, resulting in an increased inflammatory phenotype and the initiation of an anti-tumor immune
response (96).

EFFECTS OF M-CSFR SIGNALING ON NUMBERS AND
PHENOTYPE OF TAM AND OTHER TIM
The critical role of M-CSF in the turn-over of TAM is reflected
in the drastic reduction in macrophages in the primary tumor
at different stages of tumor progression to malignancy that
has been observed in the absence of M-CSF in osteopetrotic
op/op mice (97, 98). Conversely, restoration of M-CSF signaling via transgenic expression of M-CSF in the mammary
epithelium led to enhanced numbers of macrophages in primary mammary tumors (97). Similarly, strong reductions in
the number of TAM have been reported in various tumor
models upon blocking of M-CSF/M-CSFR signaling to TAM
using either blocking monoclonal antibodies (mAbs) targeting M-CSF (66, 99) or the extracellular domain of M-CSFR
(100–102) or small molecule inhibitors of the M-CSFR tyrosine kinase activity in order to block the downstream signaling
(102–105) (Figure 2).
Despite numerous reports of the differential effects of M-CSF
versus GM-CSF on macrophage polarization (44, 57), only few
studies directly addressing the effect of M-CSFR blockade on the
M1/M2 activation state and/or subpopulation heterogeneity of
TAM have recently been documented (Table 1).
One study in a mouse model based on subcutaneously inoculated colon carcinoma cells was aimed at evaluating whether
cytokine signaling could induce reprograming of the TAM phenotype in vitro. The authors reported that GM-CSF treatment
in conjunction with suppression of M-CSF signals using siRNA
against the M-CSFR resulted in an altered signal transduction
pathway of TAM, whereby expression of STAT1, STAT5, and STAT6
was increased. In this study, treatment of TAM with GM-CSF,
alone, or in conjunction with suppression of M-CSFR signals,

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FIGURE 2 | Examples of various types of M-CSFR signaling blocking
agents mentioned throughout this manuscript. In some studies,
neutralizing anti-mouse M-CSF mAb has been used for blocking
M-CSF/M-CSFR signaling (66). One study also reported on the use of a
murinized, polyethylene glycol-linked recombinant Fab fragment of the
MCSF1-033 neutralizing rabbit anti-mouse M-CSF antibody (99). Yet,
blocking mAbs targeting the extracellular domains of the M-CSFR have
more frequently been documented for blocking the M-CSF/M-CSFR
signaling axis. Typical examples of the latter that have been used in mouse
tumor models are the rat IgG1 M279 (100) and the rat IgG2A AFS98 (101,
102). A recent report documented the generation of RG7155, a humanized
anti-human M-CSFR IgG1 mAb that inhibits M-CSFR activation (106). And
also the fully human IgG1 anti-human M-CSFR mAb IMC-CS4 is currently in
clinical trials (107). M-CSFR signaling has also been inhibited via

did not alter the TAM expression pattern of M1/M2 marker
molecules (112).
In a study, whereby the tyrosine kinase inhibitor PLX3397
was used as a combination treatment with adoptive cell therapy of melanoma-targeted T cells in a syngeneic mouse model
of BRAFV600E-driven melanoma, PLX3397 as single or combination treatment resulted in a dramatic reduction of TAM and
a skewing of the subpopulation balance in the remaining TAM
from predominant M2-oriented MHC-IIlow to predominant M1oriented MHC-IIhigh macrophages (113). A similar shift in the
relative amount of TAM subpopulations was documented in the
transgenic mouse MMTV-Neu model, in which mammary carcinogenesis is driven by the mammary epithelial restricted expression of the ErbB2/Neu oncogene. Blocking M-CSFR in this model
by using the M-CSFR inhibitor GW2580, led to a significant
reduction in the amount of M2-like F4/80high TAM, which had
moderate levels of MHC-II, and not in the MHC-IIhigh F4/80low
TAM, elucidating a role for M-CSFR signaling in the maintenance or expansion of the M2-like TAM subset (95). A recent

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pharmacological, small molecule inhibitors targeting the intracellular
catalytic domains of the receptor involved in signal transduction. A number
of these tyrosine kinase inhibitors, such as CYC11645, Ki20227, GW2580, or
BLZ945, have been screened for highly selective inhibition of M-CSFR
signaling, very potent IC50 values for M-CSFR and at least a 100-fold lower
inhibitory activity for other tested receptor tyrosine kinases (66, 108–110).
Also the PLX3397 tyrosine kinase inhibitor has been used, which has higher
M-CSFR inhibitory activity as compared to GW2580, but which is less
specific since it inhibits the c-Kit receptor tyrosine kinase with similar
potency as the M-CSFR tyrosine kinase (105). In one study, the actual
contribution of M-CSFR blockade in the effect of PLX3397 has been
assessed by comparing it with the specific cKit tyrosine kinase inhibitor
imatinib and PLX5622, an M-CSFR-specific inhibitor of equal potency to
PLX3397 that does not appreciably inhibit Kit (111).

study in a mouse model of pancreatic ductal adenocarcinoma
confirmed this notion. In this model, treatment with PLX3397 or
a neutralizing anti-M-CSF mAb resulted in a drastic reduction
in TAM (114). Thereby, the authors demonstrated that blocking M-CSF/M-CSFR signaling resulted in preferential depletion
of M2-like MMRhigh TAM, whereas M1-like MMRlow TAM were
much less affected. The observation that the MMRhigh TAM subset
had significantly higher M-CSFR expression levels as compared to
the MMRlow TAM subset further supports the notion that these
M2-like cells may be more dependent on the M-CSF signal. As a
consequence, the gene-expression profile of TAM upon M-CSFR
signaling blockade featured a reduced expression of M2 markers and an increased expression of M1 markers and MHC-II. In
parallel, the TAM phenotype shifted from predominant immunosuppressive properties to improved antigen presentation capacity
(114). In a mouse glioblastoma model, in vivo M-CSFR inhibition
using the small molecule M-CSFR inhibitor BLZ945 was reported
not to result in TAM depletion. Instead, glioma-secreted factors,
including GM-CSF and IFN-γ, facilitated TAM survival in the

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TAM and M-CSF in cancer progression

Table 1 | Documented effects of M-CSFR inhibition on TAM abundance and activation state.
Mouse tumor

Tool used to inhibit

model

M-CSFR signaling

Colon carcinoma

In vitro: siRNA against

Amount of TAM

TAM M1/M2 activation

Unaltered

Unaltered

M-CSFR + GM-CSF
Melanoma

M-CSFR inhibitor:

Effect/outcome

Reference

state
Increased expression of STAT1,

(112)

STAT5, STAT6 in TAM
Skewing from M2

Improved adoptive cell therapy:

PLX3397 + adoptive

Reduced

MHC-IIlow to M1

increased amount and activation

cell therapy

MHC-IIhigh

of tumor-infiltrating lymphocytes;

(113)

reduced tumor growth
Mammary carcinoma

M-CSFR inhibitor:

Reduced (only M2-like

GW2580

MHC-IIlow TAM)

Pancreatic ductal

M-CSFR inhibitor:

Reduced (mainly M2-like

Remaining TAM are less

Increased anti-tumor T cell

adenocarcinoma

PLX3397/neutralizing

MMRhigh TAM)

immunosuppressive,

activity; enhanced response to

better Ag presenting M1

immunotherapy

Repolarization from

Reduced tumor growth

(108)

Increased amount of CD8+ T

(104)

α-M-CSF MAb
Glioblastoma

M-CSFR inhibitor:

Unaltered

BLZ945

Not assessed

Role of M-CSFR in maintenance

(95)

of M2-like TAM
(114)

pro-tumoral M2 to
phagocytic M1 TAM

Cervical and

M-CSFR inhibitor:

Reduced (both

MHC-IIlow

Not assessed

MHC-IIhigh TAM)

mammary carcinoma

BLZ945

and

Pancreatic ductal

M-CSFR inhibitor:

Reduced (mainly M1-like

Remaining TAM are less

Enhanced response to

adenocarcinoma

PLX3397 or GW2580

immunosuppressive

immunosuppressive

chemotherapy; increased CTL

MHC-IIhigh TAM)

context of M-CSFR inhibition and resulted in a repolarization
from pro-tumoral M2 to a highly phagocytic M1 phenotype, with
a decreased expression of M2 markers (108).
Despite the above examples indicating that M-CSFR blockade can shift the balance in TAM subpopulations from tumorpromoting M2-oriented MHC-IIlow or MMRhigh to anti-tumoral
M1-oriented MHC-IIhigh or MMRlow macrophages, conflicting
reports also exist. For example, the M-CSFR inhibitor BLZ945
was reported to result in a decrease in the level of TAM in cervical
and breast carcinomas by attenuating their turn-over rate. Hereby,
similar kinetics of depletion and recovery were observed for both
MHC-IIlow and MHC-IIhigh TAM subpopulations (104). And in
mice bearing transplantable pancreatic ductal adenocarcinomas,
the M-CSFR inhibitors GW2580 or PLX3397 were even reported to
significantly deplete macrophages expressing high levels of MHCII, but not the more M2-oriented MHC-IIlow or Tie2+ TAM (105).
Yet, in the latter case and in contrast to the examples above, the
MHC-IIhigh TAM were found to constitute the predominant TAM
subpopulation and to exert pro-tumoral activities by suppressing anti-tumoral CD8+ T cell responses (105). Therefore, also in
that case, the observed reduction in the level of the predominant
tumor-promoting TAM subpopulation, occurring upon M-CSFR
blockade, resulted in attenuation of cancer malignancy (Table 1).
Further studies will be required to obtain better insights into the
extent and the underlying mechanisms by which M-CSFR signaling and blockade thereof can contribute to shaping the phenotypic
and subpopulation heterogeneity of TAM, thereby re-educating
TAM toward anti-tumoral effector populations, thus contributing

Frontiers in Immunology | Molecular Innate Immunity

cells; reduced tumor growth
(105)

response; reduced metastases

to combating disease progression. It will also be of importance to
assess to what extent the remaining TAM populations detected
after M-CSFR signaling blockade in various cancer types and
tumor models are actually M-CSF-dependent macrophages for
which the depletion was incomplete or the M-CSF dependence
has been (partially) compensated for by other factors. Or do these
remaining cells in some instances represent M-CSF-independent
cells with a distinct lineage origin (such as for example certain dendritic cell types) for which the lineage surface markers
and morphological analysis used in the current studies have not
allowed to discriminate them from macrophages?
Of note, a number of recent publications evaluating the effect of M-CSFR inhibitors such as GW2580
or PLX3397 on various populations of tumor-infiltrating
myeloid cells (TIM) have documented a reduction of not
only mature CD11b+ Ly6G- Ly6Clow F4/80high TAM, but also of
CD11b+ Ly6G- Ly6Chigh F4/80mid cells, resembling the surface
receptor phenotype and morphology of inflammatory (classical)
monocytes or monocytic myeloid-derived suppressor cells (MOMDSC) (105, 114–116). Taking into account the diversity of cell
populations that can express these combinations of surface markers and the fact that an actual suppressive activity of the cells
has not been demonstrated by the authors of most of these studies, we will term these cells MO-MDSC-like cells in the current
review. It makes sense that, as monocyte-lineage cells, these tumorinfiltrating MO-MDSC-like cells are dependent on M-CSFR signaling to a similar extent as mature TAM and these MO-MDSClike cells may in fact very well represent precursors of mature TAM

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Laoui et al.

(117). In contrast to MO-MDSC-like cells, documented effects
of M-CSFR signaling inhibitors on CD11b+ Ly6Ghigh Ly6Clow
cells, resembling the surface receptor phenotype and morphology of immature granulocytes/neutrophils or polymorphonuclear
myeloid-derived suppressor cells (PMN-MDSC) and which we
will term PMN-MDSC-like cells have been more variable. Most
studies revealed no reduction (and sometimes even a limited
increase) in the number of PMN-MDSC-like cells in response
to PLX3397 or GW2580 treatment, for example in mice bearing
murine pancreatic ductal adenocarcinoma (105, 114) or in the 3LL
lung carcinoma model (115). In contrast, PLX3397 was found to
reduce both MO-MDSC-like cells and PMN-MDSC-like cells in
one study in the RM-1 and Myc-CaP prostate cancer models (116).
The variable effect of M-CSFR blockade on PMN-MDSC-like cells
suggests that this effect is most likely indirect and may depend on
other (growth) factors in the tumor microenvironment that are
affected indirectly via the M-CSFR blocking.

EFFECTS OF M-CSFR SIGNALING BLOCKADE ON CANCER
PROGRESSION AND THE ROLE OF TAM THEREIN
Depending on the tumor type/model and the blocking agents used
to impede M-CSFR signaling (Figure 2) (variable) effects of MCSFR blockade on different aspects of cancer progression have
been reported (Figure 1B).
EFFECTS ON TUMOR INCIDENCE AND PRIMARY TUMOR GROWTH

To assess the role of M-CSF in tumor development and progression, csf1op /op mice have been crossed with transgenic mice
in which mammary tumors develop due to mammary epithelial restricted expression of the Polyomavirus middle T oncogene
(PyMT). In these experiments, the drastic reduction in TAM numbers in the absence of M-CSF was reported neither to affect the
incidence nor the growth of the primary tumors but rather to delay
their development to invasive, metastatic carcinomas (97). In fact,
the PyMT model is characterized by the development of a single
primary tumor focus on the ducts emanating from the nipple,
after which other tumors arise in the ducts distant to the nipple.
Although the development of multiple foci on the distal ducts was
reduced in the csf1op /op PyMT mammary glands, the growth rate
of the primary tumor size and the proliferation rate of the cancer
cells were comparable to those in M-CSF sufficient mice.
Similarly, treatment of AE5MG mesothelioma or LLC lung carcinoma bearing mice with the M-CSFR blocking mAb M279 was
described not to result in a significant effect on tumor growth
or final tumor burden, despite a strong reduction in the number of TAM (100). In contrast, publications reporting on the
use of another mAb, AFS98, for M-CSFR blockade and ensuing
TAM inhibition, documented inhibition of primary tumor growth
in different mouse tumor models including the implanted AX
osteosarcoma model (102) and later also in the EL4 transplanted
lymphoma model, the PyMT transgenic breast carcinoma model
and the MDA-MB231 breast cancer metastasis-induced osteolysis model (101). It has been suggested that the effect of the rat
IgG1 M279 may represent the biological response to blocking
CSF-1R signaling per se, whereas the isotype of the rat IgG2A
AFS98 may result in additional effector functions such as direct
macrophage depletion upon recognition by and/or aggregation

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TAM and M-CSF in cancer progression

with other macrophages through binding of the IgG2A antibody to
the high affinity IgG receptor CD64 on mouse macrophages (52).
In human, MCF-7 mammary carcinoma cell xenografts in
immunodeficient mice, M-CSF blockade by antisense oligonucleotides and small interfering RNAs has been shown to reduce
host macrophage infiltration and suppress tumor growth (118).
Concerning the effect of pharmacologic M-CSFR blockade on
primary tumor growth, the M-CSFR tyrosine kinase inhibitor
Ki20227 was described to reduce TAM content of tumors and
retard tumor cell proliferation in osteosarcoma (102) and similar results were reported for GW2580 in papillary thyroid cancer
(103). Yet, the reduction in intratumoral proliferation in GW2580treated papillary thyroid cancers was most evident within the stromal compartment (103). These results suggest that the observed
effects of M-CSF blockade may in that case at least partially reflect
inhibition of stromal cells such as TAM rather than cancer cell
proliferation per se. In murine prostate cancer models, the MCSFR inhibitors GW2580 or PLX3397 as a single treatment were
reported to have little effect on tumor growth compared with the
control group, despite effective TAM ablation.
A recent study clearly illustrates that the specificity of the
applied inhibitors for M-CSF as compared to other tyrosine
kinases and the relative contribution of the effect on TAM as
compared to direct effects on the cancer cells should be carefully considered when interpreting the effect of M-CSFR blockers on tumor growth. In this study, PLX3397 was found to
result in effective reduction of tumor weight and cellularity in
both the KitV558del/+ transgenic murine gastrointestinal stromal
tumor (GAST) model and in human GAST xenografts (111).
These GAST cells are known to be strongly dependent on signaling via the oncogene cKit for their survival and the growth
inhibitory effect of PLX3397 was even stronger than that of the
cKit tyrosine kinase inhibitor imatinib, correlating with a superior capacity of PLX3397 as compared to imatinib to decrease
the viability of two human GAST cell lines in vitro. On the
other hand, TAM were deleted to a much greater degree in mice
treated with PLX3397 than with imatinib, correlating with a more
potent M-CSFR inhibition by PLX3397 as compared to imatinib. Therefore, one could hypothesize that the superior effect
of PLX3397 on tumor growth inhibition could at least in part
be related to superior inhibition of M-CSF signaling and consequent TAM attenuation, acting synergistically to the Kit inhibition. To address this possibility, the authors combined imatinib
with PLX5622, an M-CSFR-specific inhibitor of equal potency
to PLX3397 that does not appreciably inhibit Kit. Despite comparable levels of TAM reduction as PLX3397 therapy, treatment
with PLX5622 did not enhance the effect of imatinib on tumor
weight, cell number, or histology, suggesting that inhibition of
cKit signaling but not M-CSFR signaling is the main factor determining the capacity of tyrosine kinase inhibitors for GAST growth
inhibition (111).
Overall, despite consistent reduction in TAM content in primary tumors in the various tumor models discussed above, the
effects of M-CSF or M-CSFR blockade and consequent TAM
attenuation on primary tumor growth seem to be quite variable,
depending on the tumor model and the blocking agents used, and
thus do not seem to correlate with TAM depletion per se.

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Laoui et al.

EFFECTS ON TUMOR ANGIOGENESIS

Crossing PyMT and csf1op /op mice revealed that a low density of
macrophages in the primary tumors correlated with a delay in the
angiogenic switch, identified as the formation of a high-density
vessel network. Genetic restoration of macrophage numbers in
the tumors of these mice by the transgenic expression of MCSF specifically in the mammary epithelium thereby rescued the
vessel phenotype (119). Similarly, crossing csf1op /op mice to the
RIP1-Tag2 (RT2) mouse model of pancreatic islet cancer was documented to decrease TAM by approximately 50% during all stages
of RT2 tumor progression and to generate a substantial reduction
in cumulative tumor burden, which resulted from a significant
decrease in angiogenic switching and the number of tumors, rather
than an evident effect on the growth of established tumors or on
the cancer cell proliferative capacity (98).
In a mammary tumor model based on xenografts of human
MCF-7 breast cancer cells in athymic nude mice, mouse (host)
M-CSF expression was found to be induced as the tumors progressed. In these mice, treatment with a murinized, polyethylene
glycol-linked recombinant Fab fragment of the MCSF1-033 neutralizing rabbit anti-mouse M-CSF antibody reduced the density
of both macrophages and proliferating endothelial cells, the latter
reflecting decreased levels of angiogenic activity in the mammary
tumor xenografts (99). In an immunocompetent mouse model of
osteosarcoma, in which mice were subcutaneously transplanted
with the mouse AX osteosarcoma cell line, the M-CSF inhibitor
Ki20227 or the AFS98 rat anti-murine M-CSFR mAb dramatically
decreased peritumoral and perivascular TAM, suppressed tumor
angiogenesis and lymphangiogenesis, disorganized extracellular
matrices and concomitantly dramatically suppressed metastasis
and improved prognosis (102). In contrast to VEGF blockade,
interruption of M-CSF signaling did not promote rapid vascular
regrowth. In addition, continuous M-CSF inhibition did not affect
healthy vascular and lymphatic systems outside tumors (102).
The notion that M-CSFR+ TIM, including both TAM and MOMDSC-like cells, contribute significantly to tumor angiogenesis,
was supported by Priceman et al. (115), showing that depletion of M-CSF-dependent TAM and MO-MDSC-like cells in the
3LL lung carcinoma model, using either the M-CSFR inhibitor
GW2580 or a transgenic approach in chimeric mice, resulted
in significant reduction in angiogenesis in TIM-ablated tumors
(without a concomitant decrease in tumor growth). The authors
confirmed that, also in the orthotopic RM-1 prostate tumor model,
M-CSF blockade resulted in reduced levels of TAM, and MOMDSC-like cells, associated with reduced angiogenesis and, to a
lesser extent, lymphangiogenesis, as reflected by vessel density in
these tumors.
In the 3LL lung carcinoma model, GW2580 was in addition shown to attenuate tumor evasion of anti-angiogenic therapy. In combination with DC101, a specific blocking antibody
against VEGFR-2, GW2580 resulted in greater inhibition of
tumor angiogenesis along with synergistic tumor growth reduction compared with anti-angiogenic therapy alone. In search for
a hypothesis on the mechanism underlying the reversal of antiangiogenesis in the combination therapy, the authors provided
histological data revealing more abundant MMP9 expressing cells
with heterogeneous myeloid cell morphology in viable areas of

Frontiers in Immunology | Molecular Innate Immunity

TAM and M-CSF in cancer progression

DC101-treated tumors, which were reduced in the combination
group (115).
At a mechanistic level, M-CSF was also shown to induce VEGF
production in human monocytes through the MAPK/ERK pathway via Sp1 and was reported to enhance angiogenesis in vivo, as
evidenced in an angiogenesis assay using an in vivo polymerized
MatrigelTM plug in mice (120). Recently, the mechanistic basis of
the tumor angiogenesis-promoting effect of M-CSF was further
expanded by showing that M-CSF augments differentiation of the
subpopulation of M2 macrophages expressing the endothelial cell
tyrosine kinase receptor, Tie2. Hereby, M-CSF-mediated upregulation of Tie2 on these Tie2-expressing monocytes/macrophages
(TEM) increased branching of human umbilical vein endothelial cells (HUVECs) in vitro and enhanced angiogenesis in
PyMT tumor-bearing mice. This M-CSF-stimulated Tie2 receptor expression was found to be dependent on a synergistic
contribution from the PI3 kinase and HIF-1α pathways. (121).
As a final remark, it should be realized that high levels of angiogenesis, driven by M2-like TAM, often lead to dysfunctional blood
vessels in tumors, resulting in more malignant cancer cells under
the influence of tumor hypoxia and an easy access of these cells to
the blood circulation. TAM depletion or the conversion of M2-like
TAM to M1-like TAM, thereby results in vessel normalization and
reduced metastasis (122). In addition, normalized vessels allow a
more efficient administration of therapeutic agents to the tumor
microenvironment.
EFFECTS ON CANCER CELL INVASION AND METASTASIS

Accumulating evidence in the first decade of this century has supported the tenet that delayed development of invasive, metastatic
carcinomas in PyMT csf1op /op mice is reflective of a role for M-CSF
in promoting cancer cell invasion by regulating the infiltration
and function of TAM. Indeed, at the PyMT tumor site, expression of M-CSFR was reported to be restricted to macrophages.
Moreover, restoration of macrophage infiltration upon transgenic
expression of M-CSF in the mammary epithelium restored progression of primary tumors to the stages of invasive carcinoma
(97). In fact, macrophages and tumor cells in mammary tumors
were documented to be comigratory and to be mutually dependent
for invasion and for cancer cell intravasation (90, 123). Hereby,
M-CSF produced by carcinoma cells promotes the expression of
EGF by macrophages, which in turn promotes the formation of
elongated protrusions and cell invasion by carcinoma cells. In
addition, EGF promotes the expression of M-CSF by carcinoma
cells, thereby generating a positive feedback loop. Disruption of
this paracrine amplification loop by blockade of either EGF receptor or M-CSFR signaling was found to be sufficient for inhibiting
both macrophage and tumor cell migration and invasion (91).
A similar EGF/M-CSF paracrine interaction with macrophages,
resulting in enhanced cancer cell invasion as reported for murine
carcinoma cells, was confirmed in a mouse xenograft model of
human breast tumor derived cancer cells. Yet, for these human
breast carcinoma cells, the EGF/M-CSF paracrine feedback loop
was found to be complemented by autocrine M-CSF signaling
in the cancer cells (124). These data correlated with the expression of M-CSFR by human but not mouse breast carcinoma cells.
The possibility of macrophage-independent effects of M-CSF on

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Laoui et al.

human cancer cell invasion is also supported by a direct stimulation of in vitro invasive capacity, but not proliferation, of human
adenocarcinoma cell lines by recombinant human M-CSF (125).
The studies in csf1op /op mice also indicated a role of M-CSF
in enhancement of metastatic growth of cancer cells. In particular, M-CSF was shown to be required for the recruitment
of a population of CD11b+ F4/80+ Gr1+ host macrophages to
extravasating pulmonary metastatic cells in the PyMT model.
This recruited CD11b+ F4/80+ Gr1+ macrophage population displayed a distinct phenotype as compared to CD11b− , CD11c+
lung resident macrophages and also did not express Tie2, rendering them distinct from the M-CSF-induced pro-angiogenic Tie2expressing monocytes/macrophages. The recruited macrophages
enhanced cancer cell metastasis through effects on cancer cell
metastatic seeding, extravasation, survival, and subsequent growth
(126). The authors confirmed that the reduced metastasis in
csf1op /op PyMT mice could be recapitulated in wild-type PyMT
mice via macrophage ablation using clodronate-containing liposomes. Importantly, even after metastatic growth had been established, macrophage ablation using clodronate-containing liposomes inhibited subsequent metastatic growth (126). This effect
also seems to be M-CSF-specific since transgenic expression of
M-CSF in the mammary epithelium of both csf1op/op and wildtype tumor-prone mice led to an acceleration to the late stages
of carcinoma and to a significant increase in pulmonary metastasis. The clinical significance of these findings is illustrated by the
observation that the density of close tripartite interactions between
cancer cells, macrophages, and endothelial cells (tumor microenvironment of metastasis or TMEM) is predictive of metastasis
formation in breast cancer patients (127).
Since M-CSF signaling not only plays a critical role in the
turnover of TAM, but is also crucial for osteoclasts, blocking
M-CSFR signaling may not only attenuate metastasis via effects
on TAM, but may have additional beneficial effects on metastatic disease via inhibitory effects on osteoclasts. As an example
of this, the AFS98 M-CSFR blocking mAb was recently documented to potently block the differentiation of osteoclasts and
their bone destruction activity in a breast cancer model of bone
metastasis (101).
A recent study placed a cautionary note on blocking M-CSFR
signaling as a therapeutic modality in cancer. In that study, mice
bearing two independently derived mammary cancer cell lines
(4T1.2 and EMT6.5) injected orthotopically into the mammary
gland, were treated with the AFS98 neutralizing anti-M-CSFR
mAb, with a neutralizing anti-mouse M-CSF mAb, or with two
different small molecule inhibitors of M-CSFR (GW2580 or
CYC11645). The authors observed variable effects on reduction
of TAM in the primary tumors or metastatic lung tissue, whereby
TAM could be reduced using GW2580 or high dose of AFS98,
but were not reduced when using lower dose of AFS98. Yet in
all these cases, not only did these various modalities for blocking M-CSFR not reduce primary tumor growth, but the intended
treatment actually increased metastasis to the lung and spine (66).
The authors found that the increased spontaneous metastasis upon
blocking of M-CSFR or M-CSF was associated with increased
levels of serum granulocyte-colony-stimulating factor (G-CSF),
increased numbers of neutrophils and Ly6Chigh monocytes in the

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TAM and M-CSF in cancer progression

peripheral blood and increased frequency of neutrophils in the
primary tumor and in the lung. It is currently unclear why MCSFR blockade resulted in increased G-CSF levels in this model,
but the authors did observe that blood neutrophil numbers were
proportional to the metastatic capacity of the different mammary
carcinomas evaluated, suggesting that certain carcinomas may be
more prone to mobilize neutrophils, and leading to increased
metastasis. Interestingly, combining blockade of M-CSFR signaling with a neutralizing antibody against the G-CSF receptor
(G-CSFR), which regulates neutrophil development and function,
reduced the enhanced metastasis, and neutrophil numbers that
resulted from M-CSFR blockade. In fact, the combined blocking of
M-CSFR and G-CSFR resulted in significantly reduced metastasis
as compared to the control condition (66).
POTENTIATION OF RADIO-, CHEMO-, AND IMMUNOTHERAPY

Whereas the M-CSFR inhibitors GW2580 or PLX3397 on their
own were reported to have little effect on tumor growth in murine
prostate cancer models, when added to radiotherapy, the M-CSFR
inhibitors suppressed tumor growth more effectively than radiation alone (116). The synergistic effect of M-CSF blockade on
the efficacy of radiotherapy was explained by the observation that
irradiation resulted in increased M-CSF levels due to recruitment
of the DNA damage-induced kinase ABL1 into cell nuclei where it
bound the csf1 gene promoter and enhanced csf1 gene transcription. Consequently, enhanced recruitment of TIM, including TAM
and MO-MDSC-like and PMN-MDSC-like cells, was detected and
this enhanced TIM recruitment was counteracted via the M-CSFR
inhibitors (116). These results suggest that blockade of the MCSF/M-CSFR axis can be a promising approach for developing
more effective combination cancer therapies. The authors supported the human relevance of these findings by reporting that also
in prostate cancer patients, serum levels of M-CSF were increased
after radiotherapy.
Such potential synergistic effects in combination therapy are
not only restricted to radiotherapy, but also extend to chemotherapy. Indeed, combination therapy with a murinized, polyethylene glycol-linked antigen-binding fragment against mouse (host)
M-CSF reportedly reversed chemoresistance in athymic nude,
immunodeficient mice bearing human, and chemoresistant MCF7 breast cancer xenografts (99). Also treatment with the AFS98
anti-M-CSFR monoclonal antibody in mice already bearing established PyMT tumors was reported to prolong their survival and
potentiate the effect of chemotherapy with Paclitaxel (101). Finally,
GW2580 or PLX3397 were found to improve chemotherapeutic
efficacy in mice bearing murine pancreatic ductal adenocarcinoma
cell lines. In this tumor model, gemcitabine chemotherapy was
documented to increase M-CSF levels and consequently enhance
the tumor infiltration of T-cell suppressive TAM (and MO-MDSClike cells). This effect was blunted when chemotherapy was combined with M-CSFR blockade, resulting in increased anti-tumor
CD8+ T-cell responses and improved inhibition of tumor growth
and metastasis as compared to chemotherapy as monotherapy.
Accordingly, the higher therapeutic efficacy of combined treatment with GW2580 plus gemcitabine compared with the effects
of gemcitabine alone was shown not to occur upon depletion of
CD8+ T lymphocytes (105).

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Considering the above, it comes as no surprise that, in the
same mouse model of pancreatic ductal adenocarcinoma, MCSFR signaling blockade using PLX3397 or GW2580 was shown
to enhance the therapeutic efficacy of so-called T-cell checkpoint
immunotherapy using PD1 and CTLA4 antagonists in combination with gemcitabine (114). In this case, M-CSFR signaling blockade was reported to result in preferential depletion of
MMRhigh M2-like TAM and reprograming of the phenoptype of
the remaining TAM, with alleviated immunosuppressive activities
and enhanced antigen presentation capacity and which in turn
correlated with enhanced CD4+ and CD8+ T cell responses. Here
also, the increased therapeutic efficacy of the combination treatment was shown to be blunted upon depletion of CD4+ and CD8+
T cells (114). As another example of a synergistic effect of MCSFR blockade on immunotherapy, PLX3397 has been reported
to improve the efficacy of adoptive cell therapy of melanomatargeted T cells in a syngeneic mouse model of BRAFV600E-driven
melanoma. Mice receiving the combined treatment produced
superior anti-tumor responses and exhibited improved overall
survival compared with single treatments, correlating with a dramatic reduction of TAM (but in this setting no significant change
in already low numbers of MO-MDSC-like or PMN-MDSC-like
cells), a skewing of the subpopulation balance in the remaining
TAM from predominant M2-oriented MHC-IIlow to predominant M1-oriented MHC-IIhigh macrophages and an increase in
tumor-infiltrating lymphocytes and T cells. The authors conclude that macrophages are the targets of PLX3397 by confirming
that PLX3397 and macrophage-depleting clodronate-containing
liposomes have the same effect on tumor growth and that this
effect is not further increasing when combining both depletion
methods (113).

TAM and M-CSF in cancer progression

And especially, synergistic effects of M-CSFR blocking agents
in diminishing TAM-dependent resistance to anti-angiogenic
therapy, radiotherapy, chemotherapy, or immunotherapy offer
promising perspectives for effective combination therapy. Recent
studies thereby suggest that intratumoral M-CSF levels and their
balance with GM-CSF levels are not only critical for TAM differentiation and maintenance, but can also contribute to shaping
the M1/M2 phenotypic and subpopulation heterogeneity of TAM.
Hence, M-CSFR blocking agents may not only have the potential to counteract cancer progression by reducing TAM content in
tumors and metastatic lesions, but also by re-educating TAM from
tumor-promoting toward anti-tumoral effector populations.
Recently, more attention is in addition being given to better characterize other tumor-infiltrating myeloid cell populations
such as MDSC-like cells that are also affected by M-CSFR blockade
and to evaluate whether these contribute to the observed effects
of M-CSFR blockade on various aspects of cancer progression.
Additional effects on other cells are not necessarily a disadvantage
in the context of anti-cancer therapeutic activity, as exemplified
in reported attenuation of metastatic disease via dual inhibitory
effects on TAM and osteoclasts (101). Yet, the data recently
reported by Swierczak and colleagues on neutrophil-dependent
enhanced metastasis upon M-CSFR blockade (66) indicate that
blocking M-CSFR signaling may have variable effects according to
the tumor model and may in some cases exhibit unwanted side
effects. These cautionary findings are testaments to the notion
that successful clinical translation will be critically dependent on
proper patient stratification to focus on those patient groups in
which high M-CSF or M-CSFR expression is linked to disease
pathophysiology and correlates with worse prognosis and in which
M-CSFR/macrophage-targeted therapies are thus most likely to
exert a beneficial effect.

CONCLUDING REMARKS AND CLINICAL PERSPECTIVES
Macrophage-colony-stimulating factor receptor inhibitors are
currently in clinical development as cancer therapeutics. Plexxicon
has, for example, initiated several clinical trials of the cKit and MCSFR inhibitor PLX3397, either as a stand-alone cancer treatment
(128–130) or in an adjuvant setting with chemo- and/or radiotherapy (131–134). Phase I clinical trials of anti-M-CSFR mAbs in
patients with advanced solid tumors are currently being conducted
by Eli Lilly and Company for the fully human IgG1 IMC-CS4
(107) and by Roche for the humanized IgG1 RG7155 (135). For
the latter, it was mentioned in a recent publication that, based on
preliminary results of an ongoing clinical trial, administration of
RG7155 to diffuse-type giant cell tumor patients led to significant
reductions of M-CSFR+ CD163+ macrophages in tumor tissues,
which correlated with at least partial clinical objective responses
(106). The ultimate value of these M-CSFR targeted therapies
will need to be assessed in follow-up studies aimed at demonstrating effects that go beyond reduction in the primary tumor
burden, but extend to attenuation of metastasis and prolongation
of patient survival.
In this context, it is encouraging that numerous studies in
preclinical tumor models have revealed that blocking M-CSFR
signaling, despite variable effects on primary tumor growth per se,
has the potential to attenuate tumor-promoting effects of TAM
on tumor angiogenesis and cancer cell invasion and metastasis.

Frontiers in Immunology | Molecular Innate Immunity

AUTHOR CONTRIBUTIONS
All authors have made substantial contributions to the conception
of the manuscript. Geert Raes and Damya Laoui have drafted the
initial version of the manuscript. All other authors have critically
reviewed the manuscript for important intellectual content. All
authors approve of the version of the manuscript and agree to be
accountable for all aspects of the work in ensuring that questions
related to the accuracy or integrity of any part of the work are
appropriately investigated and resolved.

ACKNOWLEDGMENTS
The authors thank FWO-Vlaanderen, IWT-Vlaanderen, the
“Stichting tegen Kanker,” and the “Vlaamse Liga tegen Kanker”
for their support.

REFERENCES
1. Hanahan D, Weinberg RA. The hallmarks of cancer. Cell (2000) 100(1):57–70.
doi:10.1016/S0092-8674(00)81683-9
2. Hanahan D, Weinberg RA. Hallmarks of cancer: the next generation. Cell
(2011) 144(5):646–74. doi:10.1016/j.cell.2011.02.013
3. Balkwill F, Charles KA, Mantovani A. Smoldering and polarized inflammation in the initiation and promotion of malignant disease. Cancer Cell (2005)
7(3):211–7. doi:10.1016/j.ccr.2005.02.013
4. Colotta F, Allavena P, Sica A, Garlanda C, Mantovani A. Cancer-related inflammation, the seventh hallmark of cancer: links to genetic instability. Carcinogenesis (2009) 30(7):1073–81. doi:10.1093/carcin/bgp127

October 2014 | Volume 5 | Article 489 | 112

Laoui et al.

5. Mantovani A, Allavena P, Sica A, Balkwill F. Cancer-related inflammation.
Nature (2008) 454(7203):436–44. doi:10.1038/nature07205
6. Van Ginderachter JA, Movahedi K, Van den Bossche J, De Baetselier P.
Macrophages, PPARs, and Cancer. PPAR Res (2008) 2008:169414. doi:10.1155/
2008/169414
7. Laoui D, Van Overmeire E, Movahedi K, Van den Bossche J, Schouppe E, Mommer C, et al. Mononuclear phagocyte heterogeneity in cancer: different subsets
and activation states reaching out at the tumor site. Immunobiology (2011)
216(11):1192–202. doi:10.1016/j.imbio.2011.06.007
8. Bingle L, Brown NJ, Lewis CE. The role of tumour-associated macrophages in
tumour progression: implications for new anticancer therapies. J Pathol (2002)
196(3):254–65. doi:10.1002/path.1027
9. Zhang QW, Liu L, Gong CY, Shi HS, Zeng YH, Wang XZ, et al. Prognostic significance of tumor-associated macrophages in solid tumor: a meta-analysis of the
literature. PLoS One (2012) 7(12):e50946. doi:10.1371/journal.pone.0050946
10. Lewis CE, Pollard JW. Distinct role of macrophages in different tumor microenvironments. Cancer Res (2006) 66(2):605–12. doi:10.1158/0008-5472.CAN05-4005
11. Jaiswal S, Jamieson CH, Pang WW, Park CY, Chao MP, Majeti R, et al. CD47
is upregulated on circulating hematopoietic stem cells and leukemia cells to
avoid phagocytosis. Cell (2009) 138(2):271–85. doi:10.1016/j.cell.2009.05.046
12. Chao MP, Jaiswal S, Weissman-Tsukamoto R, Alizadeh AA, Gentles AJ, Volkmer
J, et al. Calreticulin is the dominant pro-phagocytic signal on multiple human
cancers and is counterbalanced by CD47. Sci Transl Med (2010) 2(63):63ra94.
doi:10.1126/scitranslmed.3001375
13. Beatty GL, Chiorean EG, Fishman MP, Saboury B, Teitelbaum UR, Sun
W, et al. CD40 agonists alter tumor stroma and show efficacy against pancreatic carcinoma in mice and humans. Science (2011) 331(6024):1612–6.
doi:10.1126/science.1198443
14. Wu QL, Buhtoiarov IN, Sondel PM, Rakhmilevich AL, Ranheim EA. Tumoricidal effects of activated macrophages in a mouse model of chronic lymphocytic
leukemia. J Immunol (2009) 182(11):6771–8. doi:10.4049/jimmunol.0801847
15. Vicetti Miguel RD, Cherpes TL, Watson LJ, McKenna KC. CTL induction of
tumoricidal nitric oxide production by intratumoral macrophages is critical for tumor elimination. J Immunol (2010) 185(11):6706–18. doi:10.4049/
jimmunol.0903411
16. Guiducci C, Vicari AP, Sangaletti S, Trinchieri G, Colombo MP. Redirecting
in vivo elicited tumor infiltrating macrophages and dendritic cells towards
tumor rejection. Cancer Res (2005) 65(8):3437–46.
17. Corthay A, Skovseth DK, Lundin KU, Rosjo E, Omholt H, Hofgaard PO, et al.
Primary antitumor immune response mediated by CD4+ T cells. Immunity
(2005) 22(3):371–83. doi:10.1016/j.immuni.2005.02.003
18. Qian BZ, Pollard JW. Macrophage diversity enhances tumor progression and
metastasis. Cell (2010) 141(1):39–51. doi:10.1016/j.cell.2010.03.014
19. Ruffell B, Affara NI, Coussens LM. Differential macrophage programming in
the tumor microenvironment. Trends Immunol (2012) 33(3):119–26. doi:10.
1016/j.it.2011.12.001
20. Condeelis J, Pollard JW. Macrophages: obligate partners for tumor cell migration, invasion, and metastasis. Cell (2006) 124(2):263–6. doi:10.1016/j.cell.
2006.01.007
21. Murdoch C, Muthana M, Coffelt SB, Lewis CE. The role of myeloid cells in
the promotion of tumour angiogenesis. Nat Rev Cancer (2008) 8(8):618–31.
doi:10.1038/nrc2444
22. Pollard JW. Trophic macrophages in development and disease. Nat Rev
Immunol (2009) 9(4):259–70. doi:10.1038/nri2528
23. De Palma M, Lewis CE. Macrophage regulation of tumor responses to anticancer therapies. Cancer Cell (2013) 23(3):277–86. doi:10.1016/j.ccr.2013.02.
013
24. Egeblad M, Nakasone ES, Werb Z. Tumors as organs: complex tissues that interface with the entire organism. Dev Cell (2010) 18(6):884–901. doi:10.1016/j.
devcel.2010.05.012
25. Movahedi K, Laoui D, Gysemans C, Baeten M, Stange G, Van den Bossche J,
et al. Different tumor microenvironments contain functionally distinct subsets of macrophages derived from Ly6C(high) monocytes. Cancer Res (2010)
70(14):5728–39. doi:10.1158/0008-5472.CAN-09-4672
26. Movahedi K, Schoonooghe S, Laoui D, Houbracken I, Waelput W, Breckpot
K, et al. Nanobody-based targeting of the macrophage mannose receptor for
effective in vivo imaging of tumor-associated macrophages. Cancer Res (2012)
72(16):4165–77. doi:10.1158/0008-5472.CAN-11-2994

www.frontiersin.org

TAM and M-CSF in cancer progression

27. Laoui D, Van Overmeire E, Di Conza G, Aldeni C, Keirsse J, Morias Y,
et al. Tumor hypoxia does not drive differentiation of tumor-associated
macrophages but rather fine-tunes the M2-like macrophage population. Cancer Res (2014) 74(1):24–30. doi:10.1158/0008-5472.CAN-13-1196
28. Biswas SK, Mantovani A. Macrophage plasticity and interaction with lymphocyte subsets: cancer as a paradigm. Nat Immunol (2010) 11(10):889–96.
doi:10.1038/ni.1937
29. Wynn TA, Chawla A, Pollard JW. Macrophage biology in development, homeostasis and disease. Nature (2013) 496(7446):445–55. doi:10.1038/nature12034
30. Van Ginderachter JA, Movahedi K, Hassanzadeh Ghassabeh G, Meerschaut S,
Beschin A, Raes G, et al. Classical and alternative activation of mononuclear
phagocytes: picking the best of both worlds for tumor promotion. Immunobiology (2006) 211(6–8):487–501. doi:10.1016/j.imbio.2006.06.002
31. Gordon S. Alternative activation of macrophages. Nat Rev Immunol (2003)
3(1):23–35. doi:10.1038/nri978
32. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism
and functions. Immunity (2010) 32(5):593–604. doi:10.1016/j.immuni.2010.
05.007
33. Mills CD. M1 and M2 Macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
34. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
35. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine
system in diverse forms of macrophage activation and polarization. Trends
Immunol (2004) 25(12):677–86. doi:10.1016/j.it.2004.09.015
36. Martinez FO, Helming L, Gordon S. Alternative activation of macrophages: an
immunologic functional perspective. Annu Rev Immunol (2009) 27:451–83.
doi:10.1146/annurev.immunol.021908.132532
37. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122(3):787–95. doi:10.1172/JCI59643
38. Hassanzadeh Ghassabeh G, De Baetselier P, Brys L, Noel W, Van Ginderachter
JA, Meerschaut S, et al. Identification of a common gene signature for type II
cytokine-associated myeloid cells elicited in vivo in different pathologic conditions. Blood (2006) 108(2):575–83. doi:10.1182/blood-2005-04-1485
39. Koning N, van Eijk M, Pouwels W, Brouwer MS, Voehringer D, Huitinga I,
et al. Expression of the inhibitory CD200 receptor is associated with alternative macrophage activation. J Innate Immun (2010) 2(2):195–200. doi:10.1159/
000252803
40. Raes G, Beschin A, Ghassabeh GH, De Baetselier P. Alternatively activated
macrophages in protozoan infections. Curr Opin Immunol (2007) 19(4):454–9.
doi:10.1016/j.coi.2007.05.007
41. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
42. Gautier EL, Shay T, Miller J, Greter M, Jakubzick C, Ivanov S, et al. Geneexpression profiles and transcriptional regulatory pathways that underlie the
identity and diversity of mouse tissue macrophages. Nat Immunol (2012)
13(11):1118–28. doi:10.1038/ni.2419
43. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental
guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
44. Hamilton JA, Achuthan A. Colony stimulating factors and myeloid cell biology in health and disease. Trends Immunol (2013) 34(2):81–9. doi:10.1016/j.it.
2012.08.006
45. Lemmon MA, Schlessinger J. Cell signaling by receptor tyrosine kinases. Cell
(2010) 141(7):1117–34. doi:10.1016/j.cell.2010.06.011
46. Sapi E, Flick MB, Gilmore-Hebert M, Rodov S, Kacinski BM. Transcriptional
regulation of the c-fms (CSF-1R) proto-oncogene in human breast carcinoma
cells by glucocorticoids. Oncogene (1995) 10(3):529–42.
47. Sherr CJ. The role of the CSF-1 receptor gene (C-fms) in cell transformation.
Leukemia (1988) 2(12 Suppl):132S–42S.
48. Sasmono RT, Oceandy D, Pollard JW, Tong W, Pavli P, Wainwright BJ,
et al. A macrophage colony-stimulating factor receptor-green fluorescent
protein transgene is expressed throughout the mononuclear phagocyte system of the mouse. Blood (2003) 101(3):1155–63. doi:10.1182/blood-2002-020569
49. Mossadegh-Keller N, Sarrazin S, Kandalla PK, Espinosa L, Stanley ER, Nutt SL,
et al. M-CSF instructs myeloid lineage fate in single haematopoietic stem cells.
Nature (2013) 497(7448):239–43. doi:10.1038/nature12026

October 2014 | Volume 5 | Article 489 | 113

Laoui et al.

50. Davies LC, Jenkins SJ, Allen JE, Taylor PR. Tissue-resident macrophages. Nat
Immunol (2013) 14(10):986–95. doi:10.1038/ni.2705
51. Wiktor-Jedrzejczak W, Bartocci A, Ferrante AW Jr., Ahmed-Ansari A, Sell
KW, Pollard JW, et al. Total absence of colony-stimulating factor 1 in the
macrophage-deficient osteopetrotic (op/op) mouse. Proc Natl Acad Sci U S
A (1990) 87(12):4828–32. doi:10.1073/pnas.87.12.4828
52. Hume DA, MacDonald KP. Therapeutic applications of macrophage colonystimulating factor-1 (CSF-1) and antagonists of CSF-1 receptor (CSF-1R) signaling. Blood (2012) 119(8):1810–20. doi:10.1182/blood-2011-09-379214
53. Davies LC, Rosas M, Jenkins SJ, Liao CT, Scurr MJ, Brombacher F, et al.
Distinct bone marrow-derived and tissue-resident macrophage lineages proliferate at key stages during inflammation. Nat Commun (2013) 4:1886.
doi:10.1038/ncomms2877
54. Greter M, Lelios I, Pelczar P, Hoeffel G, Price J, Leboeuf M, et al. Stromaderived interleukin-34 controls the development and maintenance of Langerhans cells and the maintenance of microglia. Immunity (2012) 37(6):1050–60.
doi:10.1016/j.immuni.2012.11.001
55. Wang Y, Szretter KJ, Vermi W, Gilfillan S, Rossini C, Cella M, et al. IL-34 is a
tissue-restricted ligand of CSF1R required for the development of Langerhans
cells and microglia. Nat Immunol (2012) 13(8):753–60. doi:10.1038/ni.2360
56. Martinez FO, Gordon S, Locati M, Mantovani A. Transcriptional profiling of the
human monocyte-to-macrophage differentiation and polarization: new molecules and patterns of gene expression. J Immunol (2006) 177(10):7303–11.
doi:10.4049/jimmunol.177.10.7303
57. Fleetwood AJ, Lawrence T, Hamilton JA, Cook AD. Granulocyte-macrophage
colony-stimulating factor (CSF) and macrophage CSF-dependent macrophage
phenotypes display differences in cytokine profiles and transcription factor
activities: implications for CSF blockade in inflammation. J Immunol (2007)
178(8):5245–52. doi:10.4049/jimmunol.178.8.5245
58. Fleetwood AJ, Dinh H, Cook AD, Hertzog PJ, Hamilton JA. GM-CSF- and MCSF-dependent macrophage phenotypes display differential dependence on
type I interferon signaling. J Leukoc Biol (2009) 86(2):411–21. doi:10.1189/jlb.
1108702
59. Tagliani E, Shi C, Nancy P, Tay CS, Pamer EG, Erlebacher A. Coordinate regulation of tissue macrophage and dendritic cell population dynamics by CSF-1.
J Exp Med (2011) 208(9):1901–16. doi:10.1084/jem.20110866
60. Scholl SM, Pallud C, Beuvon F, Hacene K, Stanley ER, Rohrschneider L, et al.
Anti-colony-stimulating factor-1 antibody staining in primary breast adenocarcinomas correlates with marked inflammatory cell infiltrates and prognosis.
J Natl Cancer Inst (1994) 86(2):120–6. doi:10.1093/jnci/86.2.120
61. Kawamura K, Komohara Y, Takaishi K, Katabuchi H, Takeya M. Detection
of M2 macrophages and colony-stimulating factor 1 expression in serous
and mucinous ovarian epithelial tumors. Pathol Int (2009) 59(5):300–5.
doi:10.1111/j.1440-1827.2009.02369.x
62. Espinosa I, Catasus L, D’ Angelo E, Mozos A, Pedrola N, Bertolo C, et al. Stromal signatures in endometrioid endometrial carcinomas. Mod Pathol (2014)
27:631–9. doi:10.1038/modpathol.2013.131
63. Behnes CL, Bremmer F, Hemmerlein B, Strauss A, Strobel P, Radzun HJ. Tumorassociated macrophages are involved in tumor progression in papillary renal
cell carcinoma. Virchows Arch (2014) 464(2):191–6. doi:10.1007/s00428-0131523-0
64. Espinosa I, Beck AH, Lee CH, Zhu S, Montgomery KD, Marinelli RJ, et al.
Coordinate expression of colony-stimulating factor-1 and colony-stimulating
factor-1-related proteins is associated with poor prognosis in gynecological
and nongynecological leiomyosarcoma. Am J Pathol (2009) 174(6):2347–56.
doi:10.2353/ajpath.2009.081037
65. Kluger HM, Dolled-Filhart M, Rodov S, Kacinski BM, Camp RL, Rimm DL.
Macrophage colony-stimulating factor-1 receptor expression is associated with
poor outcome in breast cancer by large cohort tissue microarray analysis. Clin
Cancer Res (2004) 10(1 Pt 1):173–7. doi:10.1158/1078-0432.CCR-0699-3
66. Swierczak A, Cook AD, Lenzo JC, Restall CM, Doherty JP, Anderson RL,
et al. The promotion of breast cancer metastasis caused by inhibition of CSF1R/CSF-1 signaling is blocked by targeting the G-CSF receptor. Cancer Immunol
Res (2014) 2:765–76. doi:10.1158/2326-6066.CIR-13-0190
67. Koh YW, Park C, Yoon DH, Suh C, Huh J. CSF-1R expression in tumorassociated macrophages is associated with worse prognosis in classical
Hodgkin lymphoma. Am J Clin Pathol (2014) 141(4):573–83. doi:10.1309/
AJCPR92TDDFARISU

Frontiers in Immunology | Molecular Innate Immunity

TAM and M-CSF in cancer progression

68. Beck AH, Espinosa I, Edris B, Li R, Montgomery K, Zhu S, et al. The macrophage
colony-stimulating factor 1 response signature in breast carcinoma. Clin Cancer Res (2009) 15(3):778–87. doi:10.1158/1078-0432.CCR-08-1283
69. Sharma M, Beck AH, Webster JA, Espinosa I, Montgomery K, Varma S,
et al. Analysis of stromal signatures in the tumor microenvironment of
ductal carcinoma in situ. Breast Cancer Res Treat (2010) 123(2):397–404.
doi:10.1007/s10549-009-0654-0
70. McDermott RS, Deneux L, Mosseri V, Vedrenne J, Clough K, Fourquet A, et al.
Circulating macrophage colony stimulating factor as a marker of tumour progression. Eur Cytokine Netw (2002) 13(1):121–7.
71. Aharinejad S, Salama M, Paulus P, Zins K, Berger A, Singer CF. Elevated CSF1
serum concentration predicts poor overall survival in women with early breast
cancer. Endocr Relat Cancer (2013) 20(6):777–83. doi:10.1530/ERC-13-0198
72. Tamimi RM, Brugge JS, Freedman ML, Miron A, Iglehart JD, Colditz GA,
et al. Circulating colony stimulating factor-1 and breast cancer risk. Cancer Res
(2008) 68(1):18–21. doi:10.1158/0008-5472.CAN-07-3234
73. Biswas SK, Gangi L, Paul S, Schioppa T, Saccani A, Sironi M, et al. A distinct and
unique transcriptional program expressed by tumor-associated macrophages
(defective NF-kappaB and enhanced IRF-3/STAT1 activation). Blood (2006)
107(5):2112–22. doi:10.1182/blood-2005-01-0428
74. Sica A, Bronte V. Altered macrophage differentiation and immune dysfunction in tumor development. J Clin Invest (2007) 117(5):1155–66. doi:10.1172/
JCI31422
75. Mantovani A, Sozzani S, Locati M, Allavena P, Sica A. Macrophage polarization: tumor-associated macrophages as a paradigm for polarized M2 mononuclear phagocytes. Trends Immunol (2002) 23(11):549–55. doi:10.1016/S14714906(02)02302-5
76. Hagemann T, Wilson J, Burke F, Kulbe H, Li NF, Pluddemann A, et al. Ovarian cancer cells polarize macrophages toward a tumor-associated phenotype.
J Immunol (2006) 176(8):5023–32. doi:10.4049/jimmunol.176.8.5023
77. Torroella-Kouri M, Silvera R, Rodriguez D, Caso R, Shatry A, Opiela S, et al.
Identification of a subpopulation of macrophages in mammary tumor-bearing
mice that are neither M1 nor M2 and are less differentiated. Cancer Res (2009)
69(11):4800–9. doi:10.1158/0008-5472.CAN-08-3427
78. Biswas SK, Allavena P, Mantovani A. Tumor-associated macrophages: functional diversity, clinical significance, and open questions. Semin Immunopathol
(2013) 35(5):585–600. doi:10.1007/s00281-013-0367-7
79. Umemura N, Saio M, Suwa T, Kitoh Y, Bai J, Nonaka K, et al. Tumorinfiltrating myeloid-derived suppressor cells are pleiotropic-inflamed monocytes/macrophages that bear M1- and M2-type characteristics. J Leukoc Biol
(2008) 83(5):1136–44. doi:10.1189/jlb.0907611
80. Mills CD, Shearer J, Evans R, Caldwell MD. Macrophage arginine metabolism
and the inhibition or stimulation of cancer. J Immunol (1992) 149(8):2709–14.
81. Zaynagetdinov R, Sherrill TP, Polosukhin VV, Han W, Ausborn JA, McLoed AG,
et al. A critical role for macrophages in promotion of urethane-induced lung
carcinogenesis. J Immunol (2011) 187(11):5703–11. doi:10.4049/jimmunol.
1100558
82. Zaki MA, Wada N, Ikeda J, Shibayama H, Hashimoto K, Yamagami T, et al.
Prognostic implication of types of tumor-associated macrophages in Hodgkin
lymphoma. Virchows Arch (2011) 459(4):361–6. doi:10.1007/s00428-0111140-8
83. Wang B, Li Q, Qin L, Zhao S, Wang J, Chen X. Transition of tumorassociated macrophages from MHC class II(hi) to MHC class II(low) mediates
tumor progression in mice. BMC Immunol (2011) 12:43. doi:10.1186/14712172-12-43
84. Zhang M, He Y, Sun X, Li Q, Wang W, Zhao A, et al. A high M1/M2 ratio of
tumor-associated macrophages is associated with extended survival in ovarian
cancer patients. J Ovarian Res (2014) 7(1):19. doi:10.1186/1757-2215-7-19
85. Hagemann T, Lawrence T, McNeish I, Charles KA, Kulbe H, Thompson RG,
et al. “Re-educating” tumor-associated macrophages by targeting NF-kappaB.
J Exp Med (2008) 205(6):1261–8. doi:10.1084/jem.20080108
86. Van Overmeire E, Laoui D, Keirsse J, Van Ginderachter JA, Sarukhan A. Mechanisms driving macrophage diversity and specialization in distinct tumor
microenvironments and parallelisms with other tissues. Front Immunol (2014)
5:127. doi:10.3389/fimmu.2014.00127
87. Van Overmeire E, Laoui D, Keirsse J, Van Ginderachter JA. Hypoxia and tumorassociated macrophages: a deadly alliance in support of tumor progression.
Oncoimmunology (2014) 3(1):e27561. doi:10.4161/onci.27561

October 2014 | Volume 5 | Article 489 | 114

Laoui et al.

88. Laoui D, Movahedi K, Van Overmeire E, Van den Bossche J, Schouppe E, Mommer C, et al. Tumor-associated macrophages in breast cancer: distinct subsets, distinct functions. Int J Dev Biol (2011) 55(7–9):861–7. doi:10.1387/ijdb.
113371dl
89. Schouppe E, De Baetselier P, Van Ginderachter JA, Sarukhan A. Instruction of
myeloid cells by the tumor microenvironment: open questions on the dynamics
and plasticity of different tumor-associated myeloid cell populations. Oncoimmunology (2012) 1(7):1135–45. doi:10.4161/onci.21566
90. Wyckoff JB, Wang Y, Lin EY, Li JF, Goswami S, Stanley ER, et al. Direct
visualization of macrophage-assisted tumor cell intravasation in mammary tumors. Cancer Res (2007) 67(6):2649–56. doi:10.1158/0008-5472.CAN06-1823
91. Goswami S, Sahai E, Wyckoff JB, Cammer M, Cox D, Pixley FJ, et al.
Macrophages promote the invasion of breast carcinoma cells via a colonystimulating factor-1/epidermal growth factor paracrine loop. Cancer Res
(2005) 65(12):5278–83. doi:10.1158/0008-5472.CAN-04-1853
92. Kedrin D, Gligorijevic B, Wyckoff J, Verkhusha VV, Condeelis J, Segall JE, et al.
Intravital imaging of metastatic behavior through a mammary imaging window. Nat Methods (2008) 5(12):1019–21. doi:10.1038/nmeth.1269
93. Egeblad M, Ewald AJ, Askautrud HA, Truitt ML, Welm BE, Bainbridge E, et al.
Visualizing stromal cell dynamics in different tumor microenvironments by
spinning disk confocal microscopy. Dis Model Mech (2008) 1(2–3):155–67.
doi:10.1242/dmm.000596
94. Ojalvo LS, Whittaker CA, Condeelis JS, Pollard JW. Gene expression analysis of
macrophages that facilitate tumor invasion supports a role for Wnt-signaling
in mediating their activity in primary mammary tumors. J Immunol (2010)
184(2):702–12. doi:10.4049/jimmunol.0902360
95. Tymoszuk P, Evens H, Marzola V, Wachowicz K, Wasmer MH, Datta S,
et al. In situ proliferation contributes to accumulation of tumor-associated
macrophages in spontaneous mammary tumors. Eur J Immunol (2014)
44:2247–62. doi:10.1002/eji.201344304
96. Casazza A, Laoui D, Wenes M, Rizzolio S, Bassani N, Mambretti M, et al. Impeding macrophage entry into hypoxic tumor areas by Sema3A/Nrp1 signaling
blockade inhibits angiogenesis and restores antitumor immunity. Cancer Cell
(2013) 24(6):695–709. doi:10.1016/j.ccr.2013.11.007
97. Lin EY, Nguyen AV, Russell RG, Pollard JW. Colony-stimulating factor 1 promotes progression of mammary tumors to malignancy. J Exp Med (2001)
193(6):727–40. doi:10.1084/jem.193.6.727
98. Pyonteck SM, Gadea BB, Wang HW, Gocheva V, Hunter KE, Tang LH,
et al. Deficiency of the macrophage growth factor CSF-1 disrupts pancreatic neuroendocrine tumor development. Oncogene (2012) 31(11):1459–67.
doi:10.1038/onc.2011.337
99. Paulus P, Stanley ER, Schafer R, Abraham D, Aharinejad S. Colony-stimulating
factor-1 antibody reverses chemoresistance in human MCF-7 breast cancer
xenografts. Cancer Res (2006) 66(8):4349–56. doi:10.1158/0008-5472.CAN05-3523
100. MacDonald KP, Palmer JS, Cronau S, Seppanen E, Olver S, Raffelt NC, et al. An
antibody against the colony-stimulating factor 1 receptor depletes the resident
subset of monocytes and tissue- and tumor-associated macrophages but does
not inhibit inflammation. Blood (2010) 116(19):3955–63. doi:10.1182/blood2010-02-266296
101. Fend L, Accart N, Kintz J, Cochin S, Reymann C, Le Pogam F, et al. Therapeutic
effects of anti-CD115 monoclonal antibody in mouse cancer models through
dual inhibition of tumor-associated macrophages and osteoclasts. PLoS One
(2013) 8(9):e73310. doi:10.1371/journal.pone.0073310
102. Kubota Y, Takubo K, Shimizu T, Ohno H, Kishi K, Shibuya M, et al. M-CSF
inhibition selectively targets pathological angiogenesis and lymphangiogenesis.
J Exp Med (2009) 206(5):1089–102. doi:10.1084/jem.20081605
103. Ryder M, Gild M, Hohl TM, Pamer E, Knauf J, Ghossein R, et al. Genetic
and pharmacological targeting of CSF-1/CSF-1R inhibits tumor-associated
macrophages and impairs BRAF-induced thyroid cancer progression. PLoS
One (2013) 8(1):e54302. doi:10.1371/journal.pone.0054302
104. Strachan DC, Ruffell B, Oei Y, Bissell MJ, Coussens LM, Pryer N, et al.
CSF1R inhibition delays cervical and mammary tumor growth in murine
models by attenuating the turnover of tumor-associated macrophages and
enhancing infiltration by CD8 T cells. Oncoimmunology (2013) 2(12):e26968.
doi:10.4161/onci.26968
105. Mitchem JB, Brennan DJ, Knolhoff BL, Belt BA, Zhu Y, Sanford DE, et al. Targeting tumor-infiltrating macrophages decreases tumor-initiating cells, relieves

www.frontiersin.org

TAM and M-CSF in cancer progression

106.

107.

108.

109.

110.

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

122.

immunosuppression, and improves chemotherapeutic responses. Cancer Res
(2013) 73(3):1128–41. doi:10.1158/0008-5472.CAN-12-2731
Ries CH, Cannarile MA, Hoves S, Benz J, Wartha K, Runza V, et al. Targeting tumor-associated macrophages with anti-CSF-1R antibody reveals a strategy for cancer therapy. Cancer Cell (2014) 25(6):846–59. doi:10.1016/j.ccr.
2014.05.016
Phase 1 study of IMC-CS4, a monoclonal antibody targeted to the CSF1 receptor (CSF-1R), in subjects with advanced solid tumors refractory to
standard therapy or for which no standard therapy is available [Internet]. Eli Lilly and Company. (2011) [Cited 2014 July 11]. Available from:
http://clinicaltrials.gov/show/NCT01346358.
Pyonteck SM, Akkari L, Schuhmacher AJ, Bowman RL, Sevenich L, Quail DF,
et al. CSF-1R inhibition alters macrophage polarization and blocks glioma progression. Nat Med (2013) 19(10):1264–72. doi:10.1038/nm.3337
Ohno H, Kubo K, Murooka H, Kobayashi Y, Nishitoba T, Shibuya M, et al. A
c-fms tyrosine kinase inhibitor, Ki20227, suppresses osteoclast differentiation
and osteolytic bone destruction in a bone metastasis model. Mol Cancer Ther
(2006) 5(11):2634–43. doi:10.1158/1535-7163.MCT-05-0313
Conway JG, McDonald B, Parham J, Keith B, Rusnak DW, Shaw E, et al.
Inhibition of colony-stimulating-factor-1 signaling in vivo with the orally
bioavailable cFMS kinase inhibitor GW2580. Proc Natl Acad Sci U S A (2005)
102(44):16078–83. doi:10.1073/pnas.0502000102
Kim TS, Cavnar MJ, Cohen NA, Sorenson EC, Greer JB, Seifert AM, et al.
Increased KIT inhibition enhances therapeutic efficacy in gastrointestinal stromal tumor. Clin Cancer Res (2014) 20(9):2350–62. doi:10.1158/1078-0432.
CCR-13-3033
Kitoh Y, Saio M, Gotoh N, Umemura N, Nonaka K, Bai J, et al. Combined
GM-CSF treatment and M-CSF inhibition of tumor-associated macrophages
induces dendritic cell-like signaling in vitro. Int J Oncol (2011) 38(5):1409–19.
doi:10.3892/ijo.2011.960
Mok S, Koya RC, Tsui C, Xu J, Robert L, Wu L, et al. Inhibition of CSF-1 receptor improves the antitumor efficacy of adoptive cell transfer immunotherapy.
Cancer Res (2014) 74(1):153–61. doi:10.1158/0008-5472.CAN-13-1816
Zhu Y, Knolhoff BL, Meyer MA, Nywening TM, West BL, Luo J, et al.
CSF1/CSF1R blockade reprograms tumor-infiltrating macrophages and
improves response to T Cell checkpoint immunotherapy in pancreatic cancer
models. Cancer Res (2014) 74:5057–69. doi:10.1158/0008-5472.CAN-13-3723
Priceman SJ, Sung JL, Shaposhnik Z, Burton JB, Torres-Collado AX, Moughon
DL, et al. Targeting distinct tumor-infiltrating myeloid cells by inhibiting CSF1 receptor: combating tumor evasion of antiangiogenic therapy. Blood (2010)
115(7):1461–71. doi:10.1182/blood-2009-08-237412
Xu J, Escamilla J, Mok S, David J, Priceman S, West B, et al. CSF1R signaling blockade stanches tumor-infiltrating myeloid cells and improves the efficacy of radiotherapy in prostate cancer. Cancer Res (2013) 73(9):2782–94.
doi:10.1158/0008-5472.CAN-12-3981
Corzo CA, Condamine T, Lu L, Cotter MJ, Youn JI, Cheng P, et al. HIF1alpha regulates function and differentiation of myeloid-derived suppressor
cells in the tumor microenvironment. J Exp Med (2010) 207(11):2439–53.
doi:10.1084/jem.20100587
Aharinejad S, Paulus P, Sioud M, Hofmann M, Zins K, Schafer R, et al. Colonystimulating factor-1 blockade by antisense oligonucleotides and small interfering RNAs suppresses growth of human mammary tumor xenografts in mice.
Cancer Res (2004) 64(15):5378–84. doi:10.1158/0008-5472.CAN-04-0961
Lin EY, Li JF, Gnatovskiy L, Deng Y, Zhu L, Grzesik DA, et al. Macrophages
regulate the angiogenic switch in a mouse model of breast cancer. Cancer Res
(2006) 66(23):11238–46. doi:10.1158/0008-5472.CAN-06-1278
Curry JM, Eubank TD, Roberts RD, Wang Y, Pore N, Maity A, et al. MCSF signals through the MAPK/ERK pathway via Sp1 to induce VEGF production and induces angiogenesis in vivo. PLoS One (2008) 3(10):e3405.
doi:10.1371/journal.pone.0003405
Forget MA, Voorhees JL, Cole SL, Dakhlallah D, Patterson IL, Gross AC,
et al. Macrophage colony-stimulating factor augments tie2-expressing monocyte differentiation, angiogenic function, and recruitment in a mouse model
of breast cancer. PLoS One (2014) 9(6):e98623. doi:10.1371/journal.pone.
0098623
Rolny C, Mazzone M, Tugues S, Laoui D, Johansson I, Coulon C, et al. HRG
inhibits tumor growth and metastasis by inducing macrophage polarization
and vessel normalization through downregulation of PlGF. Cancer Cell (2011)
19(1):31–44. doi:10.1016/j.ccr.2010.11.009

October 2014 | Volume 5 | Article 489 | 115

Laoui et al.

123. Wyckoff J, Wang W, Lin EY, Wang Y, Pixley F, Stanley ER, et al. A paracrine loop
between tumor cells and macrophages is required for tumor cell migration in
mammary tumors. Cancer Res (2004) 64(19):7022–9. doi:10.1158/0008-5472.
CAN-04-1449
124. Patsialou A, Wyckoff J, Wang Y, Goswami S, Stanley ER, Condeelis JS. Invasion
of human breast cancer cells in vivo requires both paracrine and autocrine
loops involving the colony-stimulating factor-1 receptor. Cancer Res (2009)
69(24):9498–506. doi:10.1158/0008-5472.CAN-09-1868
125. Uemura Y, Kobayashi M, Nakata H, Kubota T, Bandobashi K, Saito T, et al.
Effects of GM-CSF and M-CSF on tumor progression of lung cancer: roles of
MEK1/ERK and AKT/PKB pathways. Int J Mol Med (2006) 18(2):365–73.
126. Qian B, Deng Y, Im JH, Muschel RJ, Zou Y, Li J, et al. A distinct macrophage
population mediates metastatic breast cancer cell extravasation, establishment and growth. PLoS One (2009) 4(8):e6562. doi:10.1371/journal.pone.
0006562
127. Robinson BD, Sica GL, Liu YF, Rohan TE, Gertler FB, Condeelis JS, et al. Tumor
microenvironment of metastasis in human breast carcinoma: a potential prognostic marker linked to hematogenous dissemination. Clin Cancer Res (2009)
15(7):2433–41. doi:10.1158/1078-0432.CCR-08-2179
128. A Phase 1 study to assess safety, pharmacokinetics, and pharmacodynamics of
PLX3397 in patients with advanced, incurable, solid tumors in which the target kinases are linked to disease pathophysiology [Internet]. Plexxikon. (2009).
Available from: http://clinicaltrials.gov/show/NCT01004861.
129. A phase 1/2 safety and efficacy study of orally administered PLX3397
in adults with relapsed or refractory acute myeloid Leukemia (AML)
[Internet]. Plexxikon. (2011). Available from: http://clinicaltrials.gov/show/
NCT01349049.
130. A phase 2 study of orally administered PLX3397 in patients with recurrent
glioblastoma [Internet]. Plexxikon. (2011). Available from: http://clinicaltrials.
gov/show/NCT01349036.
131. A phase 1b study to assess the safety of PLX3397 and paclitaxel in patients
with advanced solid tumors [Internet]. Plexxikon. (2012). Available from:
http://clinicaltrials.gov/show/NCT01525602.
132. Enhancing efficacy of chemotherapy in triple negative/basal-like breast cancer
by targeting macrophages: a multicenter phase Ib/II study of PLX 3397 and

Frontiers in Immunology | Molecular Innate Immunity

TAM and M-CSF in cancer progression

eribulin in patients with metastatic breast cancer [Internet]. Plexxikon. (2012).
Available from: http://clinicaltrials.gov/show/NCT01596751.
133. An open label phase 1b/2 study of orally administered PLX3397 in combination with radiation therapy and temozolomide in patients with newly
diagnosed glioblastoma [Internet]. Plexxikon. (2013). Available from: http:
//clinicaltrials.gov/show/NCT01790503.
134. A phase 1b open label, dose escalation study to assess safety, pharmacokinetics,
pharmacodynamics, and antitumor activity of PLX3397 in combination with
vemurafenib in V600-mutated BRAF unresectable or metastatic melanoma
[Internet]. Plexxikon. (2013). Available from: http://clinicaltrials.gov/show/
NCT01826448.
135. Open-label, multicenter, dose escalation phase Ia/Ib study with expansion
phase to evaluate safety, pharmacokinetics and activity of RO5509554, administered as an intravenous infusion as monotherapy and in combination with
paclitaxel in patients with advanced solid tumors [Internet]. Hoffmann-La
Roche. (2011). Available from: http://clinicaltrials.gov/show/NCT01494688.
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 13 August 2014; accepted: 22 September 2014; published online: 07 October
2014.
Citation: Laoui D, Van Overmeire E, De Baetselier P, Van Ginderachter JA and
Raes G (2014) Functional relationship between tumor-associated macrophages and
macrophage colony-stimulating factor as contributors to cancer progression. Front.
Immunol. 5:489. doi: 10.3389/fimmu.2014.00489
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Laoui, Van Overmeire, De Baetselier, Van Ginderachter and Raes.
This is an open-access article distributed under the terms of the Creative Commons
Attribution License (CC BY). The use, distribution or reproduction in other forums is
permitted, provided the original author(s) or licensor are credited and that the original
publication in this journal is cited, in accordance with accepted academic practice. No
use, distribution or reproduction is permitted which does not comply with these terms.

October 2014 | Volume 5 | Article 489 | 116

ORIGINAL RESEARCH ARTICLE
published: 25 November 2014
doi: 10.3389/fimmu.2014.00587

Depletion of tumor-associated macrophages slows the
growth of chemically induced mouse lung
adenocarcinomas
Jason M. Fritz 1 , Meredith A. Tennis 2 , David J. Orlicky 3 , Hao Yin 1 , Cynthia Ju 1 , Elizabeth F. Redente 4 ,
Kevin S. Choo 5 , Taylor A. Staab 5 , Ronald J. Bouchard 5 , Daniel T. Merrick 3 , Alvin M. Malkinson 1 and
Lori D. Dwyer-Nield 1 *
1
2
3
4
5

Department of Pharmaceutical Sciences, Skaggs School of Pharmacy and Pharmaceutical Sciences, University of Colorado Denver, Aurora, CO, USA
Pulmonary Division, School of Medicine, University of Colorado Denver, Aurora, CO, USA
Department of Pathology, School of Medicine, University of Colorado Denver, Aurora, CO, USA
Department of Pediatrics, National Jewish Health, Denver, CO, USA
Research Division, Eastern Colorado Veterans Administration Medical Center, Denver, CO, USA

Edited by:
Laurel L. Lenz, University of Colorado
School of Medicine, USA
Reviewed by:
Janos G. Filep, University of
Montreal, Canada
Giovanna Schiavoni, Istituto Superiore
di Sanita’, Italy
Charles Dudley Mills, BioMedical
Consultants, USA
*Correspondence:
Lori D. Dwyer-Nield , Department of
Pharmaceutical Sciences, School of
Pharmacy, University of Colorado
Denver, Anschutz Medical Campus,
C238-V20, Room 4460, 12850 E.
Montview Blvd, Aurora, CO 80045,
USA
e-mail: lori.nield@ucdenver.edu

Chronic inflammation is a risk factor for lung cancer, and low-dose aspirin intake reduces
lung cancer risk. However, the roles that specific inflammatory cells and their products play
in lung carcinogenesis have yet to be fully elucidated. In mice, alveolar macrophage numbers increase as lung tumors progress, and pulmonary macrophage programing changes
within 2 weeks of carcinogen exposure. To examine how macrophages specifically affect
lung tumor progression, they were depleted in mice bearing urethane-induced lung tumors
using clodronate-encapsulated liposomes. Alveolar macrophage populations decreased to
≤50% of control levels after 4–6 weeks of liposomal clodronate treatment. Tumor burden
decreased by 50% compared to vehicle treated mice, and tumor cell proliferation, as measured by Ki67 staining, was also attenuated. Pulmonary fluid levels of insulin-like growth
factor-I, CXCL1, IL-6, and CCL2 diminished with clodronate liposome treatment. Tumorassociated macrophages expressed markers of both M1 and M2 programing in vehicle
and clodronate liposome-treated mice. Mice lacking CCR2 (the receptor for macrophage
chemotactic factor CCL2) had comparable numbers of alveolar macrophages and showed
no difference in tumor growth rates when compared to similarly treated wild-type mice
suggesting that while CCL2 may recruit macrophages to lung tumor microenvironments,
redundant pathways can compensate when CCL2/CCR2 signaling is inactivated. Depletion of pulmonary macrophages rather than inhibition of their recruitment may be an
advantageous strategy for attenuating lung cancer progression.
Keywords: macrophage, programing, lung tumor, clodronate, inflammation

INTRODUCTION
Lung cancer is responsible for 29% of all cancer deaths in North
America, making it more lethal than breast, colon, prostate, and
pancreatic cancer combined (1). Approximately 85% of lung cancer cases are smoking-related (2, 3), and tobacco smoke contains
both direct carcinogens and agents that promote the growth of
nascent tumors. Non-small cell lung cancer (NSCLC) constitutes
>75% of lung cancer cases, with adenocarcinoma (AC) being the
most frequently diagnosed subtype, regardless of smoking status (4, 5). Lung cancer has long been associated with chronic
inflammatory disease. Limiting chronic inflammation may halt
the rapid growth and progression of this disease (6, 7) since longterm, low-dose aspirin use reduces the risk of death from lung AC
by 45%. However, patients with non-AC subtypes of lung cancer
were not protected by aspirin use, suggesting that chronic inflammation may be uniquely important for AC progression (7). In
addition, increased numbers of pulmonary macrophages correlate with poor prognosis in NSCLC patients (8–10), and alveolar

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macrophage numbers also increase during lung tumor progression
in mouse models of AC (11, 12). Macrophage depletion early in
tumor formation decreases tumor multiplicity (12, 13) indicating
a role for inflammatory cells in tumor development even before
increased macrophage numbers are detected. Macrophages have
been described as obligate partners for breast cancer metastasis to
the lung in animal models, and activation of PPARγ in pulmonary
macrophages promotes lung cancer progression and metastasis
in a murine orthotopic model (14). Prolonged lung inflammation increases tumor multiplicity by promoting clonal expansion
of previously initiated cells (15), and chronic anti-inflammatory
drug therapy during chemical promotion decreases tumor multiplicity (13, 16). Chronic inflammation drives lung tumor growth
and progression in mouse models and human disease, and alveolar
macrophages facilitate much of this effect.
Alveolar macrophages produce numerous epithelial growth
factors in response to tissue damage, including insulin-like growth
factor-I (IGF-I) (8, 17). IGF-I receptor (IGF-IR) is required

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for anchorage-independent growth of epithelial cells and has
been studied in neoplastic proliferation for over 20 years. IGF-IR
inhibitors are an area of interest for lung cancer therapy (17, 18).
Macrophage IGF-I production is highly induced in response to
environmental insult (19). While resident alveolar macrophages
are a likely physiological source of lung IGF-I, this growth factor is undetectable in undifferentiated human peripheral blood
monocytes (20–22). Alternative macrophage programing occurs
early in lung tumorigenesis, corresponding with elevated IGF-I
production (23–26). Consistent with this association of tumor
growth and enhanced macrophage IGF-I production, transgenic
mice that produce twice as much IGF-I in bronchoalveolar lavage
fluid (BALF) compared to wild-type controls develop spontaneous
lung hyperplasias and adenomas after 12 months (27). Despite the
evidence linking lung inflammation, macrophage function, IGFI production, and tumor progression, the relationship between
macrophage-derived IGF-I and lung tumor cell proliferation
in vivo has not been fully explored. We previously showed that
BALF from lung tumor-bearing mice contains 3.5-times more
IGF-I than that from naïve mice, and macrophage-produced IGFI enhances neoplastic proliferation in vitro (26), indicating that
macrophage IGF-I production may play a major role in early lung
tumor progression.
Macrophages are selective targets for liposomal clodronateinduced apoptosis because they aggressively phagocytize liposomes (28). Their increased expression of phospholipases facilitates rapid release of clodronate from the liposome vehicle into
the phagocyte upon liposome engulfment (29–31). When administered intratracheally (IT), liposomes do not enter the systemic
circulation and deplete only alveolar macrophages (29, 31). Conversely, liposomes given intravenously (IV) systemically deplete
myeloid cells in the bone marrow, liver, spleen, and other tissues,
and reduce the number of circulating cells available for recruitment to the lungs (32). Herein, we use a combination of IT and IV
administration of clodronate liposomes to deplete macrophages
from the lungs of tumor-bearing mice, and measure the resulting changes in lung pathophysiology by assessing primary lung
tumor growth, macrophage depletion, programing of remaining
macrophages, and BALF cytokine contents.

MATERIALS AND METHODS
MOUSE LUNG TUMORIGENESIS

Male A/J mice (6–8 weeks old) were purchased from Jackson Laboratories (Bar Harbor, ME, USA), maintained on hardwood bedding with a 12-h light/dark cycle, and given Teklad-8640 standard
laboratory chow (Harlan Teklad; Madison, WI, USA) and water
ad libitum. CCR2+/− breeding pairs on a BALB/cJ background
were kindly provided by Cara L. Mack, M.D. Department of Pediatrics, School of Medicine, University of Colorado, Anschutz Medical Center. BALB/cJ and CCR2−/− mice were bred in the Center
for Comparative Medicine (CCM) at the University of Colorado,
Anschutz Medical Center. A/J lung tumors were initiated by a single 1 mg/g body weight intraperitoneal (IP) injection of urethane
dissolved in 0.9% NaCl (Alfa Aesar; Heysham, Lancashire, UK) as
described previously (33). CCR2−/− and wild-type BALB/cJ mice
were given six weekly 1 mg/g IP injections of urethane, a regimen shown to reproducibly induce lung tumors in this moderately

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Macrophage depletion inhibits lung tumorigenesis

resistant strain (34). At the times indicated, mice were euthanized
by IP injection of sodium pentobarbital (Sigma Aldrich; St. Louis,
MO, USA). All animal procedures were performed in accordance
with the National Institutes of Health’s Guide for the Care and
Use of Laboratory Animals and were approved by the Institutional
Animal Care and Use Committee of the University of Colorado,
Anschutz Medical Campus.
BRONCHOALVEOLAR LAVAGE

Primary alveolar macrophages and lung protein exudates were
isolated by bronchoalveolar lavage (BAL), as previously described
(23, 35). The BALF fractions were separated by centrifugation,
BAL cells counted, and inflammatory macrophages, lymphocytes,
and neutrophils differentiated by Wright Giemsa (Fisher Scientific) staining (23). BAL cell populations from both naïve and lung
tumor-bearing mice were composed predominantly of alveolar
macrophages (routinely >95%) (10).
MACROPHAGE DEPLETION BY CLODRONATE-ENCAPSULATED
LIPOSOMES

Dichloromethylene diphosphonate (clodronate, 2.5 g; Sigma) was
encapsulated in liposomes formed by a 25:1 w/w ratio of phosphatidylcholine:cholesterol (Sigma) as described (31), and the
resulting liposomes resuspended in 4 ml sterile PBS. Only 1–2% of
clodronate becomes encapsulated, yielding an estimated dose of
0.7–1.0 mg clodronate per 100 µl of liposome suspension. Saline
(vehicle) liposomes and clodronate-encapsulated liposomes were
synthesized in parallel <2 weeks before use, stored at 4°C, and gently resuspended immediately before instillation or injection (31).
Clodronate liposomes for syngeneic transfer experiments were
synthesized or purchased from ClodronateLiposomes.org (The
Netherlands).
A/J mice bearing urethane-induced lung tumors were anesthetized by a single 50 µl IP injection containing 100 mg/kg ketamine and 10 mg/kg xylazine (CU Clinical Pharmacy; Aurora, CO,
USA). Fifty microliters of vehicle or clodronate-containing liposomes were instilled into the lungs via a ball-tip gavage needle bent
to a 30° angle and guided by a rodent laryngoscope (Penn Century, Inc.). Follow-up liposome treatments were administered by
IV in all mice starting 2 days after IT instillation (100 µl of vehicle or clodronate liposomes administered IV via the tail vein) and
repeated once weekly for 5 weeks. IT administration of clodronate
is necessary to deplete resident alveolar macrophages, which are
not exposed to IV administration, but recruitment of bone marrow
macrophages to replenish the alveolar macrophage population can
be prevented by IV clodronate liposome ablation of bone marrow
monocytes (31).
TISSUE COLLECTION

Plasma was obtained by retro-orbital bleeding with heparin-lined
capillary tubes (Fisher Scientific) following administration of terminal anesthesia, and stored at −80°C. Lungs were removed following BAL, the lobes dissociated, and tumors dissected from adjacent uninvolved lung under a dissection microscope, as described
(33). Tumor diameters were measured by digital calipers, and
tumors pooled in pre-tared microfuge tubes (one tube/mouse)
and weighed (tumor burden). Tumor dissection and evaluation
were conducted in a blinded fashion.

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IMMUNOHISTOCHEMISTRY, TUMOR GRADING, AND ASSESSMENT OF
PROLIFERATION

In a similarly treated group of mice, lungs were perfused via
the pulmonary artery with 0.9% NaCl, then gently inflated with
formalin through the cannulated trachea for 1 h. Lungs were
separated into individual lobes and dissected into 14 similarly
sized portions and fixed in formalin overnight (33). Lung pieces
were embedded in paraffin and sectioned (4 µm). Sections were
processed as previously described, and incubated with anti-Ki67
primary antibody (1:200; Fisher Scientific) (33). Incubation with
a biotinylated goat anti-rabbit secondary (1:100; Vector Laboratories) was followed by incubation with horse-radish peroxidase
conjugated avidin, detection with 3,3-diaminobenzidine (Vector
Laboratories), and counterstain with hematoxylin. Sections were
evaluated at 400× magnification under an upright microscope
(BX41 Olympus), using Spot Advanced software (v4.0.1) to determine tumor area (33). The Ki-67 staining index was calculated by
dividing the number of positively staining cells in each tumor by
the corresponding tumor area (Ki-67+ /cm2 ). This Ki-67 index was
averaged per animal and then per group. Serial lung sections were
stained with hematoxylin and eosin (Fisher Scientific), and lesions
graded as hyperplasia (Hyp), atypical adenomatous hyperplasia
(AAH), adenoma (AD), adenoma containing a focus of adenocarcinoma (ADwAC) or AC following the guidelines established
by Nikitin et al. (36) and using images found at the digital atlas
of virtual histological slides as examples (37). Grading was performed in a blinded fashion by three individuals at a multiheaded
microscope and evaluated by a board certified pathologist (Daniel
T. Merrick). Particular attention was paid to nuclear morphology,
density of the lesion, and vessel involvement. We found that adenomas and AC comprised the majority of lesions in this mouse
model, and squamous cell and neuroendocrine carcinomas were
not observed.
DETERMINATION OF MACROPHAGE PROGRAMING BY
IMMUNOFLUORESCENCE

Sections were deparaffinized and rehydrated prior to antigen
retrieval as described (23). Tissue sections were then incubated
overnight at 4°C with 1:50 dilution of anti-arginase I (ArgI,
Santa Cruz Biotechnology) primary antibody followed by 1:1000
dilution of Alexa 568-conjugated anti-goat secondary antibody.
A mixture of anti-NOS2 (BD Transduction Labs; 1:50 dilution) and anti-F4/80 (ABD-Serotec; 1:50 dilution) primary antibodies were then applied for 1 h at 37°C, followed by 20 min
incubations with Alexa 488-conjugated anti-rabbit and Alexa
680-conjugated anti-rat secondaries. Nuclei were stained with
DAPI-containing mounting media (Vector Laboratories). Images
were obtained with a digital deconvolution microscopy imaging system attached to a Zeiss Axioplan 2 epi-Fluorescence
upright microscope. Macrophages were identified by positive
F4/80 staining and morphology. Total pixel counts/macrophage
were calculated for ArgI, NOS2, and F4/80 immunofluorescence
(~50/animal) using ImageJ software (38), and ArgI and NOS2
values were normalized to F4/80 staining. To confirm ArgI+
M2 programing, adjacent sections were subjected to a similar IF protocol substituting an antibody against M2 marker
phosphoTyr641 STAT6 (Cell Signaling, 1:50 dilution) for NOS2.

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Macrophage depletion inhibits lung tumorigenesis

Fluorescence intensity was calculated similarly and phosphoSTAT6/ArgI ratios determined.
BALF IGF-I AND CYTOKINE DETERMINATION

Insulin-like growth factor-I concentrations in BALF were determined by enzyme-linked immunosorbant assay (ELISA) in a 96well format as directed (R&D Systems, Inc.). For lung cytokine
levels, BALF samples were concentrated 5× by centrifugation in
3 kDa molecular weight cut off YM-3 microcon spin-columns
(Millipore), and applied to Quantibody® mouse cytokine array
slides (Raybiotech, Inc.). Fluorescent Cy3-equivilent antibody signal was read by the CU Cancer Center Microarray Core Facility
using a Perkin Elmer Scan Array Ex glass slide laser scanner (Perkin
Elmer). Analytes were quantified by regression of log-transformed
data sets against within-run standard curves.
CCL2/CCR2 INVOLVEMENT IN LUNG TUMOR PROGRESSION

Tumorigenic mouse lung epithelial E9 cells (39) were maintained
in CMRL media (Invitrogen) supplemented with 10% fetal bovine
serum. For syngeneic transplant studies, 1 × 106 log-phase E9
cells were suspended in 100 µl of serum-free CMRL media (Life
Technologies) and injected subcutaneously (s.c.) into the shaved
right flanks of syngeneic male and female BALB/c (WT) or CCL2
receptor null (CCR2−/− ) mice, a protocol previously shown to
generate rapidly growing tumors in nearly 100% of recipient
animals (40) Wild-type and CCR2−/− mice received vehicle or
clodronate-encapsulated liposomes by IV injection 1 day prior to
tumor inoculation (day −1), and once/week thereafter. Tumor
size was determined twice/weekly for 24 days, and tumor volume was calculated using the equation for an elliptical cone (as
recommended due to the non-spherical growth pattern of the
implants): V = (d 2 × l × π)/6, in which “d” is the smallest diameter, and “l” the largest. Tumor-bearing mice were euthanized and
flank tumors removed and weighed. The experiment was performed twice, with 5–6 mice/group/repetition. Previous studies
showed that both macrophage conditioned media (MΦCM; 1:1
mixture of fresh media with media harvested after 24 h incubation with MH-S murine alveolar macrophage cells) and/or IGF-I
stimulated in vitro proliferation of cultured mouse lung epithelial
cells. To determine if this was also true for E9 cells, subconfluent cultures were incubated with MΦCM or 50 ng/ml IGF-I for
48 h, harvested, and relative cell numbers were compared using
CellTiter96® proliferation assays (Promega).
STATISTICAL ANALYSIS

Continuous variables were analyzed by two-way ANOVA with
Bonferroni post hoc comparison to determine significant differences between groups and account for multiple inter-group comparisons. One-way ANOVA with student Newman–Keuls post hoc
analysis was used to determine significant differences between
three or more groups while Student’s two-tailed independent t test was used when only two groups were compared, with Welch’s
correction for unequal variances when appropriate. All statistics
including Spearman correlations were performed using Prism 5.0
software (Graphpad; La Jolla, CA, USA). Data are presented as
mean ± SEM, unless otherwise indicated. In all analyses, p < 0.05
was considered to be statistically significant.

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FIGURE 1 | (A) Alveolar macrophage numbers increase as a function of
tumor growth. Alveolar macrophage (AM) content was determined in
naïve () and urethane-treated () A/J mice as a function of time (left
axis). Tumor burden, determined by tumor weight (
, right axis), increased
similarly over time (**p < 0.01 tumor-associated vs. naïve alveolar
macrophages). Few tumors were detected in naïve mice at any time point
(data not shown). (B–E) Mice (n = 4–6 mice/group) bearing
urethane-induced lung tumors were treated with clodronate-containing

RESULTS
Tumor burden and alveolar macrophage numbers (obtained by
lavage) increased similarly over time in urethane-treated A/J mice
(Figure 1A), while alveolar macrophage numbers changed little
over the same time course in naïve mice. Few tumors are detected
in naïve mice (data not shown). Liposomal clodronate significantly
depleted alveolar macrophages in tumor-bearing mice [harvested
32 (37%) and 44 (48%) weeks after urethane injection compared
to vehicle liposome treated, tumor-bearing mice (Figure 1B)].
Tumor burden at the 44-week time point decreased by ~50% with
clodronate treatment (Figure 1C) while tumor number did not
change (Figure 1D). Comparing tumor weight at 44 weeks to
that of mice sacrificed at 32 weeks suggests that tumors did not
regress with clodronate treatment, but simply did not grow as
rapidly (Figure 1C). A significant decrease in tumors with diameter >4 mm was detected, indicating that clodronate preferentially
slowed the growth of larger tumors (Figure 1E). Immunohistochemical staining of Ki67, a marker of cell division, decreased
by >50% in tumors from 44-week clodronate-treated mice compared to the vehicle liposome-treated controls (Figures 2A,B),
affecting tumors of varying size (Figure 2D). Similar numbers of
hyperplasias, AAH, adenomas, adenomas with AC-like foci, and
AC (Figures 2C,E) were detected in the lungs of both vehicle and
clodronate-treated mice.
Programing in tumor-associated macrophages (TAMs) was
examined by immunofluorescence to determine whether clodronate liposomes targeted a specific subset of macrophages in
mice bearing 44-week lung tumors. TAMs in this study exhibited a mixed M1/M2 phenotype characterized by both NOS2 (an
M1 programing marker) and ArgI (an M2 programing marker)

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Macrophage depletion inhibits lung tumorigenesis

liposomes (black bars) or the vehicle control (gray bars) for 5 weeks prior to
lung harvest at 32 weeks and 44 weeks. (B) Alveolar macrophages are
depleted by clodronate treatment (**p < 0.01 vs. vehicle) at both 32 and
44 weeks. (C) Tumor burden decreases with macrophage depletion at
44 weeks, but not 32 weeks. (D) Tumor number remains constant
regardless of age or clodronate treatment. (E) Tumor multiplicity at
44 weeks broken down by tumor size. Significant difference (*p < 0.05)
between treatment groups occurred only in tumors >4 mm in diameter.

expression (Figure 3A). Because earlier studies showed predominantly NOS2+ macrophages in this model at this late time point, a
second M2 marker, phosphoSTAT6, was analyzed confirming the
presence of M2 markers (Figure 3B). Previously, we determined
that human lung TAMs express similar mixed (NOS2+ CD206+ )
programing (23). We saw no large-scale differences in programing marker expression between vehicle and clodronate exposed
macrophages (Figure 3C), and ArgI/phosphoSTAT6 ratios were
also similar between groups (Figure 3C) suggesting that the TAMs
in this model are similarly programed, and this programing is not
affected by clodronate exposure.
Cytokine and IGF-I levels were measured in BALF from vehicle
and clodronate-treated, tumor-bearing mice 44 weeks after urethane. BALF levels of GM-CSF, IFN-γ, IL-1α, IL-1β, IL-2, IL-4, IL5, IL-9, IL-10, IL-12, IL-17, M-CSF, and RANTES were unchanged
by clodronate treatment (data not shown). IL-3, IL-13, and TNF-α
contents were below the limit of detection in all samples (data not
shown). Clodronate treatment significantly decreased levels of IL6, CCL2, CXCL1, and IGF-I, while VEGF levels increased 1.5-fold
(Figures 4A,B). Serum levels of IGF-I did not change with clodronate exposure (Figure 4B). Levels of IGF-I, CXCL1, IL-6, and
CCL2 were higher in BALF from tumor-bearing mice than naïve
mice 32 weeks after urethane treatment, but VEGF levels were
unchanged (Figure 4C). IGF-I levels correlate significantly with
BAL macrophage numbers in naïve and tumor-bearing animals
(Figure 4D, p < 0.0001) as well as in animals exposed to vehicle
and clodronate-containing liposomes (Figure 4E, p < 0.03). Activated macrophages produce IL-6, CCL2, CXCL1, and IGF-I, but
concomitant production of all four signaling molecules also indicates a “mixed phenotype” of macrophages since IL-6 and IL-8

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FIGURE 2 | Effects of clodronate treatment on tumor growth and
progression. (A) Representative Ki67 IHC on tissue sections from similar
sized tumors from vehicle (left) and clodronate (right) treated mice (400×
final magnification). Red arrows point to positively stained tumor cells.
(B) Ki-67 index was calculated as the average number of Ki-67+ cells/cm2
tumor area for each tumor in each group (mean ± SEM, **p < 0.01 vs.

(human ortholog of murine CXCL1) are most often associated
with M1 programing and IFG-I and CCL2 with M2 programing.
Vehicle treated mice secreted 6.5-fold more CCL2 into BALF
compared to clodronate-treated mice (Figure 4A). As CCL2 is
a chemotactive factor for monocytes and could be integral in the
recruitment of macrophages to the site of tumors development, we
tested whether ablation of CCL2/CCR2 signaling affected tumor
growth in a syngeneic transplant model. Transformed mouse lung
epithelial E9 cells were injected into the flanks of immunocompetent syngeneic BALB/cJ wild-type or CCR2−/− mice. After tumors
were established, mice were subjected to systemic liposome-based
macrophage depletion. There were no significant differences in
tumor growth among the vehicle treated wild-type, vehicle treated
CCR2−/− , or clodronate-treated CCR2−/− mice. However, flank
tumors in wild-type mice receiving clodronate liposomes grew
significantly slower (Figure 5A). We previously showed that
macrophage conditioned media (MΦCM) increases proliferation
of mouse lung epithelial cells in vitro, and IGF-I is the major

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Macrophage depletion inhibits lung tumorigenesis

vehicle). (C) Representative examples of each tumor grade (400× final
magnification). (D) Ki67 index as a function of tumor size at 44 weeks in
saline and clodronate-treated mouse lung tumors. Significant differences
were seen in the number of Ki67+ cells in the smaller lung tumors.
(E) Percent of each lesion type was calculated/mouse. No significant
differences were detected.

component in MΦCM that contributes to this proliferative effect
(26). Both MΦCM and IGF-I increased E9 proliferation by more
than threefold over control media suggesting that the inhibition
of syngeneic tumor growth, resulting from systemic macrophage
depletion may be due to decreased macrophage production of
IGF-I (Figure 5B).
Ablation of CCL2/CCR2 signaling did not affect the growth
of syngeneically transplanted E9 cells, and although there was a
slight, but not significant trend of slower growth of the E9 tumors
in the CCR2−/− mice, the effect was not as dramatic as that seen
with clodronate treatment. In addition, tumor growth in CCR2−/−
mice was not significantly slowed by clodronate exposure. To test
whether CCL2/CCR2 signaling is required for the de novo development of tumors in the lung, wild-type and CCR2−/− mice were
initiated with six weekly urethane injections. Lung tumors were
harvested, counted, and tumor burden (by weight) assessed at 20,
32, 38, and 42 weeks after the initial urethane exposure. No significant decreases in tumor number, tumor burden, or alveolar

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FIGURE 3 | Immunofluorescent determination of macrophage
programing. (A) Alveolar macrophage ArgI (red), NOS2 (green), and F4/80
(blue) expression and the merged image (showing co-localization) in
tumor-bearing mice treated with saline or clodronate-containing liposomes.
(B) ArgI (red), STAT6 (green), and F4/80 (blue) expression and the merged
image showing co-localization in tumor-bearing mouse lung treated with
saline or clodronate-containing liposomes. (C) Pixel counts/macrophage of
ArgI (red) or NOS2 (green) normalized to F4/80 (blue) or STAT6 (blue)/ArgI
ratios in saline (open bars) or clodronate (cross-hatched) liposome-treated
lung (mean ± SEM, n = 135–150 macrophages/group).

macrophage numbers were detected in CCR2−/− mice at any time
point (data not shown). Results from the 42-week time point are
shown in Figures 5C–F. Not only did CCR2−/− mice show similar number of alveolar macrophages and lung tumors at this late
time point, there was a significant increase in lung tumor diameter in the CCR2−/− mice. As CCL2/CCR2 signaling was ablated
in the CCR2−/− mice, either a redundant pathway compensates
for macrophage recruitment to the lung or resident macrophages
proliferate to compensate for the lack of CCL2/CCR2-mediated
macrophage recruitment.

DISCUSSION
Depletion of pulmonary macrophages by nearly 50% decreased
the growth of the largest lung ACs in the A/J urethane model.
Decreased Ki67 staining indicated that cell division slowed as a
result of macrophage depletion in vivo, an observation complementing previous results showing that macrophage co-culture and

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Macrophage depletion inhibits lung tumorigenesis

exposure to macrophage conditioned media increased tumor cell
proliferation in vitro (26). Macrophage depletion did not cause
tumor regression as tumor number and stage did not change after
clodronate treatment. TAMs produce signals that support tumor
growth and promote tumor cell survival. When TAMs are depleted,
production of these signals decreases causing a reduction in tumor
cell proliferation. Clodronate exposure decreased BALF levels of
IGF-I, IL-6, CXCL1, and CCL2 and increased VEGF levels while
not affecting most of the other factors examined (largely TH 1 and
TH 2-associated cytokines), suggesting that alveolar macrophages
may not be their primary source in murine lungs. IGF-I involvement in lung tumorigenesis is well established, and the positive
correlation of pulmonary BAL IGF-I levels with macrophage numbers (Figures 4C,D) suggests that macrophage production of IGFI is important in maintaining tumor growth. The decrease in IGF-I
levels caused by clodronate-induced macrophage depletion may be
partially responsible for slowing lung tumor growth in vivo, which
is consistent with our previous observations of IGF-I mediation
of a significant portion of MΦCM induced lung tumor cell proliferation in vitro. The increased BALF VEGF levels detected in the
clodronate-treated animals may occur in response to clodronateinduced macrophage apoptosis, as VEGF expression is induced in
macrophages that clear apoptotic cells by efferocytosis (41, 42).
The remaining healthy alveolar macrophages may express more
VEGF as they clear the apoptotic macrophages, resulting from
clodronate depletion.
The attenuation of CCL2 production caused by clodronate
liposome exposure suggested that monocyte recruitment might
also play a role in tumor progression. CCL2 (MCP-1; monocyte
chemotactic protein-1) is involved in recruitment of monocytes
(43), T cells (44), and dendritic cells (45) to areas of inflammation induced by tissue injury or infection. High levels of CCL2
are associated with poor prognosis in breast cancer (46) and
pancreatic cancer (47). However, Zhang et al. (48) found that
CCL2 over-expression is associated with improved survival in
NSCLC patients indicating that CCL2/CCR2 signaling may have
different roles in different tissues. To determine whether CCL2
signaling affected lung tumor growth in mice, syngeneic mouse
lung tumor cells were transplanted into wild-type and CCR2−/−
mice. Ablation of CCL2/CCR2 signaling had no effect tumor
growth, and the lack of CCL2/CCR2 signaling actually nullified
the growth inhibitory effects of clodronate liposomes for reasons,
which remain to be determined. Because E9 cells are already transformed and do not require a progression phase for tumor growth
and because alveolar macrophages differ in function and response
from macrophages present near subcutaneous tumors (49), we
tested whether CCL2/CCR2 signaling was required for de novo
lung tumor formation. BAL macrophage content and tumor numbers from urethane-exposed CCR2−/− and wild-type mice were
similar between genotypes at each time point. A slight, but significant increase in tumor size in the CCR2−/− mice compared
to wild-type littermates was detected at the 44-week time point,
which is consistent with the poorer survival observed in human
NSCLC patients with low-CCL2 expression (48). Macrophages
are known to produce CCL2 in response to IL-4/IL-13 stimulation
(50), and the increase in CCL2 production we detected during
lung tumorigenesis indicates that there is a Th2 response occurring

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FIGURE 4 | Effects of macrophage depletion on BAL cytokine
content. Cytokine levels were determined in BALF from naïve (white
bars), vehicle liposome (gray bars), or clodronate liposome (black bars)
treated mice 44 weeks after urethane treatment. (A) Clodronate
administration decreased IL-6, CCL2, and CXCL1 in BALF, while VEGF
increased 1.5-fold. Data presented as mean ± SEM, n = 4–6 mice/group
(*p < 0.05 vs. vehicle treated mice). (B) BALF IGF-I levels decreased
with clodronate treatment in AC-bearing mice (***p < 0.001 vs.

in lungs during tumor formation. The CCL2 receptor (CCR2) is
expressed only in monocyte-lineage cells and T lymphocytes, so
CCL2 is unlikely to directly affect neoplastic epithelial cells (9).
CCL2 may regulate macrophage recruitment following lung injury,
as CCL2 levels in BALF are elevated just prior to the macrophage
influx that follows chemically induced pneumotoxicity (51). The
similar alveolar macrophage content in tumor-bearing lungs from
both wild-type and CCR2−/− mice suggests that either there are
redundant macrophage recruitment pathways at play in the lung
tumor microenvironment or that the increased number of TAMs
results from proliferation of resident macrophages rather than
recruitment as has been reported (52).
High-serum IL-6 levels correlate with poor survival and poor
response to chemotherapy in NSCLC patients (53, 54). IL-6 is an
inflammatory cytokine produced primarily by T cells and M1programed macrophages to stimulate immune response to injury
and trauma. IL-6 may be required for M1 programing in some
macrophages, but IL-6 production can induce M2 programing
in certain systems to limit existing inflammation (55). Recently,
Fernando et al. showed that M2 programing is enhanced by IL-6

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Macrophage depletion inhibits lung tumorigenesis

vehicle). (C) Fold change in BALF IGF-I, IL-6, CXCL1, CCL2, and VEGF
levels in age-matched naïve and tumor-bearing mice 32 weeks after
urethane exposure (*p < 0.05) (D) Correlation between IGF-I and BAL
macrophage number in naïve and urethane-treated mice 24 and
32 weeks after urethane treatment (Spearman ρ = 0.8055, p < 0.0001).
(E) Correlation between IGF-I and BAL macrophage number in 44-week
urethane mice treated with saline and clodronate liposomes (Spearman
ρ = 0.4990, p < 0.0031).

exposure in cultured macrophages and suggested that IL-6 augments cytokine expression during both M1 and M2 programing
(56). Similar to our findings, Karnevi et al. report that human
pancreatic tumor-educated macrophages display mixed M1/M2
programing and produce increased IL-6 and IL-8 (57). Pine et al.
report that increased IL-6 and IL-8 production are associated with
increased lung cancer risk (58), and IL-6 and IL-8 are both associated with increased NSCLC cell proliferation (59) although neither
directly induces increased cell division. Human IL-8 (CXCL8)
and murine keratinocyte-derived chemokine CXCL1 (KC) are
orthologous in function. CXCL1 production is high in murine
alveolar macrophages (60, 61), but not in peritoneal macrophages
(62–65). The specific combination of cytokines and surfactant
proteins intrinsic to the lung make alveolar macrophages functionally unique compared to macrophages from other tissues. Lung
production of IL-8/CXCL1 is induced by inflammatory stimuli
through NF-κB and AP-1 activation (63, 65), and both NSCLC
cells and macrophages express IL-8/CXCL1. IL-8/CXCL1 is a proangiogenic factor as well as a chemotactic factor for neutrophil
recruitment to the lungs during emphysema and lung cancer. Few

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FIGURE 5 | Clodronate reduces syngeneic implant growth in
wild-type (WT), but not CCR2−/− mice. (A) E9 cells were injected into
the flank of female BALB/cJ (WT) or CCR2−/− mice. After tumors were
established (8 days post injection), clodronate-containing or vehicle
liposomes were injected IV once/week until sacrifice. Tumors were
measured 2×/week and volumes calculated. Clodronate liposome
treatment (Cdr) decreased tumor growth in wild-type mice (*p < 0.05,
n = 10–12 mice/group from two independent experiments). (B) E9 cells

neutrophils were detected in the lungs of vehicle or clodronate
liposome-treated tumor-bearing mice (data not shown), so we
could not determine whether clodronate treatment altered their
numbers further.
Macrophage programing in mouse lung is homogeneous, probably due to the small size of the organ. As urethane-exposed A/J
mice form >30 tumors/mouse and these tumors are spread evenly
throughout the lobes, alveolar macrophages are exposed to tumor
produced factors throughout the lung. The A/J urethane model
of lung cancer is unique among most murine lung tumor models in that TAMs surrounding late-stage ACs express high levels
of NOS2 and little ArgI, indicating that they are primarily M1programed (23, 24, 34). However, BALF cytokine levels in these
mice indicate that macrophage programing might be more complex. Although ArgI expression decreases as tumors progress, production of certain M2-associated signaling molecules (i.e., IGF-1,
CCL2) increases. When these mice were exposed to either vehicle or clodronate-containing liposomes, we detected an increase
in both ArgI and phosphoSTAT6 expression (M2 programing
markers) in the same macrophages that maintained high-NOS2
expression. The M1/M2 classification represents a continuum
of plasticity and does not encompass the functional diversity of

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Macrophage depletion inhibits lung tumorigenesis

were cultured in serum-free media (vehicle), 50 ng/ml IGF-I containing
media, or macrophage conditioned media (MΦCM). Cell density was
determined by MTS and graphed as fold-increase over control
(mean ± SEM, ***p < 0.001 vs. control). (C–F) Wild-type BALB/cJ and
CCR2−/− mice were injected with 1 mg/g urethane once/week for 6 weeks.
BALF and tumors were harvested 42 weeks after the first injection and
BAL macrophages (×1000) counted, and lung tumors counted, measured,
and weighed (mean ± SEM, *p < 0.05 vs. wild-type, n = 5).

macrophages (66), and the measurement of biomarker expression
rather than activity and/or function may not yield an accurate
picture of macrophage programing. In our study, we detected
NOS2+ ArgI+ macrophages, but we did not measure NO levels
or determine arginase activity, so although both enzymes were
present, we do not know that both were active in the same cells.
The presence of phosphoSTAT6 in these TAMs indicates that they
were more “M2-like” and the presence of CCR2 and IGF-1 in the
BALF supports this. Although there were fewer macrophages in the
clodronate-treated lungs, the ratio of ArgI to NOS2 remained constant in the remaining cells indicating that the remaining tumor
microenvironment and not macrophage depletion continued to
affect the programing of the remaining macrophages.
Our previous studies indicated that TAMs near human lung
AC also expressed M1 and M2 markers simultaneously (24), and
others have seen similar mixed phenotypes in TAMs in pancreatic (57) and gastric cancer (67). Mixed macrophage programing
was also demonstrated in the resolution of liver fibrosis (68) and
pulmonary fibrosis (69) and in the early stages of diet induced
obesity (70). These mixed populations could be due to catching the macrophages as they change from one programing state
to another as in the resolution of a disease, or as they respond

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to conflicting microenvironmental signals. The detection of both
inflammatory and anti-inflammatory cytokines in BALF from
these mice indicates that these macrophages do not follow the
canonical roles of M1 or M2 macrophages. Further research to
determine phagocytic and efferocytic activity, proliferative capacity, and gene expression of TAMs from tumor-bearing mice before
and after exposure to vehicle and clodronate liposomes is necessary to characterize this novel macrophage population. The ability
of clodronate liposomes to deplete pulmonary macrophages may
be enhanced by changing the composition of the liposome delivery vehicle to more effectively and specifically target TAMs that
accumulate in the lungs of tumor-bearing mice. These mixed phenotype macrophages may express scavenger receptors such as the
mannose receptor, so adding mannose to the liposome surface
may increase their uptake, resulting in greater depletion (71). Also,
M2 macrophages exhibit enhanced efferocytosis (72) while M1
macrophages are professional phagocytes, so the presence of phosphatidyl serine in the liposome membrane to mimic apoptotic
cells and/or lipopolysaccharides on the exterior of the liposome
to mimic bacterial cell walls may lead to enhanced uptake and
possibly increased macrophage depletion. As IGF-I signaling activates survival factors such as AKT in tumor cells (26), macrophage
depletion and the subsequent decrease in BALF IGF-I levels may
make tumors more sensitive to therapy-induced death. Although
macrophage depletion cannot eliminate lung tumors, it may sensitize tumors to classic chemotherapeutics and radiation by removing both macrophage-produced survival factors from the lung
prior to cytotoxic therapies and growth promoting signals such as
IGF-I during tumor recovery. Finally, by decreasing the permissive
nature of the tumor microenvironment, macrophage depletion
may allow the host’s own defenses to recover and eliminate tumor
cells.

ACKNOWLEDGMENTS
This paper is dedicated to the memory of our late mentor, colleague, and friend Dr. Alvin M. Malkinson whose work with
inflammation and tumor promotion formed the basis for these
studies. These studies were funded by NIH CA132552 and supported by NCI Cancer Center Support Grant P30CA046934
(Microarray Core Facility).

REFERENCES
1. Ferlay J, Shin HR, Bray F, Forman D, Mathers C, Parkin DM. Estimates of
worldwide burden of cancer in 2008: GLOBOCAN 2008. Int J Cancer (2010)
127(12):2893–917. doi:10.1002/ijc.25516
2. Shopland DR, Eyre HJ, Pechacek TF. Smoking-attributable cancer mortality in
1991: is lung cancer now the leading cause of death among smokers in the
United States? J Natl Cancer Inst (1991) 83(16):1142–8. doi:10.1093/jnci/83.16.
1142
3. Peebles KA, Lee JM, Mao JT, Hazra S, Reckamp KL, Krysan K, et al. Inflammation and lung carcinogenesis: applying findings in prevention and treatment.
Expert Rev Anticancer Ther (2007) 7(10):1405–21. doi:10.1586/14737140.7.10.
1405
4. David O, Jett J, LeBeau H, Dy G, Hughes J, Friedman M, et al. PhosphoAkt overexpression in non-small cell lung cancer confers significant stageindependent survival disadvantage. Clin Cancer Res (2004) 10(20):6865–71.
doi:10.1158/1078-0432.CCR-04-0174
5. Ginsberg R, Vokes R, Rosenweig K. Non-small cell lung cancer. In: De Vitta V,
Hellman S, Rosenberg S, editors. Cancer: Principles and Practice of Oncology.
Philadelphia, PA: Lippencott Williams and Wilkins (2001). p. 925–83.

www.frontiersin.org

Macrophage depletion inhibits lung tumorigenesis

6. Schreinemachers DM, Everson RB. Aspirin use and lung, colon, and breast
cancer incidence in a prospective study. Epidemiology (1994) 5(2):138–46.
doi:10.1097/00001648-199403000-00003
7. Rothwell PM, Fowkes FG, Belch JF, Ogawa H, Warlow CP, Meade TW. Effect
of daily aspirin on long-term risk of death due to cancer: analysis of individual patient data from randomised trials. Lancet (2011) 377(9759):31–41.
doi:10.1016/S0140-6736(10)62110-1
8. Coussens LM, Werb Z. Inflammation and cancer. Nature (2002)
420(6917):860–7. doi:10.1038/nature01322
9. Miotto D, Boschetto P, Bononi I, Milani G, Legorini C, Cavallesco G, et al.
CC ligand 2 levels are increased in LPS-stimulated peripheral monocytes of
patients with non-small cell lung cancer. Respir Med (2007) 101(8):1738–43.
doi:10.1016/j.rmed.2007.02.021
10. Condeelis J, Pollard JW. Macrophages: obligate partners for tumor cell migration, invasion, and metastasis. Cell (2006) 124(2):263–6. doi:10.1016/j.cell.2006.
01.007
11. Redente EF, Higgins DM, Dwyer-Nield LD, Orme IM, Gonzalez-Juarrero
M, Malkinson AM. Differential polarization of alveolar macrophages and
bone marrow-derived monocytes following chemically and pathogen-induced
chronic lung inflammation. J Leukoc Biol (2010) 88(1):159–68. doi:10.1189/jlb.
0609378
12. Zaynagetdinov R, Sherrill TP, Polosukhin VV, Han W, Ausborn JA, McLoed AG,
et al. A critical role for macrophages in promotion of urethane-induced lung
carcinogenesis. J Immunol (2011) 187(11):5703–11. doi:10.4049/jimmunol.
1100558
13. Bauer AK, Dwyer-Nield LD, Keil K, Koski K, Malkinson AM. Butylated hydroxytoluene (BHT) induction of pulmonary inflammation: a role in tumor promotion. Exp Lung Res (2001) 27(3):197–216. doi:10.1080/01902140120756
14. Li H, Sorenson AL, Poczobutt J, Amin J, Joyal T, Sullivan T, et al. Activation of
PPARgamma in myeloid cells promotes lung cancer progression and metastasis.
PLoS One (2011) 6(12):e28133. doi:10.1371/journal.pone.0028133
15. Malkinson AM. Primary lung tumors in mice as an aid for understanding, preventing, and treating human adenocarcinoma of the lung. Lung Cancer (2001)
32(3):265–79. doi:10.1016/S0169-5002(00)00232-4
16. Moody TW, Leyton J, Zakowicz H, Hida T, Kang Y, Jakowlew S, et al.
Indomethacin reduces lung adenoma number in A/J mice. Anticancer Res (2001)
21(3B):1749–55.
17. Dziadziuszko R, Camidge DR, Hirsch FR. The insulin-like growth factor
pathway in lung cancer. J Thorac Oncol (2008) 3(8):815–8. doi:10.1097/JTO.
0b013e31818180f5
18. Chitnis MM, Yuen JS, Protheroe AS, Pollak M, Macaulay VM. The type 1 insulinlike growth factor receptor pathway. Clin Cancer Res (2008) 14(20):6364–70.
doi:10.1158/1078-0432.CCR-07-4879
19. Olbruck H, Seemayer NH, Voss B, Wilhelm M. Supernatants from quartz dust
treated human macrophages stimulate cell proliferation of different human lung
cells as well as collagen-synthesis of human diploid lung fibroblasts in vitro.
Toxicol Lett (1998) 9(6–97):85–95. doi:10.1016/S0378-4274(98)00054-X
20. Bitterman PB, Rennard SI, Hunninghake GW, Crystal RG. Human alveolar
macrophage growth factor for fibroblasts. Regulation and partial characterization. J Clin Invest (1982) 70(4):806–22. doi:10.1172/JCI110677
21. Allen JT, Bloor CA, Kedia RK, Knight RA, Spiteri MA. Expression of growth
hormone-releasing factor, growth hormone, insulin-like growth factor-1 and
its binding proteins in human lung. Neuropeptides (2000) 34(2):98–107.
doi:10.1054/npep.2000.0802
22. Nyman T, Pekonen F. The expression of insulin-like growth factors and their
binding proteins in normal human lymphocytes. Acta Endocrinol (Copenh)
(1993) 128(2):168–72.
23. Redente EF, Orlicky DJ, Bouchard RJ, Malkinson AM. Tumor signaling to
the bone marrow changes the phenotype of monocytes and pulmonary
macrophages during urethane-induced primary lung tumorigenesis in A/J mice.
Am J Pathol (2007) 170(2):693–708. doi:10.2353/ajpath.2007.060566
24. Redente EF, Dwyer-Nield LD, Merrick DT, Raina K, Agarwal R, Pao W,
et al. Tumor progression stage and anatomical site regulate tumor-associated
macrophage and bone marrow-derived monocyte polarization. Am J Pathol
(2010) 176(6):2972–85. doi:10.2353/ajpath.2010.090879
25. Martinez FO, Gordon S, Locati M, Mantovani A. Transcriptional profiling of the
human monocyte-to-macrophage differentiation and polarization: new molecules and patterns of gene expression. J Immunol (2006) 177(10):7303–11.
doi:10.4049/jimmunol.177.10.7303

November 2014 | Volume 5 | Article 587 | 125

Fritz et al.

26. Fritz JM, Dwyer-Nield LD, Malkinson AM. Stimulation of neoplastic mouse lung
cell proliferation by alveolar macrophage-derived, insulin-like growth factor-1
can be blocked by inhibiting MEK and PI3K activation. Mol Cancer (2011)
10:76. doi:10.1186/1476-4598-10-76
27. Frankel SK, Moats-Staats BM, Cool CD, Wynes MW, Stiles AD, Riches DW.
Human insulin-like growth factor-IA expression in transgenic mice promotes
adenomatous hyperplasia but not pulmonary fibrosis. Am J Physiol Lung Cell
Mol Physiol (2005) 288(5):L805–12. doi:10.1152/ajplung.00420.2004
28. Hiraoka K, Zenmyo M, Watari K, Iguchi H, Fotovati A, Kimura YN, et al. Inhibition of bone and muscle metastases of lung cancer cells by a decrease in
the number of monocytes/macrophages. Cancer Sci (2008) 99(8):1595–602.
doi:10.1111/j.1349-7006.2008.00880.x
29. Thepen T, Van Rooijen N, Kraal G. Alveolar macrophage elimination in vivo is
associated with an increase in pulmonary immune response in mice. J Exp Med
(1989) 170(2):499–509. doi:10.1084/jem.170.2.499
30. Zeisberger SM, Odermatt B, Marty C, Zehnder-Fjallman AH, Ballmer-Hofer
K, Schwendener RA. Clodronate-liposome-mediated depletion of tumourassociated macrophages: a new and highly effective antiangiogenic therapy
approach. Br J Cancer (2006) 95(3):272–81. doi:10.1038/sj.bjc.6603240
31. Van RN, Hendrikx E. Liposomes for specific depletion of macrophages from
organs and tissues. Methods Mol Biol (2010) 605:189–203. doi:10.1007/978-160327-360-2_13
32. Van RN, Kors N, Kraal G. Macrophage subset repopulation in the spleen: differential kinetics after liposome-mediated elimination. J Leukoc Biol (1989)
45(2):97–104.
33. Zerbe LK, Dwyer-Nield LD, Fritz JM, Redente EF, Shroyer RJ, Conklin E, et al.
Inhibition by erlotinib of primary lung adenocarcinoma at an early stage in male
mice. Cancer Chemother Pharmacol (2008) 62(4):605–20. doi:10.1007/s00280007-0644-z
34. Redente EF, Dwyer-Nield LD, Barrett BS, Riches DW, Malkinson AM. Lung
tumor growth is stimulated in IFN-gamma-/- mice and inhibited in IL-4Ralpha/- mice. Anticancer Res (2009) 29(12):5095–101.
35. Bauer AK, Dwyer-Nield LD, Hankin JA, Murphy RC, Malkinson AM. The lung
tumor promoter, butylated hydroxytoluene (BHT), causes chronic inflammation in promotion-sensitive BALB/cByJ mice but not in promotion-resistant
CXB4 mice. Toxicology (2001) 169(1):1–15. doi:10.1016/S0300-483X(01)
00475-9
36. Nikitin AY,Alcaraz A,Anver MR, Bronson RT, Cardiff RD, Dixon D, et al. Classification of proliferative pulmonary lesions of the mouse: recommendations of the
mouse models of human cancers consortium. Cancer Res (2004) 64(7):2307–16.
doi:10.1158/0008-5472.CAN-03-3376
37. eMICE/NCI. Proliferative Lesions of the Lung [Internet] National Cancer Institute, U.S. National Institutes of Health (2014). Available from:
http://emice.nci.nih.gov/camod-models-by-organ-site/proliferative-lesions-ofthe-lung/proliferative-lesions-of-the-lung
38. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of
image analysis. Nat Methods (2012) 9(7):671–5. doi:10.1038/nmeth.2089
39. Malkinson AM, Dwyer-Nield LD, Rice PL, Dinsdale D. Mouse lung epithelial
cell lines – tools for the study of differentiation and the neoplastic phenotype.
Toxicology (1997) 123(1–2):53–100. doi:10.1016/S0300-483X(97)00108-X
40. Porter SE, Dwyer-Nield LD, Malkinson AM. Regulation of lung epithelial cell
morphology by cAMP-dependent protein kinase type I isozyme. Am J Physiol
Lung Cell Mol Physiol (2001) 280(6):L1282–9.
41. Golpon HA, Fadok VA, Taraseviciene-Stewart L, Scerbavicius R, Sauer C,
Welte T, et al. Life after corpse engulfment: phagocytosis of apoptotic cells
leads to VEGF secretion and cell growth. FASEB J (2004) 18(14):1716–8.
doi:10.1096/fj.04-1853fje
42. Kearns MT, Dalal S, Horstmann SA, Richens TR, Tanaka T, Doe JM, et al. Vascular endothelial growth factor enhances macrophage clearance of apoptotic cells.
Am J Physiol Lung Cell Mol Physiol (2012) 302(7):L711–8. doi:10.1152/ajplung.
00116.2011
43. Rollins BJ, Walz A, Baggiolini M. Recombinant human MCP-1/JE induces
chemotaxis, calcium flux, and the respiratory burst in human monocytes. Blood
(1991) 78(4):1112–6.
44. Carr MW, Roth SJ, Luther E, Rose SS, Springer TA. Monocyte chemoattractant
protein 1 acts as a T-lymphocyte chemoattractant. Proc Natl Acad Sci U S A
(1994) 91(9):3652–6. doi:10.1073/pnas.91.9.3652

Frontiers in Immunology | Molecular Innate Immunity

Macrophage depletion inhibits lung tumorigenesis

45. Xu LL, Warren MK, Rose WL, Gong W, Wang JM. Human recombinant
monocyte chemotactic protein and other C-C chemokines bind and induce
directional migration of dendritic cells in vitro. J Leukoc Biol (1996) 60(3):
365–71.
46. Ueno T, Toi M, Saji H, Muta M, Bando H, Kuroi K, et al. Significance of macrophage chemoattractant protein-1 in macrophage recruitment,
angiogenesis, and survival in human breast cancer. Clin Cancer Res (2000)
6(8):3282–9.
47. Sanford DE, Belt BA, Panni RZ, Mayer A, Deshpande AD, Carpenter D, et al.
Inflammatory monocyte mobilization decreases patient survival in pancreatic cancer: a role for targeting the CCL2/CCR2 axis. Clin Cancer Res (2013)
19(13):3404–15. doi:10.1158/1078-0432.CCR-13-0525
48. Zhang XW, Qin X, Qin CY, Yin YL, Chen Y, Zhu HL. Expression of monocyte
chemoattractant protein-1 and CC chemokine receptor 2 in non-small cell lung
cancer and its significance. Cancer Immunol Immunother (2013) 62(3):563–70.
doi:10.1007/s00262-012-1361-y
49. Hussell T, Bell TJ. Alveolar macrophages: plasticity in a tissue-specific context.
Nat Rev Immunol (2014) 14(2):81–93. doi:10.1038/nri3600
50. Ip WK, Wong CK, Lam CW. Interleukin (IL)-4 and IL-13 up-regulate monocyte chemoattractant protein-1 expression in human bronchial epithelial cells:
involvement of p38 mitogen-activated protein kinase, extracellular signalregulated kinase 1/2 and Janus kinase-2 but not c-Jun NH2-terminal kinase
1/2 signalling pathways. Clin Exp Immunol (2006) 145(1):162–72. doi:10.1111/
j.1365-2249.2006.03085.x
51. Meyer AM, Dwyer-Nield LD, Hurteau G, Keith RL, Ouyang Y, Freed BM,
et al. Attenuation of the pulmonary inflammatory response following butylated hydroxytoluene treatment of cytosolic phospholipase A2 null mice.
Am J Physiol Lung Cell Mol Physiol (2006) 290(6):L1260–6. doi:10.1152/ajplung.
00182.2005
52. Jenkins SJ, Ruckerl D, Cook PC, Jones LH, Finkelman FD, van RN, et al. Local
macrophage proliferation, rather than recruitment from the blood, is a signature
of TH2 inflammation. Science (2011) 332(6035):1284–8. doi:10.1126/science.
1204351
53. De VF, Orditura M, Auriemma A, Infusino S, Roscigno A, Catalano G. Serum
levels of interleukin-6 as a prognostic factor in advanced non-small cell lung
cancer. Oncol Rep (1998) 5(3):649–52.
54. Chang CH, Hsiao CF, Yeh YM, Chang GC, Tsai YH, Chen YM, et al. Circulating interleukin-6 level is a prognostic marker for survival in advanced nonsmall cell lung cancer patients treated with chemotherapy. Int J Cancer (2013)
132(9):1977–85. doi:10.1002/ijc.27892
55. Mauer J, Chaurasia B, Goldau J, Vogt MC, Ruud J, Nguyen KD, et al. Signaling
by IL-6 promotes alternative activation of macrophages to limit endotoxemia
and obesity-associated resistance to insulin. Nat Immunol (2014) 15(5):423–30.
doi:10.1038/ni.2865
56. Fernando MR, Reyes JL, Iannuzzi J, Leung G, McKay DM. The proinflammatory cytokine, interleukin-6, enhances the polarization of alternatively
activated macrophages. PLoS One (2014) 9(4):e94188. doi:10.1371/journal.
pone.0094188
57. Karnevi E, Andersson R, Rosendahl AH. Tumour-educated macrophages display
a mixed polarisation and enhance pancreatic cancer cell invasion. Immunol Cell
Biol (2014) 92(6):543–52. doi:10.1038/icb.2014.22
58. Pine SR, Mechanic LE, Enewold L, Chaturvedi AK, Katki HA, Zheng YL,
et al. Increased levels of circulating interleukin 6, interleukin 8, C-reactive
protein, and risk of lung cancer. J Natl Cancer Inst (2011) 103(14):1112–22.
doi:10.1093/jnci/djr216
59. Luppi F, Longo AM, de Boer WI, Rabe KF, Hiemstra PS. Interleukin-8 stimulates cell proliferation in non-small cell lung cancer through epidermal growth
factor receptor transactivation. Lung Cancer (2007) 56(1):25–33. doi:10.1016/j.
lungcan.2006.11.014
60. Zhong L, Roybal J, Chaerkady R, Zhang W, Choi K, Alvarez CA, et al. Identification of secreted proteins that mediate cell-cell interactions in an in vitro
model of the lung cancer microenvironment. Cancer Res (2008) 68(17):7237–45.
doi:10.1158/0008-5472.CAN-08-1529
61. Elizur A, Adair-Kirk TL, Kelley DG, Griffin GL, deMello DE, Senior RM.
Tumor necrosis factor-alpha from macrophages enhances LPS-induced Clara
cell expression of keratinocyte-derived chemokine. Am J Respir Cell Mol Biol
(2008) 38(1):8–15. doi:10.1165/rcmb.2007-0203OC

November 2014 | Volume 5 | Article 587 | 126

Fritz et al.

62. Guth AM, Janssen WJ, Bosio CM, Crouch EC, Henson PM, Dow SW. Lung
environment determines unique phenotype of alveolar macrophages. Am J Physiol Lung Cell Mol Physiol (2009) 296(6):L936–46. doi:10.1152/ajplung.90625.
2008
63. Elizur A, Adair-Kirk TL, Kelley DG, Griffin GL, deMello DE, Senior RM.
Clara cells impact the pulmonary innate immune response to LPS. Am
J Physiol Lung Cell Mol Physiol (2007) 293(2):L383–92. doi:10.1152/ajplung.
00024.2007
64. Hagemann T, Robinson SC, Thompson RG, Charles K, Kulbe H, Balkwill
FR. Ovarian cancer cell-derived migration inhibitory factor enhances tumor
growth, progression, and angiogenesis. Mol Cancer Ther (2007) 6(7):1993–2002.
doi:10.1158/1535-7163.MCT-07-0118
65. Kimura YN, Watari K, Fotovati A, Hosoi F, Yasumoto K, Izumi H, et al.
Inflammatory stimuli from macrophages and cancer cells synergistically promote tumor growth and angiogenesis. Cancer Sci (2007) 98(12):2009–18.
doi:10.1111/j.1349-7006.2007.00633.x
66. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
67. Biswas SK, Gangi L, Paul S, Schioppa T, Saccani A, Sironi M, et al. A distinct and
unique transcriptional program expressed by tumor-associated macrophages
(defective NF-kappaB and enhanced IRF-3/STAT1 activation). Blood (2006)
107(5):2112–22. doi:10.1182/blood-2005-01-0428
68. Ramachandran P, Pellicoro A, Vernon MA, Boulter L, Aucott RL, Ali A, et al.
Differential Ly-6C expression identifies the recruited macrophage phenotype,
which orchestrates the regression of murine liver fibrosis. Proc Natl Acad Sci U
S A (2012) 109(46):E3186–95. doi:10.1073/pnas.1119964109
69. Redente EF, Keith RC, Janssen W, Henson PM, Ortiz LA, Downey GP, et al.
Tumor necrosis factor-alpha accelerates the resolution of established pulmonary
fibrosis in mice by targeting profibrotic lung macrophages. Am J Respir Cell Mol
Biol (2014) 50(4):825–37. doi:10.1165/rcmb.2013-0386OC
70. Shaul ME, Bennett G, Strissel KJ, Greenberg AS, Obin MS. Dynamic, M2like remodeling phenotypes of CD11c+ adipose tissue macrophages during

www.frontiersin.org

Macrophage depletion inhibits lung tumorigenesis

high-fat diet – induced obesity in mice. Diabetes (2010) 59(5):1171–81.
doi:10.2337/db09-1402
71. Kawakami S, Sato A, Nishikawa M, Yamashita F, Hashida M. Mannose receptormediated gene transfer into macrophages using novel mannosylated cationic
liposomes. Gene Ther (2000) 7(4):292–9. doi:10.1038/sj.gt.3301089
72. Korns D, Frasch SC, Fernandez-Boyanapalli R, Henson PM, Bratton DL. Modulation of macrophage efferocytosis in inflammation. Front Immunol (2011)
2:57. doi:10.3389/fimmu.2011.00057
Conflict of Interest Statement: The Guest Associate Editor Laurel L. Lenz declares
that, despite being affiliated to the same institution as the authors Jason M. Fritz,
Meredith A.Tennis, David J. Orlicky, Hao Yin, Cynthia Ju, Daniel T. Merrick, Alvin
M. Malkinson and Lori D. Dwyer-Nield, the review process was handled objectively
and no conflict of interest exists. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be
construed as a potential conflict of interest.
Received: 15 August 2014; accepted: 03 November 2014; published online: 25 November
2014.
Citation: Fritz JM, Tennis MA, Orlicky DJ, Yin H, Ju C, Redente EF, Choo KS, Staab
TA, Bouchard RJ, Merrick DT, Malkinson AM and Dwyer-Nield LD (2014) Depletion
of tumor-associated macrophages slows the growth of chemically induced mouse lung
adenocarcinomas. Front. Immunol. 5:587. doi: 10.3389/fimmu.2014.00587
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Fritz, Tennis, Orlicky, Yin, Ju, Redente, Choo, Staab, Bouchard,
Merrick, Malkinson and Dwyer-Nield. This is an open-access article distributed under
the terms of the Creative Commons Attribution License (CC BY). The use, distribution
or reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
accepted academic practice. No use, distribution or reproduction is permitted which
does not comply with these terms.

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REVIEW ARTICLE
published: 10 October 2014
doi: 10.3389/fimmu.2014.00500

Phenotypic diversity and emerging new tools to study
macrophage activation in bacterial infectious diseases
Mignane B. Ka 1 , Aurélie Daumas 1 , Julien Textoris 2 and Jean-Louis Mege 1 *
1

2

Unité de Recherche sur les Maladies Infectieuses Tropicales et Emergentes, UMR 63, CNRS 7278, IRD 198, INSERM U1095, Aix-Marseille Université, Marseille,
France
Unité Mixte bioMérieux-HCL, Hôpital Edouard Herriot, Lyon, France

Edited by:
Laurel L. Lenz, University of Colorado
School of Medicine, USA
Reviewed by:
Fulvio D’Acquisto, Queen Mary
University of London, UK
Kingston H. Mills, Trinity College
Dublin, Ireland
*Correspondence:
Jean-Louis Mege, Faculté de
Médecine, URMITE, 27 Bld. Jean
Moulin, Marseille 13005, France
e-mail: jean-louis.mege@univ-amu.fr

Macrophage polarization is a concept that has been useful to describe the different features
of macrophage activation related to specific functions. Macrophage polarization is responsible for a dichotomic approach (killing vs. repair) of the host response to bacteria; M1-type
conditions are protective, whereas M2-type conditions are associated with bacterial persistence. The use of the polarization concept to classify the features of macrophage activation
in infected patients using transcriptional and/or molecular data and to provide biomarkers for
diagnosis and prognosis has most often been unsuccessful. The confrontation of polarization with different clinical situations in which monocytes/macrophages encounter bacteria
obliged us to reappraise this concept. With the exception of M2-type infectious diseases,
such as leprosy and Whipple’s disease, most acute (sepsis) or chronic (Q fever, tuberculosis) infectious diseases do not exhibit polarized monocytes/macrophages. This is also the
case for commensals that shape the immune response and for probiotics that alter the
immune response independent of macrophage polarization. We propose that the type of
myeloid cells (monocytes vs. macrophages) and the kinetics of the immune response (early
vs. late responses) are critical variables for understanding macrophage activation in human
infectious diseases. Explorating the role of these new markers will provide important tools
to better understand complex macrophage physiology.
Keywords: macrophage, activation, polarization, infectious diseases, bacteria

INTRODUCTION
Why a new review about macrophage polarization during bacterial infectious diseases? The initial analysis of macrophage activation, based on in vitro experiments and the use of animal
models, suggested a dichotomic classification based on the production of canonical molecules associated with a specific function. The production of nitric oxide is associated with the killing
of microorganisms or tumor cells and characterizes M1-type
macrophage response whereas the expression of arginase (production of ornithine) is associated with the repair and characterizes
M2-type macrophage responses (1). The concept of M1/M2 polarization has been largely popularized because macrophage polarization was considered the reflection of Th1 and Th2 polarization of
lymphocytes, although the idea that activation by T cells is required
for macrophage polarization is likely incorrect (1). As the Th1/Th2
paradigm has progressively been replaced by several functional statuses over the past years, the meaning of a similar dichotomy of
macrophage activation is unknown. During the last years, numerous transcriptional and/or molecular markers associated with M1or M2-type macrophage responses were found but they did not
have a clear relationship with macrophage functions, which has
been a source of controversies. We feel that these new markers
could provide additional important tools to better understand
complex macrophage physiology. In addition, recent advances suggest that monocytes readily available in humans are not able to
polarize like mature tissue macrophages. As a consequence, the

Frontiers in Immunology | Inflammation

increasing number of publications in which clinical cohorts are
investigated with new tools of macrophage investigation allows
a global analysis of the cell responses, which results in a more
precise overview of the clinical data. It is likely that the concept of
M1/M2 macrophages is likely insufficient to describe human infectious diseases. While M2-type infectious diseases such as leprosy
and Whipple’s disease represent a clinical exception; most acute
(sepsis) or chronic (Q fever, tuberculosis) bacterial diseases do
not exhibit polarized monocytes/macrophages. According to the
analysis of Thomas Kuhn, the “paradigm” of macrophage polarization applied to human bacterial diseases suffers from abnormalities that could lead to a paradigm shift to a kinetic vision of
macrophage activation.

THE MACROPHAGE POLARIZATION CONCEPT
The molecular concept of the polarization of human macrophages
has been initially based on the selective expression of a few markers
that have poor specificity when expressed alone. The development
of high-throughput profiling technologies that enable the investigation of complex macrophage states (2) has increased the number
of biomarkers associated with the M1 or M2 status (Figure 1).
Among the papers reporting transcriptomic analysis of activated
macrophages that of Martinez et al. was the most contributive (3).
The authors showed that M1 and M2 polarization affect 5.2 and
0.3% of transcripts, respectively. The functional annotation reveals
the enrichment with categories such as DNA transcription, protein

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Macrophage polarization and infectious diseases

FIGURE 1 | M1 and M2 macrophage polarization. The figure represents canonical M1 and M2 agonists that induce the production of M1 and M2 markers by
human macrophages in vitro. These markers, isolated or combined, have been used to describe the polarization of monocytes and macrophages in clinical
investigations.

metabolism, G protein coupled-receptors, and lipid metabolism
in addition to well-identified cytokine and chemokine families.
Hence, the polarization of human macrophages has become more
complex than the initial descriptions.
A recent transcriptomic analysis of human macrophages stimulated by a large panel of agonists allowed a description of
macrophage activation as a spectrum. This spectrum of activation
was more complex than the M1 vs. M2 model of activation because
at least nine distinct activation programs were identified. The use
of network analyses demonstrated a central transcriptional regulator present in all activation conditions that was complemented by
regulators associated with the programs stimulated by each agonist
(4). The authors used this model of activation to analyze human
alveolar macrophages from patients who were smokers or from
patients with chronic obstructive pulmonary disease (COPD).
They found that the activation program of macrophages was
more complex than predicted in smokers and in patients with
COPD. They did not find enrichment with modules associated
with interleukin (IL)-4/IL-13 activation in patients with COPD,
as was expected, but did find a decrease in the modules associated with interferon (IFN)-γ (4). This report clearly demonstrates
that the prominent, popular point of view that cigarette smoke
and COPD increase M2-like characteristics (5) was not supported
when high-throughput approaches were used.
A proteomics approach has also been used to investigate
macrophage polarization. The MALDI-TOF mass spectrometry
(MS) technique combined with gel electrophoresis permitted the
identification of a large number of soluble or membrane proteins
in activated macrophages. This double approach allowed the identification of an M1 signature in human macrophages stimulated
with LPS and IFN-γ (6). Recently, we used MALDI-TOF MS to
characterize whole eukaryotic cells (7) and the activation status of
human macrophages (8). We found that whole-cell MALDI-TOF

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MS analysis was able to discriminate macrophages according to the
type of M1 or M2 agonists and allowed for the identification of
different subtypes of M1 or M2 macrophages. The MALDI-TOF
MS analysis of pathogen-stimulated macrophages also enabled the
detection of pathogen-associated fingerprints that did not correspond to the standard M1/M2 polarization model (8). Taken
together, the use of polarization markers other than iNOS and
arginase has been controversial. Recently, we proposed guidelines
for macrophage activation in which we favored an approach based
on a combination of markers instead of isolated canonical markers
of polarization (9).
The exploration of tissue macrophages, excepted alveolar
macrophages, requires biopsies in infected patients even if it
is possible to identify M1 and M2 macrophages in tissues
using proteomic or immunohistochemical approaches. Recently,
macrophage polarization was investigated in tissues from patients
with diseases characterized by a Th1 or Th2 response. M1
macrophages were defined as those expressing CD68 or CD163
with phosphorylated STAT1 (pSTAT1), and M2 macrophages were
defined on the basis of the co-expression of CMAF (macrophage
activation factor) with CD68 or CD163 (10). The pSTAT1 and
CMAF are preferentially associated with M1 and M2 macrophages,
respectively. In contrast, CD163, which was considered by several
authors as an M2 specific-marker (11), was unable to discriminate
M1 and M2 macrophages within pathological tissues. These findings were confirmed by a recent study in which macrophages were
differentiated by granulocyte macrophage-colony stimulating factor (GM-CSF) or macrophage-colony stimulating factor (M-CSF)
and secondarily polarized by IFN-γ or IL-4/IL-13; CD163 was
unable to discriminate the M1 status from the M2 status (12).
The investigation of macrophage activation in infected patients
concerns essentially circulating monocytes that are accessible after
blood collection and purification from blood, but the situation

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regarding their M1/M2 polarization is complex. Using a microarray approach, we showed that M1/M2 polarization, defined by
comparison with the IFN-γ and IL-4 signatures of macrophages,
was transient in human monocytes, and gene expression data
from published reports showed that not even small signatures of
polarized macrophages were found in monocytes (13). Hence, the
study of activation in tissue macrophages or circulating monocytes
suffers from the lack of convenient tools, suggesting that the concept of macrophage polarization is not convenient. Among the
recommendations for reporting macrophage activation, the recommendation precising how macrophages are isolated and which
marker combinations are used to measure macrophage activation
is likely a solution for the investigation of monocytes ex vivo (9).

MACROPHAGE POLARIZATION AND MICROBIOTA
The microorganisms present at the surfaces of mucosa mainly
consist of commensals that have developed mutualistic relationships with hosts such as human beings. Indeed, during steady-state
conditions, the microbiota influences the efficiency of digestion,
controls metabolism, and affects the differentiation and functions of intestinal immune cells, including macrophages. This
coevolution has been illustrated by numerous reports based on
studies on germ-free animals or antibiotic-treated hosts (14–16).
It has been established that the intestinal microbiota maintains
a tolerant environment that allows the development of M2-like
intestinal macrophages. Indeed, the macrophages from lamina
propria show down-regulated expression of innate response receptors and inflammatory functions, but they retain phagocytosis
and bactericidal activities (17). It is likely that commensals may
directly or indirectly shape the polarization status of intestinal
macrophages. Hence, Bacteroides fragilis and intestinal Clostridia
are known to stimulate regulatory T cells (Tregs) and polarization toward an M2 phenotype (14). The exopolysaccharide from
Bacillus subtilis prevents the intestinal disease associated with Citrobacter rodentium, and protection is transferred by peritoneal
macrophages (18). The probiotic Clostridium butyricum promotes
the development of IL-10-producing macrophages that prevent
inflammatory colitis (19). Some end-products of bacterial anaerobic fermentation, such as short-chain fatty acids (α-butyrate),
inhibit the inflammatory response of macrophages via a mechanism based on the inhibition of histone deacetylase (20). In
contrast, intestinal commensals such as Enterococcus faecalis polarize colon macrophages to an M1 phenotype in a murine model in
which macrophages are depleted with clodronate (21). These findings suggest that the diversity of commensal bacteria accounts for
the diversity of macrophage responses. Probiotics such as Lactobacillus sp. or Bifidobacterium sp. may benefit the host (14), but we
ignore their effect on macrophage polarization. The strain G-101
of Lactobacillus brevis inhibits the inflammatory response of mice
treated by trinitrobenzenesulfonic acid. This anti-inflammatory
property is related to the ability of the bacteria to prevent the
expression of M1 markers and to favor M2 markers, likely via
the production of IL-10 (22). For other authors, probiotics have
either no effect on the polarization of RAW 264.7 macrophages as
a readout (23), or these bacteria promote an activation profile of
the M1-like type in THP-1 cells stimulated with lipopolysaccharide (LPS) (24). It is noteworthy that all of these studies are limited

Frontiers in Immunology | Inflammation

Macrophage polarization and infectious diseases

to in vitro experiments or animal models, and the extrapolation to
human beings must be careful.
If the hypothesis that a breach of intestinal homeostasis is true,
the presence of pathogenic bacteria would interfere with the polarization status of intestinal and systemic macrophages. Hence, an
M1 profile would be found in patients with acute typhoid fever
due to Salmonella enterica serovar Typhi, whereas an M2 signature would be observed in convalescent patients. The M2 response
does not mean eradication of the pathogen because persistence of
the M2 status favors re-infection, relapses, and development of a
carrier state (25, 26). On the other hand, there is an increase in M1
and M2 markers in antrum from patients infected with Helicobacter pylori and uncomplicated gastritis. The presence of atrophic
gastritis is associated with the expression of M1 polarization. It
is predictable that shifting macrophage polarization from the M1
to M2 status is protective in chronic H. pylori infection. This may
be reminiscent of the association of high levels of CCL18, a typical M2 marker, with prolonged survival of patients with gastric
carcinoma (26, 27).
Imbalances in gut microbiota have also been associated with
systemic diseases such as allergy. Recently, Kim et al. reported the
induction of allergen-induced infiltration of inflammatory cells in
mice treated with antibiotics. This treatment alters macrophage
functions but reorients alveolar macrophages and circulating
monocytes toward an M2 phenotype. This latter response is
involved in allergic airway inflammation induced by allergens.
Antibiotic treatment facilitates fungal overgrowth that exacerbates
airway inflammation. The prostaglandin E2 produced by gut fungi
is responsible for eosinophil-mediated inflammation and M2
polarization of macrophages (28). If the concept of macrophage
polarization is useful for analyzing the host response to intestinal
pathogens, there is no clear evidence that it is a convenient tool to
measure the response to commensals and probiotics.

MACROPHAGE POLARIZATION AND ACUTE INFECTIOUS
DISEASES
As sepsis is a consequence of the systemic inflammatory response
to infectious aggression, it was tantalizing to consider sepsis as an
M1-associated disease (25). Sepsis can also associate a secondary
immunodeficiency in which the polarization of macrophages may
be altered, as in LPS tolerance. Indeed, LPS-tolerant macrophages
express M2 markers, but not M1 markers, and this phenotype can
be reversed by IFN-γ (29). It is thought that the evolution of sepsis is characterized by a transition from an initial M1 response to
a secondary M2 response. The interaction of macrophages with
pathogens accounts for their initial polarization, and the M1-toM2 transition should rather involve mechanisms of activation
control such as suppressors of cytokine signaling (SOCS) proteins; SOCS1 and SOCS2 are associated with M2 macrophages
whereas SOCS3 is overexpressed in M1 cells. A high SOCS1/SOCS3
expression ratio might be a biomarker of M2 cells in vivo (30).
The fact that M2 bias is associated with the resistance of mice
does not account for the poor prognosis of patients who exhibit
secondary immune deficiency with an M2 phenotype. Indeed,
this latter phase, named immune paralysis, is associated with
increased susceptibility to nosocomial infections and late lethality (31). In patients with sepsis, the percentage of monocytes

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expressing CD163 and CD206 is increased. The increase in monocytes expressing M2-like markers has been associated with a lower
proportion of IFN-γ-producing T cells or with a higher proportion of Tregs in patients with sepsis. Nevertheless, enrichment with
M2-type monocytes has no impact on sepsis prognosis (32). In
others reports, the expression of CD163 by monocytes is accurate
for discriminating patients with inflammatory presentation from
those with sepsis (33), suggesting that CD163 may be a biomarker
of prognosis and that the expression of CD163 by monocytes is
higher in non-survivors than in survivors (34). Soluble forms of
M2-type markers such as CD163 and CD206 are also increased
in patients with sepsis, and their high levels are associated with
poor prognosis in sepsis. Although membrane and soluble forms
of CD163 share the ability to be biomarkers of prognosis in sepsis, circulating CD163 reflecting the polarization of monocytes
or their activation independently of M1/M2 polarization tends
to be ignored (34, 35). The measurement of monocyte activation is a partial reflection of the altered immune functions in
tissues from patients with sepsis and does not assess the diversity of stimuli that they encounter from the initial pathological
event. It is probably more pertinent to consider the level of monocyte activation and not the bias toward a polarized status as a
biomarker.

INTERFERENCE WITH M1 POLARIZATION IN CHRONIC
INFECTIOUS DISEASES: Q FEVER
As intracellular bacteria subvert host microbicidal effectors
in vitro, we proposed that they have evolved specific strategies
to interfere with M1 polarization (25). The example of Q fever is
informative as we have assessed the concept of macrophage polarization in in vitro experiments, animal models, and patients. Q
fever is a zoonosis caused by Coxiella burnetii, an intracellular
bacterium related to Legionellae species, and for which the major
targets are monocytes and macrophages. The severity of the infectious disease is chronic evolution with a risk of endocarditis or
vascular infection (36).
The circulating monocytes exhibit a pro-inflammatory M1type response, which is consistent with epidemiological data showing bacterial clearance in most infected patients when they are
challenged by C. burnetii in vitro. More surprisingly, monocytederived macrophages are polarized toward an atypical M2-type in
response to bacterial stimulation. This latter effect is characterized by the release of IL-10, transforming growth factor (TGF)-β,
and CCL18 and the expression of the mannose receptor (MR) and
of arginase-1, but macrophages also express IL-6 and CXCL8, two
molecules that are associated with M1 polarization (37). These differences in monocyte/macrophage activation may account for the
unexplained differences in bacterial survival: C. burnetii are unable
to replicate in monocytes but replicate within macrophages (38).
Similar findings were found in vitro with Mycobacterium tuberculosis, which prevents M1 polarization and activates peroxisome
proliferator-activated receptor (PPAR)-γ, which is characteristic
of macrophage M2 polarization (25, 39).
Nevertheless, we identified IL-10 as the only cytokine able
to induce the replication of C. burnetii in monocytes and
macrophages, suggesting that IL-10-associated M2 polarization is
involved in bacterial replication and tissue persistence. The role of

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Macrophage polarization and infectious diseases

IL-10 in the pathogenesis of chronic infection is strengthened by
the correlation of the amount of IL-10 and the chronic evolution
of Q fever with the restoration of the microbicidal competence of
monocytes when IL-10 was neutralized (40, 41). The engulfment
of apoptotic cells by monocytes and macrophages is associated
with an M2 program induced by IL-10 and favors the intracellular replication of C. burnetii. In contrast, treatment of these M2
polarized myeloid cells with IFN-γ and the uptake of necrotic cells
suggest that the M1 program is sufficient to clear C. burnetii (42).
The role of IL-10 is demonstrated in transgenic mice that constitutively overexpress IL-10 in the macrophage compartment and
exhibit sustained infection, as in chronic Q fever. Macrophages
from IL-10-overexpressing mice are unable to clear C. burnetii
infection and exhibit an M2-type transcriptional program in
which arginase, MR and Yim1/2 are increased and inflammatory
markers are down-modulated (43). The infection of mice overexpressing IL-10, which mimics tuberculosis reactivation, reveals
features of M2 macrophages, as reported above in C. burnetii
infection of mice (26).
Concomitantly, we found that mice deficient for vanin-1, a
membrane-anchored pantetheinase that controls tissue inflammation, are permissive for C. burnetii and exhibit an activation program in macrophages that is skewed toward an IL-10associated M2 phenotype (44). Hence, IL-10-mediated polarization of macrophages is necessary for C. burnetii persistence in
tissues.
To test the relevance of these findings in patients, we selected
M1- and M2-related genes from the microarray analyses of IFN-γ
and IL-4-stimulated macrophages (Figure 2). The expression of
these genes was not different in patients with acute Q fever and
healthy controls. These findings did not support the hypothesis
that patients with acute Q fever, who are able to control the infection, should exhibit an M1-type phenotype. The expression of a
minority of M1/M2 genes was increased in patients with Q fever
endocarditis and who were unable to clear C. burnetii and who
were expected to exhibit an M2-type phenotype (13). The analysis
of the transcriptional profiles of patients with active tuberculosis shows the modulation of M1-related genes, but not that of
M2 genes. Similar results were obtained in infants vaccinated with
Calmette–Guerin bacillus (26, 45, 46).
In conclusion, the activation program of monocytes from
patients with acute and chronic Q fever and tuberculosis cannot be reduced to an M1/M2 dichotomy. We cannot rule out that
macrophages in tissues such as endocardium, lungs, or liver are
polarized, as suggested by in vitro studies and animal models. This
is illustrated by the example of pleural macrophages. Tuberculous
pleural effusion, an extra-pulmonary form of tuberculosis, is associated with the M1 profile in pleural fluid that is characterized by
an increase in M1 macrophages and inflammatory cytokines (47).

M2 POLARIZATION IN CHRONIC INFECTIOUS DISEASES:
LEPROSY AND WHIPPLE’S DISEASE
Two infectious diseases, leprosy and Whipple’s disease, which
share several features such as the tropism for macrophages of
Mycobacterium leprae and Tropheryma whipplei, and the role of
the immune response into features of pathogenesis, are associated
with M2 polarization (26). The overexpression of IL-10 is found in

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Macrophage polarization and infectious diseases

FIGURE 2 | Transcriptomic assessment of macrophage polarization. The figure represents the heat map of gene expression in IFN-γ- and IL-4-stimulated
macrophages. The use of microarray enables the identification of the original M1 and M2 signatures.

lepromatous lesions and likely reflects M2 polarization. The transcriptional analysis of these lesions reveals an enrichment of M2
genes, which is in contrast to what occurs in tuberculoid lesions
(48). The expression of CD163 by foamy macrophages in lepromatous lesions but not by macrophages from tuberculoid lesions
has been considered strong evidence of M2 polarization in lepromatous leprosy (26). Whether this polarization is a consequence
of the production of IL-10 or if it reflects a Th2 response is often
ignored.
Whipple’s disease is characterized by the presence of
macrophages with periodic acid-Schiff inclusions within the
lamina propria; these macrophages exhibit some features of
macrophages from mycobacterial lesions. As described above for
lepromatous leprosy, there is converging evidence that Th2 polarization of the immune response is critical for the pathophysiology

Frontiers in Immunology | Inflammation

of Whipple’s disease. An M2 macrophage signature was observed
in duodenal biopsies from one patient with intestinal Whipple’s
disease (49). Moos et al. reported the increased expression of
CD163 on duodenal macrophages and circulating monocytes,
and this finding was strengthened by an increase in IL-10 and
a decrease in inducible NO synthase expression in these cells, suggesting a functional polarization toward an M2 profile (50, 51).
The conclusion that IL-10 may be critical for T. whipplei pathogenicity was not confirmed by in vitro studies, in which we found
an increase in IL-1β, IL-16, and type I IFN production, but not
in IL-10 (52, 53). It is likely that type I IFN prevents the IFNγ-protective effect, as reported for mycobacterial infections (54).
This finding underlines the caution that must be taken regarding
conclusions about polarization when based on a limited number
of markers.

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Ka et al.

COMPLEXITY OF MACROPHAGE ACTIVATION IN INFECTIOUS
DISEASES
The analysis of infectious disease literature (see above) reveals
that modulation of monocyte/macrophage activation is frequently
observed, whereas clear-cut M1/M2 polarization is rather a rare
event. This observation is related to the history of infected patients.
Indeed, the stage of the disease is a critical parameter. For instance,
the activation of monocytes/macrophages is different in patients
with initial sepsis and those with delayed complications. In addition, numerous patients are distributed between two extreme
situations: between patients with acute Q fever and those with Q
fever endocarditis, there is a population of patients with valvular
disease and Q fever associated with a risk of chronic evolution, and
these patients overproduce IL-10 in a sustained manner. However,
the measurement of IL-10 at a given time of Q fever evolution
is not sufficient to assess the prognosis of patients with Q fever
(55). In patients with tuberculosis, the transcriptional signature
is transient at the beginning of the disease and is finished 1 year
later (45). Clearly, the analysis of the transcriptional pattern of
patients with tuberculosis will be dramatically different according to the time of the inclusion, and such an analysis is often
difficult to assess at the beginning of the disease. These different
clinical and experimental situations drove us to propose a model
of monocyte/macrophage activation in which the kinetic component of the disease was integrated. This model is based on the
comparison of the transcriptomes from activated monocytes and
macrophages. The responses of monocytes to polarizing ligands
are characterized by two early and late phases of monocyte activation. The hallmarks of the M1/M2 status are found in the early
phase but are absent from the late phase of activation. We selected
a series of early and late genes and measured their expression in
monocytes from patients with acute and chronic Q fever. Most
of the early genes were found to be up-regulated in monocytes
from patients with acute Q fever, two of them, NLRC5 and RTP4,
were up-regulated by IFN-γ, suggesting that IFN-γ plays a role in
the host response during acute Q fever. In contrast, the late genes
were up-regulated in chronic Q fever, and some early genes were
down-modulated. There was a specific association between late
genes such as ALOX15, CLEC4F, CCL13, and CCL23 and chronic
Q fever (13). It is noteworthy that some of them have been associated with the M2 program, which is a result that might lead to
incorrect conclusions about monocyte activation. We are unable
to assign a function to the modulated genes.
In conclusion, the analysis of macrophage polarization through
clinical situations revealed that the mechanisms underlying the
activation of monocytes and macrophages are distinct. This point
is critical because most clinical investigations are based on monocytes and the conclusions are extrapolated on data obtained with
macrophages. The second observation is the importance of activation kinetics in the assessment of infected patients who are at
different stages of disease history. Therefore, early and late genes
may be alternative biomarkers for analyzing infectious and inflammatory diseases. The lessons from the investigation of infected
patients do not invalidate the functional model of M1/M2 polarization. They revealed the difficulty to relate a signature and a
function. In addition, the finding of a role for these genes in
the activation of macrophages will be useful to understand the

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Macrophage polarization and infectious diseases

complexity of macrophage physiology in normal and pathological
conditions.

REFERENCES
1. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity.
J Innate Immun (2014). doi:10.1159/000364945
2. Kidd BA, Peters LA, Schadt EE, Dudley JT. Unifying immunology with informatics and multiscale biology. Nat Immunol (2014) 15(2):118–27. doi:10.1038/
ni0914-894c
3. Martinez FO, Gordon S, Locati M, Mantovani A. Transcriptional profiling of the
human monocyte-to-macrophage differentiation and polarization: new molecules and patterns of gene expression. J Immunol (2006) 177(10):7303–11.
doi:10.4049/jimmunol.177.10.7303
4. Xue J, Schmidt SV, Sander J, Draffehn A, Krebs W, Quester I, et al. Transcriptomebased network analysis reveals a spectrum model of human macrophage activation. Immunity (2014) 40(2):274–88. doi:10.1016/j.immuni.2014.01.006
5. Hussell T, Bell TJ. Alveolar macrophages: plasticity in a tissue-specific context.
Nat Rev Immunol (2014) 14(2):81–93. doi:10.1038/nri3600
6. Brown J, Wallet MA, Krastins B, Sarracino D, Goodenow MM. Proteome
bioprofiles distinguish between M1 priming and activation states in human
macrophages. J Leukoc Biol (2010) 87(4):655–62. doi:10.1189/jlb.0809570
7. Ouedraogo R, Flaudrops C, Ben Amara A, Capo C, Raoult D, Mege JL.
Global analysis of circulating immune cells by matrix-assisted laser desorption
ionization time-of-flight mass spectrometry. PLoS One (2010) 5(10):e13691.
doi:10.1371/journal.pone.0013691
8. Ouedraogo R, Daumas A, Ghigo E, Capo C, Mege JL, Textoris J.
Whole-cell MALDI-TOF MS: a new tool to assess the multifaceted activation
of macrophages. J Proteomics (2012) 75(18):5523–32. doi:10.1016/j.jprot.2012.
07.046
9. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
10. Barros MH, Hauck F, Dreyer JH, Kempkes B, Niedobitek G. Macrophage
polarisation: an immunohistochemical approach for identifying M1 and
M2 macrophages. PLoS One (2013) 8(11):e80908. doi:10.1371/journal.pone.
0080908
11. Buechler C, Ritter M, Orso E, Langmann T, Klucken J, Schmitz G. Regulation of
scavenger receptor CD163 expression in human monocytes and macrophages
by pro- and antiinflammatory stimuli. J Leukoc Biol (2000) 67(1):97–103.
doi:10.1189/jlb.0813437
12. Kittan NA, Allen RM, Dhaliwal A, Cavassani KA, Schaller M, Gallagher
KA, et al. Cytokine induced phenotypic and epigenetic signatures are key to
establishing specific macrophage phenotypes. PLoS One (2013) 8(10):e78045.
doi:10.1371/journal.pone.0078045
13. Mehraj V, Textoris J, Ben Amara A, Ghigo E, Raoult D, Capo C, et al. Monocyte responses in the context of Q fever: from a static polarized model to a
kinetic model of activation. J Infect Dis (2013) 208(6):942–51. doi:10.1093/
infdis/jit266
14. Ivanov II, Honda K. Intestinal commensal microbes as immune modulators.
Cell Host Microbe (2012) 12(4):496–508. doi:10.1016/j.chom.2012.09.009
15. Hooper LV, Macpherson AJ. Immune adaptations that maintain homeostasis with the intestinal microbiota. Nat Rev Immunol (2010) 10(3):159–69.
doi:10.1038/nri2710
16. Round JL, Mazmanian SK. The gut microbiota shapes intestinal immune
responses during health and disease. Nat Rev Immunol (2009) 9(5):313–23.
doi:10.1038/nri2515
17. Smith PD, Ochsenbauer-Jambor C, Smythies LE. Intestinal macrophages: unique
effector cells of the innate immune system. Immunol Rev (2005) 206:149–59.
doi:10.1111/j.0105-2896.2005.00288.x
18. Jones SE, Paynich ML, Kearns DB, Knight KL. Protection from intestinal inflammation by bacterial exopolysaccharides. J Immunol (2014) 192(10):4813–20.
doi:10.4049/jimmunol.1303369
19. Hayashi A, Sato T, Kamada N, Mikami Y, Matsuoka K, Hisamatsu T, et al.
A single strain of Clostridium butyricum induces intestinal IL-10-producing
macrophages to suppress acute experimental colitis in mice. Cell Host Microbe
(2013) 13(6):711–22. doi:10.1016/j.chom.2013.05.013
20. Chang PV, Hao L, Offermanns S, Medzhitov R. The microbial metabolite butyrate regulates intestinal macrophage function via histone deacetylase

October 2014 | Volume 5 | Article 500 | 133

Ka et al.

inhibition. Proc Natl Acad Sci U S A (2014) 111(6):2247–52. doi:10.1073/pnas.
1322269111
21. Yang Y, Wang X, Huycke T, Moore DR, Lightfoot SA, Huycke MM. Colon
macrophages polarized by commensal bacteria cause colitis and cancer
through the bystander effect. Transl Oncol (2013) 6(5):596–606. doi:10.1593/
tlo.13412
22. Jang SE, Hyam SR, Han MJ, Kim SY, Lee BG, Kim DH. Lactobacillus brevis G-101
ameliorates colitis in mice by inhibiting NF-κB, MAPK and AKT pathways and
by polarizing M1 macrophages to M2-like macrophages. J Appl Microbiol (2013)
115(3):888–96. doi:10.1111/jam.12273
23. Christoffersen TE, Hult LT, Kuczkowska K, Moe KM, Skeie S, Lea T, et al. In vitro
comparison of the effects of probiotic, commensal and pathogenic strains
on macrophage polarization. Probiotics Antimicrob Proteins (2014) 6(1):1–10.
doi:10.1007/s12602-013-9152-0
24. Habil N, Al-Murrani W, Beal J, Foey AD. Probiotic bacterial strains differentially modulate macrophage cytokine production in a strain-dependent and
cell subset-specific manner. Benef Microbes (2011) 2(4):283–93. doi:10.3920/
BM2011.0027
25. Benoit M, Desnues B, Mege JL. Macrophage polarization in bacterial infections.
J Immunol (2008) 181(6):3733–9. doi:10.4049/jimmunol.181.6.3733
26. Mege JL, Mehraj V, Capo C. Macrophage polarization and bacterial infections.
Curr Opin Infect Dis (2011) 24(3):230–4. doi:10.1097/QCO.0b013e328344b73e
27. Quiding-Jarbrink M, Raghavan S, Sundquist M. Enhanced M1 macrophage
polarization in human Helicobacter pylori-associated atrophic gastritis and in
vaccinated mice. PLoS One (2010) 5(11):e15018. doi:10.1371/journal.pone.
0015018
28. Kim YG, Udayanga KG, Totsuka N, Weinberg JB, Nunez G, Shibuya A. Gut
dysbiosis promotes M2 macrophage polarization and allergic airway inflammation via fungi-induced PGE2. Cell Host Microbe (2014) 15(1):95–102.
doi:10.1016/j.chom.2013.12.010
29. Chen J, Ivashkiv LB. IFN-γ abrogates endotoxin tolerance by facilitating Tolllike receptor-induced chromatin remodeling. Proc Natl Acad Sci U S A (2010)
107(45):19438–43. doi:10.1073/pnas.1007816107
30. Wilson HM. SOCS proteins in macrophage polarization and function. Front
Immunol (2014) 5:357. doi:10.3389/fimmu.2014.00357
31. Hotchkiss RS, Karl IE. The pathophysiology and treatment of sepsis. N Engl
J Med (2003) 348(2):138–50. doi:10.1056/NEJMra021333
32. Brunialti MK, Santos MC, Rigato O, Machado FR, Silva E, Salomao R. Increased
percentages of T helper cells producing IL-17 and monocytes expressing markers
of alternative activation in patients with sepsis. PLoS One (2012) 7(5):e37393.
doi:10.1371/journal.pone.0037393
33. Moller HJ, Moestrup SK, Weis N, Wejse C, Nielsen H, Pedersen SS, et al.
Macrophage serum markers in pneumococcal bacteremia: prediction of survival by soluble CD163. Crit Care Med (2006) 34(10):2561–6. doi:10.1097/01.
CCM.0000239120.32490.AB
34. Kjaergaard AG, Rodgaard-Hansen S, Dige A, Krog J, Moller HJ, Tonnesen
E. Monocyte expression and soluble levels of the haemoglobin receptor
(CD163/sCD163) and the mannose receptor (MR/sMR) in septic and critically
ill non-septic ICU patients. PLoS One (2014) 9(3):e92331. doi:10.1371/journal.
pone.0092331
35. Rodgaard-Hansen S, Rafique A, Christensen PA, Maniecki MB, Sandahl TD,
Nexo E, et al. A soluble form of the macrophage-related mannose receptor
(MR/CD206) is present in human serum and elevated in critical illness. Clin
Chem Lab Med (2014) 52(3):453–61. doi:10.1515/cclm-2013-0451
36. Raoult D, Marrie T, Mege JL. Natural history and pathophysiology of Q fever.
Lancet Infect Dis (2005) 5(4):219–26. doi:10.1016/S1473-3099(05)70052-9
37. Benoit M, Barbarat B, Bernard A, Olive D, Mege JL. Coxiella burnetii, the agent of
Q fever, stimulates an atypical M2 activation program in human macrophages.
Eur J Immunol (2008) 38(4):1065–70. doi:10.1002/eji.200738067
38. Ben Amara A, Bechah Y, Mege JL. Immune response and Coxiella burnetii invasion. Adv Exp Med Biol (2012) 984:287–98. doi:10.1007/978-94-007-4315-1_15
39. Rajaram MV, Brooks MN, Morris JD, Torrelles JB, Azad AK, Schlesinger
LS. Mycobacterium tuberculosis activates human macrophage peroxisome
proliferator-activated receptor gamma linking mannose receptor recognition
to regulation of immune responses. J Immunol (2010) 185(2):929–42. doi:10.
4049/jimmunol.1000866
40. Ghigo E, Capo C, Raoult D, Mege JL. Interleukin-10 stimulates Coxiella burnetii replication in human monocytes through tumor necrosis factor downmodulation: role in microbicidal defect of Q fever. Infect Immun (2001)
69(4):2345–52. doi:10.1128/IAI.69.4.2345-2352.2001

Frontiers in Immunology | Inflammation

Macrophage polarization and infectious diseases

41. Ghigo E, Honstettre A, Capo C, Gorvel JP, Raoult D, Mege JL. Link between
impaired maturation of phagosomes and defective Coxiella burnetii killing in
patients with chronic Q fever. J Infect Dis (2004) 190(10):1767–72. doi:10.1086/
425041
42. Benoit M, Ghigo E, Capo C, Raoult D, Mege JL. The uptake of apoptotic cells
drives Coxiella burnetii replication and macrophage polarization: a model for
Q fever endocarditis. PLoS Pathog (2008) 4(5):e1000066. doi:10.1371/journal.
ppat.1000066
43. Meghari S, Bechah Y, Capo C, Lepidi H, Raoult D, Murray PJ, et al. Persistent
Coxiella burnetii infection in mice overexpressing IL-10: an efficient model for
chronic Q fever pathogenesis. PLoS Pathog (2008) 4(2):e23. doi:10.1371/journal.
ppat.0040023
44. Meghari S, Berruyer C, Lepidi H, Galland F, Naquet P, Mege JL. Vanin-1 controls granuloma formation and macrophage polarization in Coxiella burnetii
infection. Eur J Immunol (2007) 37(1):24–32. doi:10.1002/eji.200636054
45. Berry MP, Graham CM, McNab FW, Xu Z, Bloch SA, Oni T, et al. An interferoninducible neutrophil-driven blood transcriptional signature in human tuberculosis. Nature (2010) 466(7309):973–7. doi:10.1038/nature09247
46. Fletcher HA, Keyser A, Bowmaker M, Sayles PC, Kaplan G, Hussey G, et al.
Transcriptional profiling of mycobacterial antigen-induced responses in infants
vaccinated with BCG at birth. BMC Med Genomics (2009) 2:10. doi:10.1186/
1755-8794-2-10
47. Tang Y, Hua SC, Qin GX, Xu LJ, Jiang YF. Different subsets of macrophages
in patients with new onset tuberculous pleural effusion. PLoS One (2014)
9(2):e88343. doi:10.1371/journal.pone.0088343
48. Montoya D, Cruz D, Teles RM, Lee DJ, Ochoa MT, Krutzik SR, et al. Divergence
of macrophage phagocytic and antimicrobial programs in leprosy. Cell Host
Microbe (2009) 6(4):343–53. doi:10.1016/j.chom.2009.09.002
49. Desnues B, Lepidi H, Raoult D, Mege JL. Whipple disease: intestinal infiltrating
cells exhibit a transcriptional pattern of M2/alternatively activated macrophages.
J Infect Dis (2005) 192(9):1642–6. doi:10.1086/491745
50. Moos V, Schmidt C, Geelhaar A, Kunkel D, Allers K, Schinnerling K, et al.
Impaired immune functions of monocytes and macrophages in Whipple’s disease. Gastroenterology (2010) 138(1):210–20. doi:10.1053/j.gastro.2009.07.066
51. Geelhaar-Karsch A, Schinnerling K, Conrad K, Friebel J, Allers K, Schneider T,
et al. Evaluation of arginine metabolism for the analysis of M1/M2 macrophage
activation in human clinical specimens. Inflamm Res (2013) 62(9):865–9.
doi:10.1007/s00011-013-0642-z
52. Al Moussawi K, Ghigo E, Kalinke U, Alexopoulou L, Mege JL, Desnues B.
Type I interferon induction is detrimental during infection with the Whipple’s
disease bacterium, Tropheryma whipplei. PLoS Pathog (2010) 6(1):e1000722.
doi:10.1371/journal.ppat.1000722
53. Desnues B, Raoult D, Mege JL. IL-16 is critical for Tropheryma whipplei replication in Whipple’s disease. J Immunol (2005) 175(7):4575–82. doi:10.4049/
jimmunol.175.7.4575
54. Rayamajhi M, Humann J, Kearney S, Hill KK, Lenz LL. Antagonistic crosstalk
between type I and II interferons and increased host susceptibility to bacterial
infections. Virulence (2010) 1(5):418–22. doi:10.4161/viru.1.5.12787
55. Honstettre A, Imbert G, Ghigo E, Gouriet F, Capo C, Raoult D, et al. Dysregulation of cytokines in acute Q fever: role of interleukin-10 and tumor necrosis factor in chronic evolution of Q fever. J Infect Dis (2003) 187(6):956–62.
doi:10.1086/368129
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 24 July 2014; accepted: 25 September 2014; published online: 10 October
2014.
Citation: Ka MB, Daumas A, Textoris J and Mege J-L (2014) Phenotypic diversity
and emerging new tools to study macrophage activation in bacterial infectious diseases.
Front. Immunol. 5:500. doi: 10.3389/fimmu.2014.00500
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Ka, Daumas, Textoris and Mege. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The
use, distribution or reproduction in other forums is permitted, provided the original
author(s) or licensor are credited and that the original publication in this journal is cited,
in accordance with accepted academic practice. No use, distribution or reproduction is
permitted which does not comply with these terms.

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REVIEW ARTICLE
published: 08 September 2014
doi: 10.3389/fimmu.2014.00428

The role of myeloid cell activation and arginine metabolism
in the pathogenesis of virus-induced diseases
Kristina S. Burrack and Thomas E. Morrison*
Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Masaaki Murakami, Hokkaido
University, Japan
Mayda Gursel, Middle East Technical
University, Turkey
*Correspondence:
Thomas E. Morrison, Department of
Immunology and Microbiology,
University of Colorado School of
Medicine, 12800 East 19th Avenue,
Mail Stop 8333, Aurora, CO 80045,
USA
e-mail: thomas.morrison@
ucdenver.edu

When an antiviral immune response is generated, a balance must be reached between two
opposing pathways: the production of proinflammatory and cytotoxic effectors that drive
a robust antiviral immune response to control the infection and regulators that function
to limit or blunt an excessive immune response to minimize immune-mediated pathology
and repair tissue damage. Myeloid cells, including monocytes and macrophages, play an
important role in this balance, particularly through the activities of the arginine-hydrolyzing
enzymes nitric oxide synthase 2 (Nos2; iNOS) and arginase 1 (Arg1). Nitric oxide (NO)
production by iNOS is an important proinflammatory mediator, whereas Arg1-expressing
macrophages contribute to the resolution of inflammation and wound repair. In the context
of viral infections, expression of these enzymes can result in a variety of outcomes for the
host. NO has direct antiviral properties against some viruses, whereas during other virus
infections NO can mediate immunopathology and/or inhibit the antiviral immune response
to promote chronic infection. Arg1 activity not only has important wound healing functions but can also inhibit the antiviral immune response during some viral infections. Thus,
depending on the specific virus and the tissue(s) involved, the activity of both of these
arginine-hydrolyzing enzymes can either exacerbate or limit the severity of virus-induced
disease. In this review, we will discuss a variety of viral infections, including HIV, SARSCoV, LCMV, HCV, RSV, and others, where myeloid cells influence the control and clearance
of the virus from the host, as well as the severity and resolution of tissue damage, via
the activities of iNOS and/or Arg1. Clearly, monocyte/macrophage activation and arginine
metabolism will continue to be important areas of investigation in the context of viral
infections.
Keywords: arginase, iNOS, viral pathogenicity, macrophages, immunity, cellular

INTRODUCTION
Tissue -resident and monocyte-derived macrophages are innate
immune cells that play a key role in normal tissue homeostasis, presentation of foreign and self antigens following infection or injury, pathogen clearance, and resolution of inflammation and wound healing. Depending on the microenvironment,
macrophages can be programed to adopt a variety of proinflammatory, regulatory, resolving, and immunosuppressive activation phenotypes, particularly in vivo. These activation states
exist as a complex continuum of overlapping phenotypes; however, macrophage subsets with distinct functions have been
defined (1). Macrophages are considered M1-polarized when
stimulated by IFN-γ or Toll-like receptor (TLR) ligands, such
as lipopolysaccharide (LPS), to express inducible nitric oxide
synthase (iNOS; Nos2) and produce nitric oxide (NO). NOS
enzymes metabolize l-arginine to citrulline and NO. NO is a
short-lived gaseous messenger with physiological and pathological effects. Nanomolar concentrations of NO, generated by
endothelial NOS and neuronal NOS, are important for maintaining homeostasis, regulating vasodilation, and for the aggregation,
recruitment, and adhesion of platelets to the vascular endothelium. iNOS generates micromolar levels of NO that modulates

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various pathophysiological processes and is important for killing
intracellular pathogens (2).
In contrast, M2-polarized macrophages result following stimulation of cells with a variety of stimuli, including type 2 cytokines
such as interleukin (IL)-4 or IL-13. M2-polarized macrophages
express a distinct l-arginine-metabolizing enzyme, arginase 1
(Arg1), which hydrolyzes l-arginine to l-ornithine and urea. lOrnithine can be further metabolized to polyamines, which participate in a variety of fundamental cellular functions (e.g., proliferation, cell membrane transport), and l-proline, which is an essential
component of collagen. In addition to playing important roles
in defense against extracellular parasites and tissue repair, Arg1
expression and activity in myeloid cells have emerged as a major
regulator of innate and adaptive immune responses (3). Other
M2-like suppressive or anti-inflammatory macrophages include
myeloid-derived suppressor cells (MDSCs) and tumor-associated
macrophages (TAMs). MDSCs are considered to be an immature population of myeloid cells, including both monocyte-like
(GR-1/Ly-6C+ ) and neutrophil-like (GR-1/Ly-6G+ ) populations,
associated with tumors or infections that suppress proinflammatory responses (4, 5). Depending on the context, MDSCs have
been shown to mediate their suppressive activity via NO- and/or

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Burrack and Morrison

Arg1-dependent mechanisms. Importantly, macrophages are not
permanently programed, but are considered “plastic” – that is,
macrophages have been shown to change activation phenotypes
depending on the local environment.
Although the M1/iNOS and M2/Arg1 division is generally
appropriate, Arg1 can be induced in M1-like macrophages under
certain conditions. Thus, due to the spectrum of activation states
for macrophages, a framework for macrophage-activation nomenclature was recently suggested (6). In an attempt to avoid confusion
in this review, we focused on the specific effects of the l-arginine
metabolizing enzymes iNOS or Arg1 on the pathogenesis of viral
infections, noting other activation markers where appropriate.
Increasing evidence suggests that myeloid cell programing,
iNOS, and Arg1 contribute to the pathogenesis of numerous virus
infections, suggesting that therapies that target these cells and
pathways may be beneficial for the treatment of some virus diseases. In this review, we highlight recent studies of viral infections
where myeloid cell polarization – resulting in expression of iNOS
or Arg1 – contribute to viral control or the development of chronic
virus infection and mediate the resolution of tissue damage or
cause immunopathology.

NO PRODUCTION CAN BE BENEFICIAL DURING VIRUS
INFECTION
NO has antimicrobial activity against a number of bacteria, parasites, and fungi (7, 8). Additionally, NO has been shown to have
direct antiviral effects in vitro and/or in vivo against several viruses,
including DNA viruses such as herpes simplex virus type-1 (HSV1), ectromelia virus (EV), and vaccinia virus (VV) (9, 10), as well
as some RNA viruses such as vesicular stomatitis virus (VSV) (11),
Japanese encephalitis virus (JEV) (12), dengue virus (DENV) (13),
and coxsackievirus (Table 1) (14–17). There are several advantages
of using NO as an antiviral agent. For instance, unlike complement
and antibody, NO can readily pass through cellular membranes
into neighboring cells as well as some viruses. Additionally, NO is
likely to act on a variety of both viral and virally exploited cellular
targets, inhibiting viral replication as well as limiting the capacity
of viruses to develop resistance. Lastly, the effect of NO is independent of immune recognition of the infected cell, in contrast to
that of antiviral lymphocytes, which could be important in virusinfected cells where expression of MHC class I molecules may be
downregulated and in some virally infected tissues such as the
brain where expression of MHC class I and II molecules is limited.
In initial studies in vitro, inhibition of EV, VV, and HSV-1 replication in mouse RAW 264.7 macrophages and in primary mouse
macrophages following IFN-γ treatment was shown to be largely
dependent on NO production (9, 10). Additionally, pharmacologic inhibition of NOS or genetic deletion of Nos2 resulted in
increased viral titers and mortality following EV infection in mice
(9, 18). Moreover, NO affects several events in the late stages of
the life cycle of VV, including viral DNA replication, viral protein
synthesis, and virion maturation in vitro (32). These studies provided some of the first evidence that macrophage-produced NO
has direct antiviral effects.
In addition to inhibiting HSV-1 replication in vitro,
macrophage-derived NO has been shown to have anti-HSV properties in vivo. In a mouse model of HSV-1-mediated corneal

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Arginine metabolism in virus pathogenesis

disease, iNOS was highly induced in the trigeminal ganglion (TG)
of HSV-1-infected mice, and its expression was markedly reduced
in mice depleted of macrophages (22). Depletion of macrophages
prior to HSV-1 infection resulted in markedly reduced iNOS
expression and higher viral loads in the TG of infected mice
(22, 23), suggesting that macrophages were the main source of
iNOS expression in the affected tissues following HSV-1 infection and that NO had important anti-HSV-1 properties in vivo.
Consistent with these data, inhibition of NOS activity resulted in
increased viral loads in the TG (22). Additional studies showed
that F4/80+ GR-1+ inflammatory monocytes were recruited to
the eye via an IFN-α-driven CCL2 gradient and restricted HSV-1
replication in that tissue via NO production (24). It was further
shown that NO production by F4/80+ macrophages in the brains
of HSV-1-infected mice blocked viral replication in a partially
TLR2- and TLR9-dependent mechanism (25). Finally, following
footpad inoculation, HSV-1-infected Nos2 −/− mice displayed a
delayed clearance of virus from the dorsal root ganglia (DRG)
and exhibited an increase in the frequency of virus reactivation
in DRG (26).
The reactivity of NO and its higher oxides and nitrosothiol
products (84) makes it likely that a variety of molecular targets are involved in its antiviral action. It has been shown that
NO can inhibit ribonucleotide reductase (85, 86), a rate-limiting
enzyme in DNA synthesis, and NO can lead to the deamination of mammalian and bacterial DNA (87, 88), which may be
important antiviral mechanisms. Indeed, HSV-1 encodes its own
ribonucleotide reductase and although it is not required for HSV1 replication in vitro, it is necessary under conditions where the
intracellular pool of deoxynucleotides is limited (89, 90). Thus, by
inactivating this cellular and/or viral enzyme, NO may halt virus
replication by directly inhibiting viral DNA synthesis.
In addition to HSV-1, treatment of primary human cells with
an NO donor following infection with human cytomegalovirus
(HCMV), a beta-herpesvirus, resulted in a significant reduction
of early and late viral protein expression (28). Consistent with
these in vitro data, Nos2 −/− mice (129/Sv/Ev x C57BL/6 F1) exhibited increased viral titers and mortality following infection with
murine CMV (MCMV; Smith VR194 strain) (29).
Nitric oxide has also been shown to have antiviral properties
on a chicken herpesvirus, Marek’s disease virus (MDV), which can
cause T cell lymphomas in chickens: Addition of NO-generating
compounds inhibited viral replication in chicken fibroblasts (33).
Additionally, the treatment of chickens with an inhibitor of iNOS
increased the level of MDV replication in vivo (34). Further studies demonstrated that NO production was limited to chickens that
were genetically resistant to tumor development following MDV
infection or to chickens that were vaccinated before being inoculated with MDV (35). Thus, NO appeared to be produced in both
types of resistance to tumor development in Marek’s disease, either
acquired after vaccination or genetic. Together, these findings suggest a role of NO in the protective immune mechanisms against
Marek’s disease, possibly through its activity on viral replication.
Finally, studies with HBV, a hepadnavirus associated with acute
and chronic hepatitis, demonstrated that HBV replicated to higher
levels in the livers of HBV-transgenic Nos2 −/− mice than control transgenic mice, and transgenic Nos2 −/− mice had increased

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Arginine metabolism in virus pathogenesis

Table 1 | Differential roles for the L-arginine-hydrolyzing enzymes iNOS and Argl in virus-induced diseases.

Beneficial

Antiviral

iNOS/NO

Reference

Ectromelia virus (EV)

(9, 10, 18)

for host

Tissue repair/

Arg1

Reference

Respiratory syncytial virus (RSV)

(19–21)

Influenza

(27)

regeneration
Herpes simplex virus-1 (HSV-1)
Cytomegalovirus

(CMV)a

(9, 10, 22–26)
(28, 29)

Vaccinia virus (VV)

(9, 10, 32)

Marek’s disease virus (MDV)b

(33–35)

Hepatitis B virus (HBV)

(36)

Vesicular stomatitis virus (VSV)

(11)

Japanese encephalitis virus (JEV)

(12)

Dengue virus (DENV)

(13, 37–41)

West Nile virus (WNV)

(42)

Sindbis virus (SINV)

(43)

Reovirus (T3A strain)

(44)

Coxsackievirus B3 and B4 (CVB3,

(14–17, 91)

Coxsackievirus B3

(CVB3)d

(30, 31)

CVB4)
Detrimental Immunopato host

Influenza

(45–55)

thologic

Immunopa-

SARS-CoV

(56, 57)

Murine γ-herpesvirus-68

(61–63)

thologic
HSV-1

(58–60)

(MHV-68)e

Promotes viral

Feline immunodeficiency virus

(64, 65)

Cytomegalovirus (CMV)c

(66)

persistence

Promotes viral

LCMV clone 13

(67)

Ross River virus (RRV),

(69)

persistence
rJ2.2 strain of mouse hepatitis virus

(68)

(neurotropic coronavirus)

Lymphocytic choriomeningitis virus

chikungunya virus (CHIKV)

(70)

(LCMV) clone 13

Marek’s disease virus (MDV)f

(35)

Hepatitis C virus (HCV)

(71)

Human immunodeficiency virus

(72–77)

(HIV)

a

(78–80)

Influenza

(81, 82)

Human papillomavirus (HPV)g

(83)

HCMV, in vitro; MCMV, in 129/Sv/Ev x C57BL/6 F1 mice.

b

For resistant chickens.

c

MCMV, in BALB/c mice.

d

In female BALB/c mice.

e

In Ifngr −/− mice.

f

Hepatitis B virus (HBV)

For susceptible chickens.

g

Mouse model of HPV-induced cancer.

liver disease (36). It was further demonstrated that NO production by mononuclear cells, most likely macrophages, in the liver
mediated most of the antiviral activity resulting from IFN-γ production by virus-specific T cells (36), suggesting an antiviral role
for macrophage-derived NO following HBV infection in mice.
In addition to DNA viruses, macrophage-derived NO also
exerts antiviral effects against a number of RNA viruses. Inhibition
of JEV, a mosquito-transmitted flavivirus that causes encephalitis
in humans, in IFN-γ-activated RAW 264.7 macrophages in vitro

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correlated with NO production, and IFN-γ-activated RAW 264.7
macrophage-mediated inhibition of JEV replication in murine
neuroblastoma N18 cells was NO-dependent (12). Moreover, inhibition of NOS activity led to increased mortality in JEV-infected
mice (12).
In terms of its mechanism of action, NO was found to inhibit
JEV RNA synthesis, viral protein accumulation, and virus release
from infected cells in vitro (12). These data suggest that NO may
be directly or indirectly inhibiting viral enzymes and/or other

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Burrack and Morrison

cellular components required for viral replication, and this may
subsequently block viral protein synthesis. Additionally, NO may
interfere with the release and/or maturation of virions.
Monocyte/macrophage-derived NO may also block replication of DENV, another mosquito-transmitted flavivirus. Infection
with DENV resulted in increased levels of NO in patients with
dengue fever, the classic form of the disease (37). Additionally,
iNOS expression was induced in CD14+ monocytes from a subset of acutely infected individuals (13). It was further shown that
ex vivo infection of human monocytes with DENV-1 resulted in
increased iNOS expression, and inhibition of iNOS activity led to
increased DENV antigen detection in these cells (13). Moreover,
treatment of C6/36 mosquito cells with an NO donor resulted in
reduced DENV-positive cells (13). These data suggest that DENV
replication is susceptible to NO-mediated inhibition. Consistent
with this, Nos2 −/− mice were shown to be more susceptible to
DENV infection, resulting in more severe disease and increased
lethality in mouse models of DENV-2 and DENV-3 infection (38,
39). It was further demonstrated that, following DENV infection
in vivo, IL-12 and IL-18 induced IFN-γ production, resulting in
iNOS expression and NO production, which contributed to viral
control (38, 39).
In addition to monocyte/macrophage-derived NO, a recent
study demonstrated that platelets isolated from patients with
dengue fever had increased l-arginine transport and increased
NO production compared to platelets from healthy controls
(40). However, NO has anti-aggregatory properties, and MendesRibeiro et al. (40) found that dengue patients exhibited decreased
collagen-induced platelet aggregation, consistent with the vascular leak and hemorrhagic manifestations of dengue hemorrhagic
fever/dengue shock syndrome (DHF/DSS), thus establishing an
association between reduced platelet aggregation, enhancement
of the l-arginine–NO pathway, and DHF/DSS (41).
In contrast, Getts and colleagues showed that experimentally
abrogating NO activity during West Nile virus (WNV) encephalitis, a related flavivirus, in NO-competent mice at a specific, relatively late time point prolonged survival of infected mice, while
pharmacological inactivation throughout disease did not (42).
Combined, these data suggest that although during DENV infection IFN-γ-induced NO production has a role in antiviral defense,
it is likely that dysregulation of the IL-12/18–IFN-γ–NO axis
leads to immune-mediated damage in certain flavivirus infections.
Along these lines, it has also been shown that treatment of mice
with a NOS inhibitor increased mortality rates following Sindbis
virus (SINV) infection (43), suggesting a protective role for NO
during this particular CNS infection. However, SINV replication
in the brain was unaffected. Furthermore, treatment of neuroblastoma cells with NO donors had little effect on SINV replication but
increased cell viability (43). These data suggest that NO protects
mice from fatal SINV-induced encephalitis by a distinct mechanism that does not directly involve the inhibition of virus growth
but rather may enhance survival of the infected neuron until the
immune response can control virus replication.
Nitric oxide also plays an antiviral role during CNS infection
with reovirus. Infection of neonatal mice with the prototypic neurotropic reovirus strain (T3A) induced iNOS expression in brain
areas demonstrating reovirus antigen expression and associated

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Arginine metabolism in virus pathogenesis

virus-induced injury (44). Reovirus also induced iNOS expression
following in vitro infection of primary neuronal and glial cultures.
Reovirus was shown to infect a subpopulation of microglial cells
in vitro (44), suggesting that direct virus interaction may induce
iNOS in this specialized population of macrophages. Treatment
of neuronal cultures with an NO donor inhibited viral replication
whereas a NOS inhibitor increased viral growth (44), suggesting
iNOS has the potential to exert antiviral activity in vivo.
Finally, coxsackievirus infection has been shown to induce
expression of iNOS in macrophages infiltrating the hearts
of infected mice (17). Treatment of WT mice with a NOS
inhibitor and infection of Nos2 −/− mice resulted in more severe
coxsackievirus-induced pancreatitis and myocarditis, elevated
viral loads in tissues, and decreased survival compared to WT
mice following coxsackievirus B3 (CVB3) infection (14, 15, 17).
Similarly, Nos2 −/− mice infected with coxsackievirus B4 exhibited decreased survival and delayed viral clearance compared to
WT mice (16). These data suggest an antiviral effect of NO against
coxsackievirus infection. Consistent with this, it was demonstrated
that NO inhibits the 2A and 3C proteinases of CVB3 in vitro (91).
Additionally, CVB3-infected outbred mice showed significantly
reduced signs of myocarditis after treatment with NO donors (91).

NO PRODUCTION CAN BE DETRIMENTAL TO HOST
MYELOID CELL PRODUCTION OF NO CAN BE IMMUNOPATHOLOGIC
DURING VIRUS INFECTIONS

Despite its protective capacity during some viral infections, NO
can also contribute to immunopathology. The pathological effects
of NO are likely due, at least in part, to oxidative damage caused by
the interaction of NO

with oxygen radicals such as the superoxide
anion radical O−
2 and hydrogen peroxide (H2 O2 ).
For example, although addition of an NO donor to virusinfected MDCK cells reduced influenza A and B viral burden
in vitro (45), treatment of mice with inhaled NO (iNO) did not
decrease the viral load of influenza A (mouse-adapted H1N1
strain)-infected mice; in fact, prophylactic treatment with iNO
resulted in enhanced weight loss and decreased survival following
infection (46), suggesting a pathogenic role for NO. Consistent
with this, chickens, which show a high level of mortality and associated pathology following avian influenza infection, had higher
levels of iNOS expression in the lungs compared with H5N1
influenza-infected ducks, which show relatively minor symptoms
following influenza infection (47). Additionally, Akaike and colleagues (48) found evidence of the production of peroxynitrite,
which is generated through the reaction of NO and O2 – , in the
lungs of influenza A (mouse-adapted H2N2 strain)-infected mice.
Moreover, inhibition of NOS resulted in enhanced survival and
decreased pneumonia, but not decreased viral loads, in influenzainfected mice (48, 49), suggesting that NO was contributing to
pathogenesis rather than having direct antiviral effects. Nos2 −/−
mice also survived a lethal dose of influenza A virus (PR/8/34
strain) infection with little histopathologic evidence of pneumonitis; however, in these studies no infectious virus was detected in
Nos2 −/− mice at day 6 after infection (49). The enhanced viral
control in Nos2 −/− mice was shown to require the activity of
IFN-γ (51), with Nos2 −/− mice also producing increased virusspecific IgG2a antibody titers (50). Additionally, genetic deletion

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Burrack and Morrison

of Nos2 or pharmacologic inhibition of NOS enhanced survival of
mice inoculated with the highly pathogenic (non-mouse-adapted)
1918 influenza virus strain, although mice exhibited similar viral
loads to control mice in lung tissue at the peak of viral replication (51). Influenza infection in vitro was shown to induce
apoptosis, and a reduction in influenza-mediated apoptosis was
noted in cells treated with a NOS inhibitor (52). Similarly, fewer
apoptotic cells were found in the lungs of influenza-infected
Nos2 −/− mice, suggesting that NO mediates cell death following
influenza infection (52). The cellular source of iNOS/NO following influenza infection in mice was shown to be CCR2+ inflammatory monocytes that accumulate in the lungs: CCR2−/− mice
survived a lethal challenge of influenza infection (PR/8/34 strain)
and had significantly reduced accumulation of iNOS-expressing
macrophages in the lung, with no associated increase in viral titers
or dissemination (53).
It was also recently shown that a subset of monocyte-derived
dendritic cells (DCs), described as TNF-α/iNOS-producing DCs
(tipDCs), accumulate in greater numbers during the course of
lethal versus sublethal influenza infections, suggesting a pathogenic role for this subpopulation of myeloid cells (54). Interestingly, though, Aldridge et al. (54) found that the tipDCs also
stimulated a local, protective CD8+ T cell response in the virusinfected respiratory tract, indicating both protective and pathogenic roles for these cells in influenza infection. It was further
shown that partially compromising tipDC recruitment via treatment with pioglitazone, a synthetic agonist of the peroxisome
proliferator-activated receptor-γ (PPAR-γ), was protective against
lethal influenza challenge (54). Pioglitazone treatment led to a
reduction in the levels of CCL2 (MCP-1) and MCP-3 in the BAL
fluid of influenza-infected mice (54). Pioglitazone has also been
shown to reduce the production of a wide range of proinflammatory molecules, including iNOS (55), providing further evidence
for the importance of NO production by monocyte-derived cells
in the pathogenesis of influenza infection.
Pharmacologic inhibition of NOS using l-NMMA also
decreased pneumonitis and increased survival following intranasal
infection of CBA/J mice with HSV-1, despite a 17-fold increase in
viral titers in the lung at day 3 after inoculation (58). In contrast, treatment of BALB/c mice with a different NOS inhibitor
[aminoguanidine (AG), administered intranasally] resulted in
enhanced pneumonitis, viral titers, and mortality following infection with a different strain of HSV-1 (59). Thus, the precise role
of NO in HSV-1 pneumonitis remains to be determined. NO and
other ROS/RNS were also shown to be pathogenic in the brains
of mice with herpes encephalitis: iNOS was induced in CD11b+
resident microglia following intranasal infection with HSV-1, and
oxidative and nitrative damage was found in the brains of infected
animals (60).
A common neurological complication of HIV infection in
the developed world is sensory neuronal injury accompanied by
inflammation, which is clinically manifested as disabling pain and
gait instability. Feline immunodeficiency virus (FIV) infection
of cats, which causes similar neuroinflammation together with
immunosuppression in cats, resulted in induction of iNOS and
STAT-1, which were predominantly produced by macrophages, in
DRG (64). Additionally, inhibition of NOS resulted in reduced

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Arginine metabolism in virus pathogenesis

nitrotyrosine and prevented neuronal injury in FIV-infected DRG
cultures in vitro (64). These data suggest that lentivirus infection
contributes to axonal and neuronal injury through a mechanism
involving M1 macrophage immune activation mediated by STAT1 and iNOS activation. In addition to these studies, infection of
mice with the retrovirus LP-BM5, which causes profound immunodeficiency, induces CD11b+ GR-1+ Ly-6C+ MDSC-like cells that
inhibit both T- and B-cell responses in an iNOS/NO-dependent
but arginase-independent fashion (65). This study identified an
important – and only recently appreciated – role for iNOSexpressing myeloid cell-mediated suppression of B cell responses
in retrovirus infection.
MYELOID CELL PRODUCTION OF NO CAN INHIBIT VIRAL CLEARANCE

The oxidative effects of NO have also been shown to inhibit
immune cells, particularly T cells. This phenomenon has been
appreciated for a number of years in the context of tumors (92),
where myeloid suppressor cells can inhibit the anti-tumor T cell
response via the effects of NO in addition to other mechanisms (2,
4). In a similar manner, it has been shown that NO can inhibit the
antiviral immune response.
MCMV clearance from BALB/c mice is predominantly CD8+
T cell-mediated. A recent report showed that MCMV infection
in BALB/c mice induced CD11b+ Ly-6Chi inflammatory monocyte recruitment from the bone marrow to infected tissues that
was dependent on CCR2 signaling (66). This recruitment was
shown to inhibit antigen-specific CD8+ T cell activation, expansion, and cytotoxic activity via NO production, thus facilitating
viral persistence (66).
In a similar fashion, NO may contribute to a defective immune
response following infection of mice with an attenuated neurotropic coronavirus (rJ2.2 strain of mouse hepatitis virus). rJ2.2infected WT mice exhibited mild acute encephalitis, followed by a
non-lethal, chronic demyelinating disease (68). In marked contrast, rJ2.2 infection of mice that transgenically express CCL2
in the brain (CCL2 Tg) ineffectively cleared virus and rapidly
succumbed to the infection (68). CCL2 Tg mice mounted a dysregulated immune response, characterized by increased accumulation
of iNOS-expressing macrophages and microglia as well as regulatory T cells, but decreased Arg1 expression (68). These data suggest
that persistent CCL2 overexpression establishes and sustains an
immunological milieu that may predispose mice to a defective
immune response to a typically minimally virulent virus.

ARGINASE ACTIVITY CAN BE BENEFICIAL FOR TISSUE
REPAIR FOLLOWING VIRUS INFECTION
Arginase activity is important for wound healing and tissue regeneration through the production of polyamines and proline (2).
In the context of some viral infections, arginase activity and M2
macrophage activation have been shown to be beneficial for tissue
repair following virus-induced damage. For instance, resolution of
severe respiratory syncytial virus (RSV)-induced bronchiolitis in
mice is mediated by M2 macrophages that counteract cyclooxygenase (COX)-2-induced lung pathology (19, 20). Arg1 was induced
in the lungs of RSV-infected mice, and its induction was shown to
be IL-4Rα-dependent (19). Additionally, WT macrophages adoptively transferred into RSV-infected IL-4Rα−/− mice restored the

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Burrack and Morrison

M2 phenotype in the lungs and decreased lung pathology (19). It
was further shown that the lipoxogenase pathway was important
for M2 macrophage activation and lung resolution following RSV
infection (20). Most recently it was demonstrated that treating
mice with agents that sustain Arg1 expression (e.g., IL-4/anti-IL-4
immune complexes) limited RSV-induced lung pathology (21).
Consistent with a pathogenic role for iNOS/NO following
influenza infection (described above), it was recently shown that
the presence of airway bacteria polarize alveolar macrophages
into a M2 phenotype, thus limiting influenza-mediated lethal
lung inflammation. Wang and colleagues (27) demonstrated
that priming with Staphylococcus aureus, which commonly colonizes the upper respiratory mucosa, attenuated influenzamediated lung injury via TLR2 signaling that recruited peripheral CCR2+ CD11b+ monocytes into the alveoli (27). These
monocytes polarized alveolar macrophages into a M2 phenotype
characterized by high Arg1 as well as Ym1, FIZZ1, and IL-10
expression (27). It was further shown that S. aureus-primed M2
alveolar macrophages inhibited inflammatory cell recruitment
to the lung, including neutrophils, NK cells, and CD8 T cells
(27). S. aureus-primed M2 alveolar macrophages also expressed
higher levels of the inhibitory ligand PD-L1 (27), suggesting that
expression of a combination of anti-inflammatory cytokines and
inhibitory ligands could be the mechanisms by which S. aureusprimed M2 alveolar macrophages limit influenza-mediated lung
inflammation.
As discussed above, coxsackievirus B3 (CVB3) infection causes
myocarditis in human beings as well as in male BALB/c mice.
Although female mice do not develop severe myocarditis, both
male and female mice have comparable numbers of infiltrating
macrophages and viral titers in the heart following CVB3 infection (30). The macrophages infiltrating the heart in male mice were
skewed toward a M1 phenotype characterized by high expression
of iNOS (17) as well as M1-associated cytokines such as IFN-γ and
IL-12 (30). Additionally, inhibition of NOS resulted in increased
viral titers and higher mortality in CVB3-infected mice (17), consistent with an antiviral role for NO during CVB3 infection (see
above). However, in contrast to male mice, the heart-infiltrating
macrophages in female mice were skewed toward a M2 phenotype characterized by high expression of Arg1 as well as IL-4 and
IL-10 (30). Moreover, adoptive transfer of ex vivo-programed M1
macrophages significantly increased myocarditis in both male and
female mice. Strikingly, transfer of M2-programed macrophages
into susceptible male mice alleviated myocardial inflammation by
modulating the local cytokine profile from a M1 to M2 phenotype
and promoting peripheral regulatory T cell (Treg) differentiation
(30). Using different variants of CVB3, one that caused myocarditis in C57BL/6 mice and one that did not, it was additionally
shown that the myocarditic variant induced a M1 macrophage
phenotype (31). In contrast, the amyocarditic variant induced a
M2 macrophage phenotype, which was also associated with the
activation of NKT cells that promoted a Treg response (31). The
ability of NKT cells to suppress myocarditis was shown by adoptive
transfer of purified NKT cells into NKT knockout (Jα18 knockout)
mice infected with the myocarditic CVB3 variant, which inhibited
cardiac inflammation and increased Treg response (31). Cardiac
virus titers were equivalent in all mouse strains indicating that

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Arginine metabolism in virus pathogenesis

NKT cells did not participate in control of virus infection (31).
Thus, although NO appears to have antiviral properties against
CVB3, these data indicate an important role for Arg1-expressing
M2 macrophages in controlling CVB3-induced myocarditis.

ARGINASE ACTIVITY CAN PROMOTE VIRAL PERSISTENCE
AND/OR EXACERBATED IMMUNOPATHOLOGY
ARGINASE ACTIVITY CAN INHIBIT VIRAL CLEARANCE

As a consequence of their co-evolution with their hosts, viruses
have developed numerous strategies to evade the host immune system and ensure their own replication and survival. Recent studies
have identified a new evasion strategy for viruses: exploitation of
the host’s anti-inflammatory, wound repair response to promote
chronic infection.
Two strains of LCMV – Armstrong (Arm) and clone 13 (C13) –
have been studied for decades as models for acute and chronic
infections (93). Infection of mice with the Arm strain leads
to a robust CD8+ T cell response that rapidly clears the virus
(94), whereas infection with C13 results in T cells with impaired
functionality, enabling the virus to persist (95). It was recently
demonstrated that C13 infection led to an enhanced and sustained expansion of cells that resembled MDSCs (70). These
suppressive myeloid cells inhibited T cell proliferation ex vivo
via an iNOS/NO-dependent but Arg1-independent mechanism.
Another study, however, found that Arg1-expressing immunoregulatory antigen presenting cells induced during C13 infection
suppressed T cell responses (67). Most recently, it was demonstrated that T cell responses were improved – resulting in clearance
of the normally chronic C13 infection – when either myeloid cells
or T cells lacked IL-10 production (96). Overall, these data demonstrate the importance of iNOS/Arg1-expressing myeloid cells in
viral persistence.
Similar to LCMV C13 infection, it was recently demonstrated
that infection of mice with the arthritogenic alphaviruses Ross
River virus (RRV) and chikungunya virus (CHIKV) resulted in the
induction of Arg1 in macrophages in the infected and inflamed
musculoskeletal tissues (69). It was further shown that genetic
deletion of myeloid cell Arg1 resulted in enhanced viral control
in inflamed muscle tissue and reduced tissue pathology following RRV infection in mice (69), suggesting an important role for
Arg1-expressing macrophages in the persistence of these chronic
viruses.
Infection of mice with Theiler’s murine encephalomyelitis virus
(TMEV) results in persistent virus infection in the CNS, which
contributes to the development of a demyelinating disease that
has similarities with multiple sclerosis. Bowen and Olson (97)
showed that CD11b+ Ly-6C+ cells infiltrated the CNS following infection and were the dominant cell type during the innate
immune response. Depletion of the CD11b+ Ly-6C+ cells via
administration of an anti-GR-1 Ab resulted in reduced development of demyelinating disease and enhanced virus-specific CD4+
and CD8+ T cell responses (97). Additionally, TMEV-infected,
anti-GR-1 Ab-treated mice had decreased myelin-specific CD4+
T cell responses compared to control Ab-treated mice during the
demyelinating disease at a later time post-infection (97). Although
the expression of Arg1 was not investigated in this study, TMEVinfected mice had elevated expression of IL-10 in the brain and

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spinal cord (97), suggesting a role for this cytokine in the suppression of antiviral T cell responses, potentially through the effects
of Arg1.
Interestingly, a role for the modulation of arginine metabolism
in viral control versus persistence along with associated disease
has recently been demonstrated for the tumor-inducing, chickenspecific herpesvirus MDV. We mentioned above that MDV was
vulnerable to the antiviral properties of NO, with iNOS being
induced in genetically resistant chickens and in vaccinated chickens (35). In contrast, MDV induced strong macrophage arginase
activity in cell extracts from adherent monocytes from genetically susceptible chickens, but not in chickens that were resistant
to Marek’s disease, either genetically or acquired after vaccination (35). Together, these data suggest that in the case of Marek’s
disease, the state of resistance versus sensitivity to disease was correlated with a reciprocal balance of NOS versus arginase activities
in macrophages.
This phenomenon of Arg1-mediated T cell suppression has also
been recognized in human viral infections. Arg1 mRNA and protein levels were elevated in HCV-infected liver cell lines in vitro and
in HCV-infected liver samples compared with paired hepatocellular carcinoma samples from the same patients or with uninfected
liver tissues (71). Additionally, the number of MDSCs in chronic
HCV patients correlated with levels of plasma HCV-RNA (98). Cai
et al. (98) also found that MDSCs from patients with chronic HCV
infection suppressed T cell function via an Arg1-dependent mechanism. An additional study found that more PBMCs from chronic
HCV patients expressed the phenotypic markers of MDSCs than
PBMCs from healthy controls, and these cells expressed increased
levels of p47phox , a component of the NADPH oxidase complex
(99), suggesting a role for ROS in MDSC-mediated suppression. Consistent with this, CD33+ mononuclear cells co-cultured
with HCV-infected hepatocytes or HCV core protein suppressed
T cell proliferation in a ROS-dependent manner (99). Overall,
these data suggest that multiple mechanisms – including arginine
metabolism and ROS – may be at play in myeloid cell-mediated
suppression of anti-HCV T cell responses.
It has been suggested that prolonged immune activation during chronic virus infections, such as HCV and HIV, provides
an environment that drives viral replication and disease progression (100, 101). Moreover, immune activation can drive an
anti-inflammatory response to limit immunopathology, which
can be characterized by the presence of M2-like macrophages.
Indeed, similar to HCV infection, a role for arginase and M2polarized MDSC-like cells has been identified in the suppression of antiviral T cell responses following HIV infection. Individuals with detectable HIV-1 infection showed an increase
in the frequency of CD163+ CD16+ CD14+ monocytes, which
are thought to be precursors of M2 macrophages, when compared to seronegative or HIV-1-infected persons with undetectable viral loads, and monocyte frequency correlated positively
with HIV-1 viremia and negatively with CD4+ T cell counts
(in patients with counts <450 cells/µl) (72). Furthermore, Qin
and colleagues (73) observed elevated levels of MDSCs, defined
as HLA-DR−/low CD11b+ CD33+/high CD14+ CD15− cells, in the
peripheral blood of HIV-1-seropositive subjects compared with
healthy controls, and these MDSCs suppressed T cell responses

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Arginine metabolism in virus pathogenesis

in an Arg1-dependent manner. Moreover, PBMCs from HIVseropositive patients exhibited increased levels of arginase activity
(73). Cloke and colleagues (74) found that increased arginase
activity correlated with lower CD4+ T cell counts, and this association was abrogated following antiretroviral treatment (75).
Additionally, exposure of PBMCs to HIV gp120 expanded T cellsuppressive MDSCs in vitro (76). These data point to a direct role
for arginase-expressing MDSC-like cells in the suppression of antiHIV T cell responses. Consistent with that, individuals co-infected
with HIV and Leishmania parasites had increased arginase activity
in PBMCs and plasma compared with Leishmania-only infected
individuals, even though Leishmania infection alone results in
increased arginase activity (77). In addition, the parasite load in
the spleen was significantly higher in co-infected patients (77).
The arginase-expressing cells were identified as low-density granulocytes (77). These results suggest that increased arginase might
contribute to the poor immune responses and disease outcome
characteristic of patients with Leishmania and HIV co-infection.
Hepatitis B virus (HBV) infection is another common chronic
viral infection, with estimates as high as 350 million chronically
infected humans (102). Bility and colleagues (78) recently developed a humanized mouse model with both a human immune
system and human liver cells, named the A2/NSG-hu HSC/Hep
humanized mouse model, to study the pathogenesis of HBV
infection. Following HBV infection, the mice developed persistent HBV infection as well as chronic hepatitis and liver fibrosis
(78). The liver disease was associated with a high level of infiltrating human macrophages with a M2-like activation phenotype
(78). Similarly, M2-like macrophage accumulation was seen in
chronic HBV-infected patients, and M2-like macrophage induction in the liver was associated with accelerated liver fibrosis and
necrosis in patients with acute HBV-induced liver failure (78),
suggesting a role for M2 macrophages in persistent HBV infection. Additionally, patients with acute HBV infection had increased
serum levels of arginase, and this serum inhibited IFN-γ production by CD8+ T cells (79). Das et al. (80) also found decreased
l-arginine levels in the circulation of chronic HBV patients with
marked liver inflammation (>100 ALT) and increased arginase
activity in liver extracts taken directly ex vivo from patients with
chronic HBV compared with those from patients with other types
of liver pathology (80). They further showed that CD8+ T cells
from chronic HBV patients, regardless of their antigen specificity, exhibited less IL-2 but not IFN-γ or TNF-α production
and impaired proliferation following TCR-dependent stimulation,
indicating an aberrant antiviral T cell response in chronic HBV
infection (80). In the A2/NSG-hu HSC/Hep humanized mouse
model, HBV-infected mice had impaired liver T cell responses,
and M2 macrophages were associated with T cells in the liver
(78). Expression of the TCR signaling molecule CD3ζ was reduced
in both peripheral and intrahepatic CD8+ T cells from chronic
HBV patients; similarly, CD28 was also downregulated on CD8+
T cells from high viral load HBV patients (80). Downregulation
of the CD3ζ molecule has previously been shown to occur in the
arginine-depleted tumor microenvironment. Consistent with this,
in vitro transfection of CD3ζ and CD28 restored IL-2 production and supplementation of l-arginine partially restored CD3ζ
expression and T cell proliferation (80). These data suggest a role

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for arginase activity and arginine depletion in the impairment of
anti-HBV T cells functions.
In the absence of iNKT cells, influenza A (PR/8 strain) infection
was shown to induce the expansion of CD11b+ GR-1+ MDSCs in
the lungs of mice, which suppressed influenza-specific T cell and
antibody responses through the activity of both arginase and NOS,
resulting in higher viral titers and increased mortality (81). Adoptive transfer of iNKT cells reversed this phenotype; mice had an
increased survival rate, reduced viral titers, and increased virusspecific immune responses, suggesting a novel immunomodulatory role for iNKT cells during influenza virus infection (81).
Moreover, these authors identified that influenza infection in
humans induced the expansion of CD11b+ myeloid cells with
suppressive activity that could be reduced by iNKT cell activation
or the inhibition of arginase and NOS activity. Similarly, it was
recently shown that highly pathogenic H5N1 and H1N1 influenza
virus infection induced the accumulation of CD11b+ GR-1+ cells
and the expression of Arg1 in the lungs (82), further supporting a role for M2-polarized MDSC-like cells in promoting viral
persistence and immunopathology.
Helminth infection induces the expression of type 2 cytokines
and is associated with M2 macrophage activation, as determined by Arg1, FIZZ1, and Ym1 expression. Indeed, Osborne
and colleagues (83) found that Arg1, FIZZ1, and Ym1 were
highly induced in the ileum of mice infected with the helminth
Trichinella spiralis (Ts). Interestingly, they further showed that
co-infection of mice with Ts and murine norovirus (MNV)
resulted in decreased frequencies and numbers of MNV-specific
CD8+ and CD4+ T cells within the small intestine and spleen
as well as decreased polyfunctionality of these T cells, compared to Ts-only infected mice (83). Additionally, the defective
T cell responses were associated with increased viral loads in the
double-infected mice compared to the mono-infected controls
(83), suggesting that Ts-elicited M2-activated macrophages inhibited the antiviral T cell response to MNV. Lastly, neutralization
of Ym1, a chitinase-like molecule, in co-infected mice partially
restored antiviral immunity and was associated with enhanced
control of viral replication (83). These data point to a new mechanism by which Arg1-expressing macrophages inhibit antiviral
responses.
Cumulatively, these data are reminiscent of macrophages found
in tumors (e.g., MDSCs, TAMs) that have been shown to suppress anti-tumor T cell responses via a variety of NO- and/or
Arg1-dependent mechanisms (4, 5). Indeed, in a mouse model
of human papillomavirus (HPV)-induced cancer, Arg1-expressing
CD11b+ F4/80+ macrophages infiltrated the tumors and inhibited
T cell responses, including virus-specific T cells, by suppressing T cell proliferation and promoting a regulatory phenotype
(103). Moreover, depletion of the tumor-infiltrating macrophages
resulted in reduced tumor growth and increased tumor infiltration
by virus-specific CD8+ T cells (103). Thus, increasing evidence
points to a direct role for arginase-expressing M2-polarized cells in
the suppression of antiviral T cell responses and the persistence of a
variety of important pathogenic viruses. In addition to the actions
of iNOS and Arg1, MDSC-like cells can employ other mechanisms
to promote chronic viral infections, which were recently reviewed
by Goh and colleagues (104).

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Arginine metabolism in virus pathogenesis

M2 MACROPHAGE ACTIVATION CAN PROMOTE IMMUNOPATHOLOGY

In contrast to some parasitic infections where M2 macrophages
limit Th2 cell-mediated immunopathology, M2-polarized
macrophages have been shown to promote immunopathology
in some viral infections. For example, it was recently demonstrated that SARS-CoV infection of mice induced suppressive
alveolar macrophages that inhibited the induction of antiviral
T cell responses, a phenotype that was reversed by the adoptive transfer of activated bone marrow-derived DCs into mice
prior to virus infection (56). Additionally, SARS-CoV-infected
mice lacking hematopoietic STAT-1 expression were shown to
have greater weight loss and lung pathology, and this was associated with the activation of M2 macrophages (57). To further test
the role of M2 macrophages in enhanced pathogenesis following
SARS-CoV infection, the authors generated STAT-1/STAT-6 double knockout mice due to the established role for STAT-6 in driving
M2 macrophage activation in response to IL-4/IL-13 stimulation.
STAT-1/STAT-6 double knockout mice, which reversed the upregulation of M2 macrophages observed in STAT-1-deficient mice,
had reduced lung disease and prefibrotic lesions (57). These data
support the notion that M2 macrophages contribute to SARS-CoV
pathogenesis.
In another example, mice deficient in the IFN-γR exhibit
more severe disease following infection with murine gammaherpesvirus-68 (MHV-68), including interstitial and intra-alveolar
fibrosis that is reminiscent of idiopathic pulmonary fibrosis (IPF)
in human beings. In this model, alveolar macrophages were
recruited to the lungs of MHV-68-infected IFN-γR−/− mice, were
associated with areas of fibrosis, and exhibited a M2-polarized
phenotype characterized by the expression of FIZZ1, Ym1, and
Arg1 (61). Additionally, lung tissue from patients with IPF showed
increased expression of Arg1 in alveolar macrophages compared
with normal lung (61). These results suggest that virus-induced
upregulation of Arg1 could be mediating lung fibrogenesis. MHV68 infection in IFN-γR−/− mice also resulted in fibrosis in lymphoid tissues such as the spleen, which is a site of latent MHV-68
infection, and the liver (62, 63). Similar to the lung, MHV-68 infection in the absence of IFN-γR signaling induced a M2 macrophage
response in the spleen, characterized by high Arg1 expression
along with FIZZ1 and M2/Th2 cytokines such as IL-13, resulting in fibrotic disease in the spleen (105). Moreover, depletion of
T cells prevented MHV-68-mediated fibrosis in IFN-γR−/− mice
(62), suggesting that M2 macrophages were further driving Th2
activation to possibly create a M2/Th2 cytokine-induced cycle,
resulting in the exaggerated pathology. In contrast to IFN-γR−/−
mice, iNOS was induced in the spleen of MHV-68-infected WT
mice (105), indicating an important role for IFN-γ in inducing a
M1-associated immune response to control gamma-herpesvirus
infection and limiting Arg1-mediated immunopathology.

CONCLUSION
Macrophages and other myeloid cells have marked phenotypic heterogeneity, as a result of distinct cellular differentiation programs,
distribution in tissues, and responsiveness to various endogenous
and exogenous stimuli. Indeed, macrophages have well-established
roles in development, tissue homeostasis, coordinating the adaptive immune response and inflammation, as well as directing

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tissue resolution and repair following damage – processes that
are often modulated via the actions of the arginine-hydrolyzing
enzymes Nos2 and Arg1. We have highlighted a number of viral
infections in which these enzymes have a beneficial effect: NO
has antiviral properties against a variety of viruses, and arginase
activity can mediate tissue repair and regeneration following a
viral insult (Table 1). However, NO production can also result
in immunopathology in some virus infections, and the suppressive functions of Arg1-expressing macrophages can promote
immunopathology. Additionally, some viruses have exploited the
immune-suppressive properties of iNOS- and/or Arg1-expressing
macrophages to evade the immune response, particularly the
antiviral T cell response, resulting in chronic viral infections.
Clearly, iNOS- and/or Arg1-mediated responses are important in many viral infections. Thus, there is the potential to
develop the means to selectively stimulate or inhibit either M1
or M2 responses to mediate viral clearance or repair tissue damage. Due to the overlap in immunosuppressive mechanisms of
iNOS- and/or Arg1-expressing suppressor cells, therapeutic strategies under development to limit the immunosuppressive effects
of myeloid cells in cancer may be beneficial in treating persistent/chronic virus infections. However, as described above, iNOS
and Arg1 activity can be both beneficial and detrimental during
certain viral infections. Therefore, further research is needed to
define the molecular and tissue-specific mechanism(s) by which
iNOS and Arg1 influence the clearance of viral pathogens as well as
the injury and repair of tissues. In addition, a better understanding
of the pathways regulating macrophage polarization (specifically
iNOS and/or Arg1 induction and activity), macrophage trafficking, and the precise effects of iNOS and Arg1 activity on other
immune cells following different virus infections will inform the
development of therapeutics that target critical effector molecules
to promote viral control and limit immunopathology.

ACKNOWLEDGMENTS
Work in Dr. Morrison’s laboratory is supported by NIH-NIAID
grants U19 AI109680 and R01 AI108725. Kristina S. Burrack was
supported by NIH-NIAID training grant T32 AI052066.

REFERENCES
1. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol (2011) 11(11):723–37. doi:10.1038/nri3073
2. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
3. Munder M. Arginase: an emerging key player in the mammalian immune
system. Br J Pharmacol (2009) 158(3):638–51. doi:10.1111/j.1476-5381.2009.
00291.x
4. Gabrilovich DI, Ostrand-Rosenberg S, Bronte V. Coordinated regulation of
myeloid cells by tumours. Nat Rev Immunol (2012) 12(4):253–68. doi:10.1038/
nri3175
5. Kong YY, Fuchsberger M, Xiang SD, Apostolopoulos V, Plebanski M. Myeloid
derived suppressor cells and their role in diseases. Curr Med Chem (2013)
20(11):1437–44. doi:10.2174/0929867311320110006
6. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental
guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
7. Nathan CF, Hibbs JB Jr. Role of nitric oxide synthesis in macrophage antimicrobial activity. Curr Opin Immunol (1991) 3(1):65–70. doi:10.1016/09527915(91)90079-G
8. James SL. Role of nitric oxide in parasitic infections. Microbiol Rev (1995)
59(4):533–47.

www.frontiersin.org

Arginine metabolism in virus pathogenesis

9. Karupiah G, Xie QW, Buller RM, Nathan C, Duarte C, MacMicking JD. Inhibition of viral replication by interferon-gamma-induced nitric oxide synthase.
Science (1993) 261(5127):1445–8. doi:10.1126/science.7690156
10. Croen KD. Evidence for antiviral effect of nitric oxide. Inhibition of herpes simplex virus type 1 replication. J Clin Invest (1993) 91(6):2446–52.
doi:10.1172/JCI116479
11. Bi Z, Reiss CS. Inhibition of vesicular stomatitis virus infection by nitric oxide.
J Virol (1995) 69(4):2208–13.
12. Lin YL, Huang YL, Ma SH, Yeh CT, Chiou SY, Chen LK, et al. Inhibition of
Japanese encephalitis virus infection by nitric oxide: antiviral effect of nitric
oxide on RNA virus replication. J Virol (1997) 71(7):5227–35.
13. Neves-Souza PC, Azeredo EL, Zagne SM, Valls-de-Souza R, Reis SR, Cerqueira
DI, et al. Inducible nitric oxide synthase (iNOS) expression in monocytes during acute dengue fever in patients and during in vitro infection. BMC Infect
Dis (2005) 5:64. doi:10.1186/1471-2334-5-64
14. Zaragoza C, Ocampo C, Saura M, Leppo M, Wei XQ, Quick R, et al. The
role of inducible nitric oxide synthase in the host response to coxsackievirus
myocarditis. Proc Natl Acad Sci U S A (1998) 95(5):2469–74. doi:10.1073/pnas.
95.5.2469
15. Zaragoza C, Ocampo CJ, Saura M, Bao C, Leppo M, Lafond-Walker A, et al.
Inducible nitric oxide synthase protection against coxsackievirus pancreatitis.
J Immunol (1999) 163(10):5497–504.
16. Flodstrom M, Horwitz MS, Maday A, Balakrishna D, Rodriguez E, Sarvetnick N. A critical role for inducible nitric oxide synthase in host survival following coxsackievirus B4 infection. Virology (2001) 281(2):205–15.
doi:10.1006/viro.2000.0801
17. Lowenstein CJ, Hill SL, Lafond-Walker A, Wu J, Allen G, Landavere M, et al.
Nitric oxide inhibits viral replication in murine myocarditis. J Clin Invest
(1996) 97(8):1837–43. doi:10.1172/JCI118613
18. Karupiah G, Chen JH, Nathan CF, Mahalingam S, MacMicking JD. Identification of nitric oxide synthase 2 as an innate resistance locus against ectromelia
virus infection. J Virol (1998) 72(9):7703–6.
19. Shirey KA, Pletneva LM, Puche AC, Keegan AD, Prince GA, Blanco JC, et al.
Control of RSV-induced lung injury by alternatively activated macrophages
is IL-4R alpha-, TLR4-, and IFN-beta-dependent. Mucosal Immunol (2010)
3(3):291–300. doi:10.1038/mi.2010.6
20. Shirey KA, Lai W, Pletneva LM, Karp CL, Divanovic S, Blanco JC, et al. Role of
the lipoxygenase pathway in RSV-induced alternatively activated macrophages
leading to resolution of lung pathology. Mucosal Immunol (2013) 7(3):549–57.
doi:10.1038/mi.2013.71
21. Shirey KA, Lai W, Pletneva LM, Finkelman FD, Feola DJ, Blanco JC, et al.
Agents that increase AAM differentiation blunt RSV-mediated lung pathology.
J Leukoc Biol (2014). doi:10.1189/jlb.4HI0414-226R
22. Kodukula P, Liu T, Rooijen NV, Jager MJ, Hendricks RL. Macrophage control
of herpes simplex virus type 1 replication in the peripheral nervous system.
J Immunol (1999) 162(5):2895–905.
23. van Den Broek M, Bachmann MF, Kohler G, Barner M, Escher R,
Zinkernagel R, et al. IL-4 and IL-10 antagonize IL-12-mediated protection
against acute vaccinia virus infection with a limited role of IFN-gamma
and nitric oxide synthetase 2. J Immunol (2000) 164(1):371–8. doi:10.4049/
jimmunol.164.1.371
24. Conrady CD, Zheng M, Mandal NA, van Rooijen N, Carr DJ. IFN-alpha-driven
CCL2 production recruits inflammatory monocytes to infection site in mice.
Mucosal Immunol (2013) 6(1):45–55. doi:10.1038/mi.2012.46
25. Zolini GP, Lima GK, Lucinda N, Silva MA, Dias MF, Pessoa NL, et al. Defense
against HSV-1 in a murine model is mediated by iNOS and orchestrated by
the activation of TLR2 and TLR9 in trigeminal ganglia. J Neuroinflammation
(2014) 11:20. doi:10.1186/1742-2094-11-20
26. MacLean A, Wei XQ, Huang FP, Al-Alem UA, Chan WL, Liew FY. Mice lacking inducible nitric-oxide synthase are more susceptible to herpes simplex
virus infection despite enhanced Th1 cell responses. J Gen Virol (1998) 79(Pt
4):825–30.
27. Wang J, Li F, Sun R, Gao X, Wei H, Li LJ, et al. Bacterial colonization
dampens influenza-mediated acute lung injury via induction of M2 alveolar
macrophages. Nat Commun (2013) 4:2106. doi:10.1038/ncomms3106
28. Bodaghi B, Goureau O, Zipeto D, Laurent L, Virelizier JL, Michelson S. Role
of IFN-gamma-induced indoleamine 2,3 dioxygenase and inducible nitric
oxide synthase in the replication of human cytomegalovirus in retinal pigment
epithelial cells. J Immunol (1999) 162(2):957–64.

September 2014 | Volume 5 | Article 428 | 143

Burrack and Morrison

29. Noda S, Tanaka K, Sawamura S, Sasaki M, Matsumoto T, Mikami K, et al. Role
of nitric oxide synthase type 2 in acute infection with murine cytomegalovirus.
J Immunol (2001) 166(5):3533–41. doi:10.4049/jimmunol.166.5.3533
30. Li K, Xu W, Guo Q, Jiang Z, Wang P, Yue Y, et al. Differential macrophage
polarization in male and female BALB/c mice infected with coxsackievirus
B3 defines susceptibility to viral myocarditis. Circ Res (2009) 105(4):353–64.
doi:10.1161/CIRCRESAHA.109.195230
31. Liu W, Moussawi M, Roberts B, Boyson JE, Huber SA. Cross-regulation of T
regulatory-cell response after coxsackievirus B3 infection by NKT and gammadelta T cells in the mouse. Am J Pathol (2013) 183(2):441–9. doi:10.1016/j.
ajpath.2013.04.015
32. Harris N, Buller RM, Karupiah G. Gamma interferon-induced, nitric oxidemediated inhibition of vaccinia virus replication. J Virol (1995) 69(2):910–5.
33. Djeraba A, Bernardet N, Dambrine G, Quere P. Nitric oxide inhibits Marek’s
disease virus replication but is not the single decisive factor in interferongamma-mediated viral inhibition. Virology (2000) 277(1):58–65. doi:10.1006/
viro.2000.0576
34. Xing Z, Schat KA. Inhibitory effects of nitric oxide and gamma interferon
on in vitro and in vivo replication of Marek’s disease virus. J Virol (2000)
74(8):3605–12. doi:10.1128/JVI.74.8.3605-3612.2000
35. Djeraba A, Musset E, van Rooijen N, Quere P. Resistance and susceptibility to
Marek’s disease: nitric oxide synthase/arginase activity balance. Vet Microbiol
(2002) 86(3):229–44. doi:10.1016/S0378-1135(02)00010-X
36. Guidotti LG, McClary H, Loudis JM, Chisari FV. Nitric oxide inhibits hepatitis B virus replication in the livers of transgenic mice. J Exp Med (2000)
191(7):1247–52. doi:10.1084/jem.191.7.1247
37. Valero N, Espina LM, Anez G, Torres E, Mosquera JA. Short report: increased
level of serum nitric oxide in patients with dengue. Am J Trop Med Hyg (2002)
66(6):762–4.
38. Fagundes CT, Costa VV, Cisalpino D,Amaral FA, Souza PR, Souza RS, et al. IFNgamma production depends on IL-12 and IL-18 combined action and mediates
host resistance to dengue virus infection in a nitric oxide-dependent manner.
PLoS Negl Trop Dis (2011) 5(12):e1449. doi:10.1371/journal.pntd.0001449
39. Costa VV, Fagundes CT, Valadao DF, Cisalpino D, Dias AC, Silveira KD, et al.
A model of DENV-3 infection that recapitulates severe disease and highlights
the importance of IFN-gamma in host resistance to infection. PLoS Negl Trop
Dis (2012) 6(5):e1663. doi:10.1371/journal.pntd.0001663
40. Mendes-Ribeiro AC, Moss MB, Siqueira MA, Moraes TL, Ellory JC, Mann GE,
et al. Dengue fever activates the l-arginine-nitric oxide pathway: an explanation for reduced aggregation of human platelets. Clin Exp Pharmacol Physiol
(2008) 35(10):1143–6. doi:10.1111/j.1440-1681.2008.04970.x
41. Achike FI. The l-arginine-nitric oxide pathway: a potential therapeutic target in
dengue haemorrhagic fever. Clin Exp Pharmacol Physiol (2008) 35(10):1135–6.
doi:10.1111/j.1440-1681.2008.05022.x
42. Getts DR, Terry RL, Getts MT, Muller M, Rana S, Deffrasnes C, et al. Targeted blockade in lethal West Nile virus encephalitis indicates a crucial role for
very late antigen (VLA)-4-dependent recruitment of nitric oxide-producing
macrophages. J Neuroinflammation (2012) 9:246. doi:10.1186/1742-2094-9246
43. Tucker PC, Griffin DE, Choi S, Bui N, Wesselingh S. Inhibition of nitric
oxide synthesis increases mortality in Sindbis virus encephalitis. J Virol (1996)
70(6):3972–7.
44. Goody RJ, Hoyt CC, Tyler KL. Reovirus infection of the CNS enhances iNOS
expression in areas of virus-induced injury. Exp Neurol (2005) 195(2):379–90.
doi:10.1016/j.expneurol.2005.05.016
45. Rimmelzwaan GF, Baars MM, de Lijster P, Fouchier RA, Osterhaus AD. Inhibition of influenza virus replication by nitric oxide. J Virol (1999) 73(10):8880–3.
46. Darwish I, Miller C, Kain KC, Liles WC. Inhaled nitric oxide therapy fails
to improve outcome in experimental severe influenza. Int J Med Sci (2012)
9(2):157–62. doi:10.7150/ijms.3880
47. Burggraaf S, Bingham J, Payne J, Kimpton WG, Lowenthal JW, Bean AG.
Increased inducible nitric oxide synthase expression in organs is associated
with a higher severity of H5N1 influenza virus infection. PLoS One (2011)
6(1):e14561. doi:10.1371/journal.pone.0014561
48. Akaike T, Noguchi Y, Ijiri S, Setoguchi K, Suga M, Zheng YM, et al. Pathogenesis of influenza virus-induced pneumonia: involvement of both nitric
oxide and oxygen radicals. Proc Natl Acad Sci U S A (1996) 93(6):2448–53.
doi:10.1073/pnas.93.6.2448

Frontiers in Immunology | Antigen Presenting Cell Biology

Arginine metabolism in virus pathogenesis

49. Karupiah G, Chen JH, Mahalingam S, Nathan CF, MacMicking JD. Rapid interferon gamma-dependent clearance of influenza A virus and protection from
consolidating pneumonitis in nitric oxide synthase 2-deficient mice. J Exp Med
(1998) 188(8):1541–6. doi:10.1084/jem.188.8.1541
50. Jayasekera JP, Vinuesa CG, Karupiah G, King NJ. Enhanced antiviral antibody
secretion and attenuated immunopathology during influenza virus infection
in nitric oxide synthase-2-deficient mice. J Gen Virol (2006) 87(Pt 11):3361–71.
doi:10.1099/vir.0.82131-0
51. Perrone LA, Belser JA, Wadford DA, Katz JM, Tumpey TM. Inducible nitric
oxide contributes to viral pathogenesis following highly pathogenic influenza
virus infection in mice. J Infect Dis (2013) 207(10):1576–84. doi:10.1093/infdis/
jit062
52. Mgbemena V, Segovia JA, Chang TH, Tsai SY, Cole GT, Hung CY, et al. Transactivation of inducible nitric oxide synthase gene by Kruppel-like factor 6
regulates apoptosis during influenza A virus infection. J Immunol (2012)
189(2):606–15. doi:10.4049/jimmunol.1102742
53. Lin KL, Suzuki Y, Nakano H, Ramsburg E, Gunn MD. CCR2+ monocytederived dendritic cells and exudate macrophages produce influenzainduced pulmonary immune pathology and mortality. J Immunol (2008)
180(4):2562–72. doi:10.4049/jimmunol.180.4.2562
54. Aldridge JR Jr, Moseley CE, Boltz DA, Negovetich NJ, Reynolds C, Franks
J, et al. TNF/iNOS-producing dendritic cells are the necessary evil of lethal
influenza virus infection. Proc Natl Acad Sci U S A (2009) 106(13):5306–11.
doi:10.1073/pnas.0900655106
55. Konturek PC, Brzozowski T, Kania J, Konturek SJ, Kwiecien S, Pajdo R, et al.
Pioglitazone, a specific ligand of peroxisome proliferator-activated receptorgamma, accelerates gastric ulcer healing in rat. Eur J Pharmacol (2003)
472(3):213–20. doi:10.1016/S0014-2999(03)01932-0
56. Zhao J, Van Rooijen N, Perlman S. Evasion by stealth: inefficient immune activation underlies poor T cell response and severe disease in SARS-CoV-infected
mice. PLoS Pathog (2009) 5(10):e1000636. doi:10.1371/journal.ppat.1000636
57. Page C, Goicochea L, Matthews K, Zhang Y, Klover P, Holtzman MJ, et al.
Induction of alternatively activated macrophages enhances pathogenesis during severe acute respiratory syndrome coronavirus infection. J Virol (2012)
86(24):13334–49. doi:10.1128/JVI.01689-12
58. Adler H, Beland JL, Del-Pan NC, Kobzik L, Brewer JP, Martin TR, et al. Suppression of herpes simplex virus type 1 (HSV-1)-induced pneumonia in mice by
inhibition of inducible nitric oxide synthase (iNOS, NOS2). J Exp Med (1997)
185(9):1533–40. doi:10.1084/jem.185.9.1533
59. Gamba G, Cavalieri H, Courreges MC, Massouh EJ, Benencia F. Early inhibition of nitric oxide production increases HSV-1 intranasal infection. J Med
Virol (2004) 73(2):313–22. doi:10.1002/jmv.20093
60. Marques CP, Cheeran MC, Palmquist JM, Hu S, Lokensgard JR. Microglia
are the major cellular source of inducible nitric oxide synthase during experimental herpes encephalitis. J Neurovirol (2008) 14(3):229–38. doi:10.1080/
13550280802093927
61. Mora AL, Torres-Gonzalez E, Rojas M, Corredor C, Ritzenthaler J, Xu J, et al.
Activation of alveolar macrophages via the alternative pathway in herpesvirusinduced lung fibrosis. Am J Respir Cell Mol Biol (2006) 35(4):466–73. doi:10.
1165/rcmb.2006-0121OC
62. Dutia BM, Clarke CJ, Allen DJ, Nash AA. Pathological changes in the spleens
of gamma interferon receptor-deficient mice infected with murine gammaherpesvirus: a role for CD8 T cells. J Virol (1997) 71(6):4278–83.
63. Ebrahimi B, Dutia BM, Brownstein DG, Nash AA. Murine gammaherpesvirus68 infection causes multi-organ fibrosis and alters leukocyte trafficking in interferon-gamma receptor knockout mice. Am J Pathol (2001)
158(6):2117–25. doi:10.1016/S0002-9440(10)64683-4
64. Zhu Y, Jones G, Tsutsui S, Opii W, Liu S, Silva C, et al. Lentivirus infection
causes neuroinflammation and neuronal injury in dorsal root ganglia: pathogenic effects of STAT-1 and inducible nitric oxide synthase. J Immunol (2005)
175(2):1118–26. doi:10.4049/jimmunol.175.2.1118
65. Green KA, Cook WJ, Green WR. Myeloid-derived suppressor cells in murine
retrovirus-induced AIDS inhibit T- and B-cell responses in vitro that are used
to define the immunodeficiency. J Virol (2013) 87(4):2058–71. doi:10.1128/
JVI.01547-12
66. Daley-Bauer LP, Wynn GM, Mocarski ES. Cytomegalovirus impairs antiviral CD8+ T cell immunity by recruiting inflammatory monocytes. Immunity
(2012) 37(1):122–33. doi:10.1016/j.immuni.2012.04.014

September 2014 | Volume 5 | Article 428 | 144

Burrack and Morrison

67. Wilson EB, Kidani Y, Elsaesser H, Barnard J, Raff L, Karp CL, et al. Emergence of distinct multiarmed immunoregulatory antigen-presenting cells
during persistent viral infection. Cell Host Microbe (2012) 11(5):481–91.
doi:10.1016/j.chom.2012.03.009
68. Trujillo JA, Fleming EL, Perlman S. Transgenic CCL2 expression in the central nervous system results in a dysregulated immune response and enhanced
lethality after coronavirus infection. J Virol (2013) 87(5):2376–89. doi:10.1128/
JVI.03089-12
69. Stoermer KA, Burrack A, Oko L, Montgomery SA, Borst LB, Gill RG, et al.
Genetic ablation of arginase 1 in macrophages and neutrophils enhances
clearance of an arthritogenic alphavirus. J Immunol (2012) 189(8):4047–59.
doi:10.4049/jimmunol.1201240
70. Norris BA, Uebelhoer LS, Nakaya HI, Price AA, Grakoui A, Pulendran B.
Chronic but not acute virus infection induces sustained expansion of myeloid
suppressor cell numbers that inhibit viral-specific T cell immunity. Immunity
(2013) 38(2):309–21. doi:10.1016/j.immuni.2012.10.022
71. Cao W, Sun B, Feitelson MA, Wu T, Tur-Kaspa R, Fan Q. Hepatitis C virus targets over-expression of arginase I in hepatocarcinogenesis. Int J Cancer (2009)
124(12):2886–92. doi:10.1002/ijc.24265
72. Fischer-Smith T, Tedaldi EM, Rappaport J. CD163/CD16 coexpression by circulating monocytes/macrophages in HIV: potential biomarkers for HIV infection and AIDS progression. AIDS Res Hum Retroviruses (2008) 24(3):417–21.
doi:10.1089/aid.2007.0193
73. Qin A, Cai W, Pan T, Wu K, Yang Q, Wang N, et al. Expansion of monocytic myeloid-derived suppressor cells dampens T cell function in HIV-1seropositive individuals. J Virol (2013) 87(3):1477–90. doi:10.1128/JVI.0175912
74. Cloke TE, Garvey L, Choi BS, Abebe T, Hailu A, Hancock M, et al. Increased
level of arginase activity correlates with disease severity in HIV-seropositive
patients. J Infect Dis (2010) 202(3):374–85. doi:10.1086/653736
75. Cloke TE, Abebe T, Hailu A, Munder M, Taylor GP, Muller I, et al. Antiretroviral
therapy abrogates association between arginase activity and HIV disease severity. Trans R Soc Trop Med Hyg (2010) 104(11):746–8. doi:10.1016/j.trstmh.
2010.08.004
76. Garg A, Spector SA. HIV type 1 gp120-induced expansion of myeloid derived
suppressor cells is dependent on interleukin 6 and suppresses immunity. J Infect
Dis (2014) 209(3):441–51. doi:10.1093/infdis/jit469
77. Takele Y, Abebe T, Weldegebreal T, Hailu A, Hailu W, Hurissa Z, et al. Arginase
activity in the blood of patients with visceral leishmaniasis and HIV infection.
PLoS Negl Trop Dis (2013) 7(1):e1977. doi:10.1371/journal.pntd.0001977
78. Bility MT, Cheng L, Zhang Z, Luan Y, Li F, Chi L, et al. Hepatitis B virus infection and immunopathogenesis in a humanized mouse model: induction of
human-specific liver fibrosis and M2-like macrophages. PLoS Pathog (2014)
10(3):e1004032. doi:10.1371/journal.ppat.1004032
79. Sandalova E, Laccabue D, Boni C, Watanabe T, Tan A, Zong HZ, et al. Increased
levels of arginase in patients with acute hepatitis B suppress antiviral T cells.
Gastroenterology (2012) 143(1): 78–87.e3. doi:10.1053/j.gastro.2012.03.041
80. Das A, Hoare M, Davies N, Lopes AR, Dunn C, Kennedy PT, et al. Functional
skewing of the global CD8 T cell population in chronic hepatitis B virus infection. J Exp Med (2008) 205(9):2111–24. doi:10.1084/jem.20072076
81. De Santo C, Salio M, Masri SH, Lee LY, Dong T, Speak AO, et al. Invariant
NKT cells reduce the immunosuppressive activity of influenza A virus-induced
myeloid-derived suppressor cells in mice and humans. J Clin Invest (2008)
118(12):4036–48. doi:10.1172/JCI36264
82. Long JP, Kotur MS, Stark GV, Warren RL, Kasoji M, Craft JL, et al. Accumulation of CD11b(+)Gr-1(+) cells in the lung, blood and bone marrow of mice
infected with highly pathogenic H5N1 and H1N1 influenza viruses. Arch Virol
(2013) 158(6):1305–22. doi:10.1007/s00705-012-1593-3
83. Osborne LC, Monticelli LA, Nice TJ, Sutherland TE, Siracusa MC, Hepworth
HR, et al. Virus-helminth coinfection reveals a microbiota-independent mechanism of immunomodulation. Science (2014) 345(6196):578–82. doi:10.1126/
science.1256942
84. Stamler JS, Singel DJ, Loscalzo J. Biochemistry of nitric oxide and its redoxactivated forms. Science (1992) 258(5090):1898–902. doi:10.1126/science.
1281928
85. Lepoivre M, Chenais B, Yapo A, Lemaire G, Thelander L, Tenu JP. Alterations of
ribonucleotide reductase activity following induction of the nitrite-generating
pathway in adenocarcinoma cells. J Biol Chem (1990) 265(24):14143–9.

www.frontiersin.org

Arginine metabolism in virus pathogenesis

86. Kwon NS, Stuehr DJ, Nathan CF. Inhibition of tumor cell ribonucleotide reductase by macrophage-derived nitric oxide. J Exp Med (1991) 174(4):761–7.
doi:10.1084/jem.174.4.761
87. Wink DA, Kasprzak KS, Maragos CM, Elespuru RK, Misra M,
Dunams TM, et al. DNA deaminating ability and genotoxicity of nitric oxide
and its progenitors. Science (1991) 254(5034):1001–3. doi:10.1126/science.
1948068
88. Nguyen T, Brunson D, Crespi CL, Penman BW, Wishnok JS, Tannenbaum
SR. DNA damage and mutation in human cells exposed to nitric oxide
in vitro. Proc Natl Acad Sci U S A (1992) 89(7):3030–4. doi:10.1073/pnas.89.7.
3030
89. Goldstein DJ, Weller SK. Factor(s) present in herpes simplex virus type 1infected cells can compensate for the loss of the large subunit of the viral
ribonucleotide reductase: characterization of an ICP6 deletion mutant. Virology (1988) 166(1):41–51. doi:10.1016/0042-6822(88)90144-4
90. Jacobson JG, Leib DA, Goldstein DJ, Bogard CL, Schaffer PA, Weller SK, et al.
A herpes simplex virus ribonucleotide reductase deletion mutant is defective for productive acute and reactivatable latent infections of mice and for
replication in mouse cells. Virology (1989) 173(1):276–83. doi:10.1016/00426822(89)90244-4
91. Zell R, Markgraf R, Schmidtke M, Gorlach M, Stelzner A, Henke A, et al.
Nitric oxide donors inhibit the coxsackievirus B3 proteinases 2A and 3C
in vitro, virus production in cells, and signs of myocarditis in virus-infected
mice. Med Microbiol Immunol (2004) 193(2–3):91–100. doi:10.1007/s00430003-0198-6
92. Mills CD. Molecular basis of “suppressor” macrophages. Arginine metabolism via the nitric oxide synthetase pathway. J Immunol (1991) 146(8):
2719–23.
93. Ahmed R, Salmi A, Butler LD, Chiller JM, Oldstone MB. Selection of
genetic variants of lymphocytic choriomeningitis virus in spleens of persistently infected mice. Role in suppression of cytotoxic T lymphocyte response
and viral persistence. J Exp Med (1984) 160(2):521–40. doi:10.1084/jem.160.
2.521
94. Butz EA, Bevan MJ. Massive expansion of antigen-specific CD8+ T cells during
an acute virus infection. Immunity (1998) 8(2):167–75. doi:10.1016/S10747613(00)80469-0
95. Wherry EJ, Blattman JN, Murali-Krishna K, van der Most R, Ahmed R.
Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment. J Virol (2003)
77(8):4911–27. doi:10.1128/JVI.77.8.4911-4927.2003
96. Richter K, Perriard G, Behrendt R, Schwendener RA, Sexl V, Dunn R, et al.
Macrophage and T cell produced IL-10 promotes viral chronicity. PLoS Pathog
(2013) 9(11):e1003735. doi:10.1371/journal.ppat.1003735
97. Bowen JL, Olson JK. Innate immune CD11b+Gr-1+ cells, suppressor cells,
affect the immune response during Theiler’s virus-induced demyelinating disease. J Immunol (2009) 183(11):6971–80. doi:10.4049/jimmunol.0902193
98. Cai W, Qin A, Guo P, Yan D, Hu F, Yang Q, et al. Clinical significance and
functional studies of myeloid-derived suppressor cells in chronic hepatitis
C patients. J Clin Immunol (2013) 33(4):798–808. doi:10.1007/s10875-0129861-2
99. Tacke RS, Lee HC, Goh C, Courtney J, Polyak SJ, Rosen HR, et al. Myeloid
suppressor cells induced by hepatitis C virus suppress T-cell responses through
the production of reactive oxygen species. Hepatology (2012) 55(2):343–53.
doi:10.1002/hep.24700
100. Cassol E, Cassetta L, Alfano M, Poli G. Macrophage polarization and HIV-1
infection. J Leukoc Biol (2010) 87(4):599–608. doi:10.1189/jlb.1009673
101. Lawn SD, Butera ST, Folks TM. Contribution of immune activation to the
pathogenesis and transmission of human immunodeficiency virus type 1 infection. Clin Microbiol Rev (2001) 14(4):753–77. doi:10.1128/CMR.14.4.753-777.
2001
102. Liaw YF, Chu CM. Hepatitis B virus infection. Lancet (2009) 373(9663):582–92.
doi:10.1016/S0140-6736(09)60207-5
103. Lepique AP, Daghastanli KR, Cuccovia IM, Villa LL. HPV16 tumor associated macrophages suppress antitumor T cell responses. Clin Cancer Res (2009)
15(13):4391–400. doi:10.1158/1078-0432.CCR-09-0489
104. Goh C, Narayanan S, Hahn YS. Myeloid-derived suppressor cells: the dark
knight or the joker in viral infections? Immunol Rev (2013) 255(1):210–21.
doi:10.1111/imr.12084

September 2014 | Volume 5 | Article 428 | 145

Burrack and Morrison

105. Gangadharan B, Hoeve MA, Allen JE, Ebrahimi B, Rhind SM, Dutia BM, et al.
Murine gammaherpesvirus-induced fibrosis is associated with the development of alternatively activated macrophages. J Leukoc Biol (2008) 84(1):50–8.
doi:10.1189/jlb.0507270
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 29 July 2014; paper pending published: 17 August 2014; accepted: 22 August
2014; published online: 08 September 2014.

Frontiers in Immunology | Antigen Presenting Cell Biology

Arginine metabolism in virus pathogenesis

Citation: Burrack KS and Morrison TE (2014) The role of myeloid cell activation and
arginine metabolism in the pathogenesis of virus-induced diseases. Front. Immunol.
5:428. doi: 10.3389/fimmu.2014.00428
This article was submitted to Antigen Presenting Cell Biology, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Burrack and Morrison. This is an open-access article distributed
under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s)
or licensor are credited and that the original publication in this journal is cited, in
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ORIGINAL RESEARCH ARTICLE
published: 28 October 2014
doi: 10.3389/fimmu.2014.00537

Classical and alternative activation and metalloproteinase
expression occurs in foam cell macrophages in male and
female ApoE null mice in the absence of T and
B lymphocytes
Elaine Mo Hayes, Aikaterini Tsaousi , Karina Di Gregoli , S. Rhiannon Jenkinson, Andrew R. Bond ,
Jason L. Johnson, Laura Bevan, Anita C. Thomas † and Andrew C. Newby *†
Bristol Heart Institute, School of Clinical Sciences, University of Bristol, Bristol, UK

Edited by:
Klaus Ley, La Jolla Institute for Allergy
& Immunology, USA
Reviewed by:
Carolyn Louise Geczy, University of
New South Wales, Australia
Norbert Leitinger, University of
Virginia, USA
Almudena Ortega-Gomez,
Ludwig-Maximilians-Universität
München, Germany
*Correspondence:
Andrew C. Newby , Bristol Heart
Institute, School of Clinical Sciences,
University of Bristol, Bristol Royal
Infirmary, Bristol BS2 8HW, UK
e-mail: a.newby@bristol.ac.uk
†

Anita C. Thomas and Andrew C.
Newby have contributed equally to
this work.

Background: Rupture of advanced atherosclerotic plaques accounts for most lifethreatening myocardial infarctions. Classical (M1) and alternative (M2) macrophage activation could promote atherosclerotic plaque progression and rupture by increasing production
of proteases, including matrix metalloproteinases (MMPs). Lymphocyte-derived cytokines
may be essential for generating M1 and M2 phenotypes in plaques, although this has not
been rigorously tested until now.
Methods and results: We validated the expression of M1 markers (iNOS and COX-2)
and M2 markers (arginase-1, Ym-1, and CD206) and then measured MMP mRNA levels in mouse macrophages during classical and alternative activation in vitro. We then
compared mRNA expression of these genes ex vivo in foam cells from subcutaneous
granulomas in fat-fed immune-competent ApoE knockout (KO) and immune-compromised
ApoE/Rag-1 double-KO mice, which lack all T and B cells. Furthermore, we performed
immunohistochemistry in subcutaneous granulomas and in aortic root and brachiocephalic
artery atherosclerotic plaques to measure the extent of M1/M2 marker and MMP protein
expression in vivo. Classical activation of mouse macrophages with bacterial lipopolysaccharide in vitro increased MMPs-13, -14, and -25 but decreased MMP-19 andTIMP-2 mRNA
expressions. Alternative activation with IL-4 increased MMP-19 expression. Foam cells in
subcutaneous granulomas expressed all M1/M2 markers and MMPs at ex vivo mRNA and
in vivo protein levels, irrespective of Rag-1 genotype. There were also similar percentages
of foam cell macrophages (FCMs) carrying M1/M2 markers and MMPs in atherosclerotic
plaques from ApoE KO and ApoE/Rag-1 double-KO mice.
Conclusion: Classical and alternative activation leads to distinct MMP expression patterns
in mouse macrophages in vitro. M1 and M2 polarization in vivo occurs in the absence of T
and B lymphocytes in either granuloma or plaque FCMs.
Keywords: atherosclerosis, macrophages, lymphocytes, cytokines, plaque rupture

INTRODUCTION
Atherosclerotic plaque rupture underlies most myocardial infarctions and thromboembolic strokes, which are principal causes
of mortality and morbidity worldwide (https://apps.who.int/
infobase/mortality.aspx). Macrophages play a key role in atherosclerosis progression, as demonstrated by their abundance in
human plaques as foam cells and by the dramatic reduction in
atherosclerosis in mice after genetic (1) or pharmacological (2)
deletion of macrophages. Furthermore, production of mediators
from activated macrophages is believed to be important in making plaques vulnerable to rupture (3). Reactive oxygen species,
cytokines, and cell surface modifying proteinases produced by
macrophages promote apoptosis, thereby contributing to the cellular rarefaction of vulnerable plaques (4). Moreover, extracellular
proteinases, in particular matrix metalloproteinases (MMPs) can

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directly degrade the extracellular matrix and promote plaque
instability (5). These proposed mechanisms are supported by the
histological appearance of vulnerable plaques, which contain an
abundance of macrophages expressing MMPs but a relative lack
of smooth muscle cells (SMC) and extracellular matrix proteins,
importantly collagens, which contribute tensile strength to the
plaque cap (5). Intervening to diminish the production of these
harmful mediators is therefore a rational approach to new therapies and this motivates efforts to understand the cellular and
molecular mechanisms involved.
Early work highlighted the functional diversity of macrophages
(6) and the existence of distinct phenotypes has become increasingly discussed (7). Polarization of macrophage into a so-called
classically activated or M1 phenotype was recognized, possibly
reflecting the context of infection where pathogen associated

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Hayes et al.

molecular patterns (PAMPs) and pro-inflammatory lymphocytederived cytokines, particularly interferons (IFNs) and interleukins
(ILs) occurred in the same microenvironments. Consistent with
this, the M1 phenotype is simulated in vitro by the combined
action of PAMPs acting through Toll-like receptors (TLRs) and
IFNγ (8), with some evidence for synergy. Mechanisms underlying synergy include the ability of IFNγ to prime responses to
PAMPs by inducing expression of TLRs and their co-activators (9).
Synergy also results from the combined activation of differing signaling pathways for TLRs through nuclear factor-κB (NF-κB) (9)
and IFNγ through signal transducer and activator of transcription
(STAT-1) (10).
The actions of IFNγ have led to the hypothesis that Thelper1
(Th1)-lymphocytes may be essential for, or at least prominent
contributors to, M1 polarization in vivo, a concept that has been
acknowledged in papers dealing with atherosclerosis (11–13). It is
consistent with studies showing that knockout (KO) of IFNγ and
its receptors reduces atherosclerosis progression (3), although the
impact on M1 polarization was not measured directly in any of
these IFNγ KO investigations. On the other hand, it is well recognized that other cytokines and combinations of cytokines that
activate the pathways leading to M1 states could bypass the requirement for IFNγ (9, 14). Conversely, macrophages can polarized to
a variety of alternatively activated or M2 states. The so-called M2a
phenotype occurs after treatment with IL-4 or IL-13, which are
potentially produced by Thelper2 (Th2) lymphocytes (15). Overall, this leads to the idea that T-lymphocytes, and, by implication,
adaptive immunity play an essential part in M1/M2 macrophage
polarization in atherosclerotic plaques. On the other hand, data
from other areas of inflammation support the conclusion that
M1/M2 polarization is primarily a function of innate immunity
and that lymphocytes play a minor, at best modulatory role (16).
Resolving this controversy is important for human atherosclerosis
since a variety of immunotherapies have been proposed and some
have already entered clinical trials (17).
Our present aim was formally to test the requirement for T and
B lymphocytes in M1 and M2 macrophage activation and MMP
production using an established ApoE-KO mouse model of atherosclerosis formation. To do this, we compared the expression of
M1 and M2 markers, MMPs and the endogenous tissue inhibitors
of MMPs (TIMPs) in foam cell macrophages (FCMs) from subcutaneous granulomas and atherosclerotic plaques in ApoE KO and
ApoE/Rag-1 double-knockout (DKO) mice, which lack all T and
B lymphocytes. As it is already known that there are differences
in inflammation and plaque development between genders (18,
19), we used both male and female mice in this study. The mice
were fed on a high-fat diet because it was previously shown that
these conditions produced similar sized plaques in both genotypes
(20). Hence, we could be sure that measurements of the prevalence
of macrophages phenotypes would not be an artifact of different
stages of atherosclerosis progression or plaque sizes.

MATERIALS AND METHODS
BONE MARROW MONOCYTE ISOLATION AND DIFFERENTIATION TO
BONE MARROW-DERIVED MACROPHAGES

Mouse femurs and tibias were excised from C57BL/6 mice (Charles
River, UK) on a normal diet. The bone ends were cut and bone

Frontiers in Immunology | Inflammation

Foamy macrophage activation without lymphocytes

marrow was flushed out with sterile phosphate-buffered saline
(PBS). Erythrocytes were removed using ACK lysis buffer (Life
Technologies, UK) and the pelleted white blood cells resuspended
in PBS and counted, giving an average yield of 40 × 106 cells per
mouse. Cells were plated into 24-well plates at 0.5 × 106 cells/well
and grown in RPMI 1640 media (Life Technologies) supplemented
with antibiotics, glutamine, and 20% fetal calf serum (FCS) (Life
Technologies) in the presence of 20 ng/ml recombinant human
macrophage colony-stimulating factor (M-CSF, R&D Systems,
USA). The media was changed every 3–4 days. After 7–10 days,
when the cells had differentiated into macrophages, M-CSF was
removed and the cells exposed to selected cytokines for 18 h in
serum-free media (SFM), as specified in the text. These included
mouse (m) IFNγ at 20 ng/ml (Miltenyi Biotec, USA), human
(h) tissue necrosis factor (TNF)-α at 10 ng/ml (R&D Systems,
USA), mIL-4 at 10 ng/ml (PreproTech, UK), and lipopolysaccharide (LPS) at 10 ng/ml (Sigma-Aldrich, USA, L2654). Human
cytokines were used only when their efficacy on mouse cells had
been previously documented by the suppliers.
PERIPHERAL BLOOD MONONUCLEAR CELL ISOLATION AND
DIFFERENTIATION

Pooled peripheral blood was collected by cardiac puncture from
C57BL/6 mice with heparin as anti-coagulant. Monocytes were
then isolated density gradient separation on Ficoll Paque Plus and
differentiated to macrophages as previously described (21).
RNA EXTRACTION, REVERSE TRANSCRIPTION, AND QUANTITATIVE PCR

RNA lysates from cultured macrophages were collected using RLT
solution (Qiagen Ltd, UK) with β-mercaptoethanol and the total
RNA extracted using the Qiagen RNeasy kit (Qiagen Ltd), according to the manufacturer’s instructions. The quantity and quality
of resulting RNA was assessed using a NanoDrop ND-1000 spectrophotometer (LabTech International, UK). Samples of cDNA
were generated using QuantiTect Reverse Transcription Kit (Qiagen Ltd), according to the manufacturer’s instructions and the
resulting cDNA was diluted 1:1 in 10 mM TrisHCl, pH 8.0. Real
time quantitative PCR was performed in a Roche Light Cycler 1.5
(Roche, UK) to quantify the steady-state concentration of RNA,
using the QuantiTect SYBR Green PCR Kit (Qiagen Ltd). The
primers used are listed in Table 1. Each reaction contained 2.5–7 ng
RNA and 0.5 µM primers. Initial denaturation (15 min at 95°C)
was followed by 55 cycles of denaturation (15 s at 95°C), annealing
(20 s at 60°C), and extension (25 s at 72°C). Copy numbers of gene
transcripts per total nanogram RNA input were calculated using
standard curves constructed as recommended by from purified
amplicon (Bioline, USA).
IN VIVO STUDIES

Rag-1 KO mice that do not produce mature T or B cells (B6.129S7Rag-1tm1Mom /J) and ApoE KO mice on a C57BL/6 background
(B6.129P2-Apoetm1Unc /J) were purchased from The Jackson Laboratories (USA), and bred together to create ApoE± /Rag-1± mice.
Breeding stocks of ApoE KO and ApoE/Rag-1 DKO mice were
obtained by crossing the resulting F1 generation. Mice were kept
in scantainers and given sterile food and water ad libitum. All
animal work was in accordance with the Home Office Guidance

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Foamy macrophage activation without lymphocytes

Table 1 | Primer sequences used for quantitative RT-PCR.
Gene
ARG-1

Primer sequence
AGTCTGGCAGTTGGAAGCATCTCT
TTCCTTCAGGAGAAAGGACACAGG

COX-2

ATACTGGAAGCCGAGCACCTTTGG
ATGGTGGCTGTTTTGGTAGGCTGT

CD206

CCATTTATCATTCCCTCAGCAAGC
AAATGTCACTGGGGTTCCATCACT

FIZZ1

AGAGGTGGAGAACCCAGCTTTGAT
TTTCAAGAAGCAGGGTAAATGGGCA

IL-12p35

CCACAACAAGAGGGAGCTGCCTGCCC
AGTGCTGCGTTGATGGCCTGGAACT

IL-12p40

AGACCAGGCAGCTCGCAGCAAAGCA
GACACATCCCACTCCCACGCTGCC

iNOS

CTCATGACATCGACCAGAAGCGT
TATATTGCTGTGGCTCCCATGTTG

MMP-2

GGCTGACATCATGATCAACTTTGG
GCCATCAGCCGTTCCCATACTTTAC

MMP-3

GCATCCCCTGATGTCCTCGTGG
TCCCCGGAGGGTGCTGACTG

MMP-8

TGCCTCGATGTGGAGTGCCTGA

MMP-9

AGAGAGGAGTCTGGGGTCTGGTTT

MMP-12

AATTACACTCCGGACATGAAGCGT

MMP-13

ATGATGATGAAACCTGGACAAGCA

GCCCTTGACAGCTGTGGCGT
GAGAACACCACCGAGCTATCCACT
GGCTAGTGTACCACCTTTGCCATC
ATAGGGCTGGGTCACACTTCTCTG
MMP-14

ACCACAAGGACTTTGCCTCTGAAG
CACCGAGCTGTGAGATTCCCTTGA

MMP-19

GATGAACTGGCCAGAACTGACCTT
GTCCCCGGTTGATGAGTTAGTGTC

MMP-23

CAAGGTTGGTGAGAGAGGGTAGGA
AGGAGTAGGTGCTGAGAACACGCT

MMP-25

CTCTGAGTGGCAGTGTTTGGAAGA
TGATGTCAGGCTCCTGGTACTGAG

TIMP-1

AGGAACGAAATTTGCACATCAGT
CAAAGTGACGGCTCTGGTAGTCCT

TIMP-2

GACTCCCCCTCAGACTCTCCCTAC
CATATTGATACCACCGCACAGGAA

TIMP-3

CACATCAAGGTGCCATTCAGGTAG
GTTCTCTCCTCCTCAACCCAAACA

Ym1

CAGGTCTGGCAATTCTTCTG
GTCTTGCTCATGTGTGTAAGTG

on the operation of the Animals (Scientific Procedures) Act 1986
and conforms to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health
(NIH Publication No. 85–23, revised 1996). Genotyping of the
mice was performed on ear or tail pieces, using the Direct PCR
kit (Bioquote, UK or Viagen Biotech, USA) after a Proteinase K
digestion (Sigma, UK). PCR was performed using Crimson Taq
Pol (New England Biolabs, UK), with dNTPs from Bioline (UK),
using primers designed by The Jackson Laboratories. Male and

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female mice commenced a sterile high-fat diet (21–23% fat, Special
Diet Services, UK) at 5 weeks of age, and were sacrificed with an
anesthetic overdose 12 weeks later. Blood was taken via cardiac
puncture and heparinized plasma was subsequently analyzed for
total, HDL, and LDL/VLDL cholesterol (Cholesterol/Cholesterol
Ester Quantification Kit, Abcam, UK), after a minor adaptation
of the manufacturer’s instructions. The levels of selected M1 and
M2 cytokines was assessed in additional samples of mouse plasma
using a Bio-Plex Pro Mouse Cytokine Th1/Th2 Panel 8-plex (BioRad, USA). After the cardiac puncture, fresh tissue samples were
taken (tail tip, spleen, liver lobe) and the animals perfused via
the heart with PBS, then 10% formalin, at a constant pressure of
100 mmHg, with outflow through the left jugular vein. The brachiocephalic artery (BCA) (with a small piece of aortic arch), heart
and remaining ascending and descending aorta were cleaned and
removed from each mouse. Other tissues harvested included thymus and remaining liver. Tissue blocks of spleen less than 0.5 cm
thick were post-fixed in 10% formalin for 24 h for subsequent
histological examination, as described below.
SPONGE IMPLANTATION AND FOAM CELL MACROPHAGE ISOLATION

ApoE KO or DKO mice were fed the same high-fat diet as above
from 6 weeks of age. Two weeks later these mice had 0.5 cm3 sterile
polyurethane sponges containing ~50 µl of Matrigel (VWR, UK)
placed under the dorsal skin under halothane anesthesia to generate FCMs as described previously (22, 23). The mice were fed
the high-fat diet for a further 4 weeks. Recovered sponges were
either fixed and embedded for immunohistological examinations
as described below or the FCM were isolated and studied ex vivo.
Fresh sponges were treated with 0.75 ml undiluted Dispase (VWR,
UK or BD Biosciences, USA) and then squeezed to obtain a cellular
exudate. FCM were then purified, as previously described (22, 24)
by flotation after centrifugation on a metrizamide gradient (1.3507
refractive index, Sigma) followed by differential adherence. Only
foam cells (validated by Oil-red-O staining) float because of the
relatively low buoyant density of lipid.
Samples were taken from each preparation immediately for
protein or RNA isolation, and mRNA levels quantified as described
above. Other cell preparations were cultured for a short period, to
allow adherence to coverslips. Oil-Red-O (2% Oil-red-O in isopropanol; Sigma) staining was performed to confirm lipid content,
and immunocytochemistry performed to confirm cell purity. Cells
were also assessed for their proliferative capacity [BrdU (Sigma)
incorporation, 8 h pulse] or in situ zymography (25). In this assay,
the gelatinolytic capacity of the macrophages isolated from the
sponges was determined using the EnzChek gelatinase/collagenase
assay kit (Invitrogen, USA). Controls included cells treated with
EDTA, 1,10-phrenanthroline (Sigma) or GM6001 (Millipore, UK),
to prevent MMP activity. Cells were fixed in paraformaldehyde
and mounted in Vectorshield + DAPI (Vector Labs, USA). Several
fields were photographed on each coverslip and the proportion of
cells with gelatinase activity as indicated by the loss of fluorescence
of the DQ-gelatin substrate determined.
HISTOLOGICAL METHODS

The proximal aorta and BCA from each mouse were embedded
in paraffin and 3 µm sections cut at 3 µm intervals from the

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Hayes et al.

Foamy macrophage activation without lymphocytes

atherosclerosis-prone areas of these vascular beds, as described
previously (23, 26). The first section after the bifurcation of the
BCA from the aorta was cleared and rehydrated and then stained
using Miller’s elastin/van Gieson (EVG) and plaque dimensions
were measured using image analysis software (Image Pro, DataCell, Maidenhead, UK), as described previously (23). The aortic
sinus from each mouse was treated and examined in a similar
fashion, with the first leaflet section (from the aorta) stained
using EVG, with subsequent image analysis being performed (26).
For immunohistochemistry, 3 µm sections were brought to water
and antigen retrieval performed using citrate buffer. Non-specific
binding blocked with 10% goat serum (Sigma) in PBS. Primary
antibodies for SMC (α-smooth muscle actin), macrophages [Griffonia simplicifolia Lectin II (GSL)], iNOS, COX-2, CD206, arg-1,
Ym-1, MMP-12, MMP-13, MMP-14, and TIMP-3 (see Table 2)
were added to the sections and incubated either overnight at
4°C or for 1 h at room temperature. After washing and further incubations with goat anti-rabbit-biotin (Dako or Sigma)
and ExtrAvidin-HRP (Sigma) staining was visualized using 3,3’diaminobenzidine (DAB, Sigma). A negative control where the
primary antibody was replaced with the relevant species IgG at
the same dilution was always included. The percentage of the

plaque area stained with each cell-specific or phenotypic marker
or MMP/TIMPs antibody was determined using the same image
analysis software detailed above. The number of buried layers was
assessed manually on sections stained with EVG and on sections
using antibodies that recognize SMC. Paraffin-embedded sponge
sections were treated similarly, and the presence of markers of
macrophage activation examined. Oil-Red-O staining was performed en face, and the percentage of fatty deposits in each aorta
was measured using NIH ImageJ v1.43.
STATISTICAL METHODS

All analyses were performed using GraphPad InStat v3.05 (GraphPad Software, Inc. USA) or SPSS v21 (IBM, USA) software. Data
were checked for normality (Kolmogorov and Smirnov normality
test), and logarithmic transformation of data performed if necessary. Regression analyses were performed using Pearson’s correlation co-efficient. Statistical analyses of data were performed using
Students t -test, a Mann–Whitney U -test or 1- or 2-way ANOVAs,
with the 1-way ANOVA followed by a Bonferroni or Tukey-Kramer
post-test. Data are expressed as arithmetic mean ± SEM or geometric mean and 95% confidence limits, and statistical significance
defined as P < 0.05.

RESULTS
Table 2 | Primary and secondary antibodies and Griffonia simplicifolia
Lectin II.
Catalog

Type

Supplier

Vector labs

number
Primary
GSL II

B1215

Lectin

α-Smooth muscle actin

M0851

M_Mab

Dako

arginase 1

sc-20150

Rb_PAb

Santa Cruz

BrdU

B2531

M_Mab

Sigma

COX2

ab15191

Rb_PAb

Abcam

iNOS

ab15323

Rb_PAb

Abcam

MOMA-2

ab33451

R_Mab

Abcam

MMP-12

ab52897

Rb_MAb

Abcam

MMP-13

ab39012

Rb_PAb

Abcam

MMP-14

ab51074

Rb_MAb

Abcam

STAT1p (phospho Y701)

ab30645

Rb_PAb

Abcam

STAT6p (phospho Tyr641)

06–937

Rb_PAb

Millipore

TIMP-3

Ab39206

Rb_PAb

Abcam

Ym1/2

01404

Rb_PAb

Stemcell
Technologies

rabbit IgG negative control

I5006

Sigma

mouse IgG2a negative

M5409

Sigma

control
Goat serum

G9023

Sigma

Rabbit serum

R9133

Sigma

Secondary antibody
GtαM

E0433

Biotinylated

Dako

GtαM

B6649

Biotinylated

Sigma

GtαRb

E0432

Biotinylated

Dako

GtαRb

B6649

Biotinylated

Sigma

RbαR

E0468

Biotinylated

Dako

Frontiers in Immunology | Inflammation

IN VITRO STUDIES IN MOUSE BONE MARROW MACROPHAGES

Bone marrow proved a convenient source of large quantities of
mouse monocytes that were converted to bone marrow-derived
macrophages (BMDM) using M-CSF. BMDM were 97% F4/80
and CD11b double positive by flow cytometry (results not shown).
We used mRNA expression for established M1 and M2 marker
genes as positive controls for classic or alternatively activation. As
expected from previous literature (15), classical activation with
LPS alone or LPS plus IFNγ increased mRNA levels of inducible
NO synthase (iNOS, NOS-2) and cyclooxygenase-2 (COX-2)
(Figure 1A), whereas alternative activation with IL-4 increased
mRNA expression of arginase-I (arg-1), Ym-1, and CD206. We
then investigated the concomitant regulation of a wide spectrum of
MMPs and TIMPs, many of which have been previously implicated
in atherosclerosis (27). The most abundant mRNAs under unstimulated conditions were MMP-12 > MMP-8 = MMP-19 = MMP14 = TIMP-2> other MMPs and TIMPs (Figure 1B). Among the
MMPs studied, MMP-13 showed the most dramatic 121-fold stimulation by LPS + IFNγ (Figure 1B, note the scale is logarithmic).
LPS + IFNγ treatment also increased expressions of MMP-14
(11.3-fold) and MMP-25 (14.6-fold); and decreased expressions
of MMP-19 (4.5-fold) and TIMP-2 (9.0-fold) (Figure 1B). Classical activation with a different mediator, tumor necrosis factorα
(TNFα), significantly increased expressions of MMP-2 (77-fold),
MMP-9 (3.5-fold), and MMP-14 (3.5-fold) but not of MMP-13
(Figure 1B). Classical activation with TNFα also increased MMP9 and MMP-14 expression in blood derived macrophages, similar
to BMDM, but did not affect any of the other MMPs or TIMPs
(results not shown). Treatment with IFNγ did not increase mRNA
levels of any MMP or TIMP in BMDM either alone or in the presence of LPS (Figures 1C,D). No effect of IFNγ was observed in
blood derived macrophages either (results not shown). Alternative
activation with IL-4 increased mRNA expression of only MMP-19

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FIGURE 1 | Continued

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Hayes et al.

Foamy macrophage activation without lymphocytes

FIGURE 1 | Continued
Effect of M1/M2 polarization on mRNAs for markers, MMPs, and
TIMPs in mouse bone marrow-derived macrophages. Monocytes
from mouse bone marrow were differentiated to macrophages in M-CSF
and then treated with bacterial lipopolysaccharide (LPS), interferon-γ
(IFNγ), tumor necrosis factor-α (TNFα), or interleukin-4 (IL-4) alone or in
combination, as indicated. SFM = serum-free medium. (A) The effect of

polarization by IFNγ, LPS, LPS + IFNγ, or IL-4 on M1/M2 markers
(n = 4–5). (B) The effect of polarization by IFNγ + LPS, TNFα, or IL-4 on
MMPs and TIMPs (n = 3–8). (C) Effect of IFNγ alone on macrophages
incubated in SFM. (D) Effect of IFNγ on macrophages incubated in SFM
containing LPS. Values are expressed as mean and SEM (A) or mean and
95% confidence intervals (B–D). *M1 different from M0, # M2 different
from M0, P < 0.05.

Table 3 | Concentration of inflammatory and anti-inflammatory cytokines in plasma from ApoE KO or ApoE/Rag1 KO mice.
Male ApoE KO
Mean

SEM

Male DKO

P-value (males)

Mean

SEM

Female ApoE KO

Female DKO

Mean

SEM

Mean

SEM

P-value (females)

IFNγ

11.88

4.353

5.680

2.902

0.2440

11.33

6.345

6.012

2.595

0.6275

TNFα

297.4a

56.00

206.3

23.16

0.1495

167.9

21.46

191.0

19.26

0.4331

GM-CSF

118.0

24.55

106.4

16.66

0.6959

72.85

11.24

113.6

13.22

0.0304

IL-12p70

133.6

38.79

129.9

37.60

0.9457

58.65

12.61

127.7

29.90

0.0302

IL-10

15.23

3.023

23.99

3.852

0.0964

10.32

3.0.03

25.38

2.486

0.0012

Concentrations (picograms per milliliter) are expressed as mean and SEM, n = 9–10. Bold type indicates P < 0.05 for ApoE KO vs. DKO.
a

Indicates significant gender difference for that genotype.

Table 4 | Concentration of cholesterol-containing lipoproteins in plasma from ApoE KO or ApoE/Rag1 KO mice.
Male ApoE KO

Male DKO

Mean

SEM

Mean

Total

1360a

116.5

1215a

47.04

HDL

58.64

32.99

108.0

21.20

LDL +VLDL

931.3a

71.77

508.4

99.79

P-value (males)

SEM

Female ApoE KO
Mean

SEM

0.3939

802.4

0.2364

14.39

0.0063

504.4

Female DKO

P-value (females)

Mean

SEM

64.65

707.3

59.73

0.3050

3.177

85.86

41.76

0.0022

55.29

362.3

68.75

0.1384

Concentrations (milligrams per deciliter) are expressed as mean and SEM, n = 6. Bold type indicates P < 0.05 for ApoE KO vs. DKO.
a

Indicates significant gender difference for that genotype.

in BMDM (Figure 1B). These results revealed widely different
levels of expression and divergent patterns of regulation of MMPs
and TIMPs, which informed our choice of genes to measure in the
subsequent in vivo experiments (see below).
CIRCULATING INFLAMMATORY CYTOKINE AND CHOLESTEROL LEVELS
IN ApoE KO AND ApoE/Rag-1 DOUBLE-KNOCKOUT MICE

To investigate the impact of lymphocytes on M1/M2 polarization and MMP/TIMP expression, we compared ApoE KO and
ApoE/Rag-1 DKO mice, which were genotyped using PCR to verify the deletion of ApoE alone or ApoE and Rag-1. The weight
of the spleens of ApoE KO mice (115 ± 6 mg, n = 8) was reduced
to (56 ± 9 mg, n = 10) in DKO mice and histological analysis of
cut sections confirmed the absence of lymphocytes from DKO
spleens (not shown). As expected, the expression of CD3 mRNA
in the spleen, as a further marker of the presence of lymphocytes, was measurable in all ApoE KO mice tested (mean value
1.6 ± 0.5 copies/ng RNA, n = 10) but undetectable in all DKO mice
tested. We used blood levels of cytokines related to Th1 and Th2
lymphocytes to further characterize the inflammatory status of
ApoE KO and DKO mice. As in Table 3, circulating levels of IFNγ
were low and not different between genotypes, whereas IL-4 levels

Frontiers in Immunology | Inflammation

were below the limits of detection of our assays. Of the more
abundant cytokines, TNFα levels were not different, whereas GMCSF, IL-12, and IL-10 levels were elevated (1.6, 2.2, and 2.5-fold,
respectively) in female DKO compared to ApoE KO mice. The
same trend was seen for IL-10 levels in male mice (Table 3). These
results showed, unexpectedly, that cytokines associated with M1
polarization (i.e., GM-CSF and IL-12) and M2 polarization (IL10) were at least the same or even elevated in DKO mice. We
also noted that TNF-α levels were significantly 1.8-fold higher in
male than female ApoE KO mice but not different in DKO mice
(Table 3).
Plasma cholesterol levels play a fundamental role in determining the extent of atherosclerosis in human beings and animal
models. As shown in Table 4, the plasma concentrations of total
cholesterol were not different between ApoE KO and DKO mice.
HDL cholesterol was significantly increased sixfold in female DKO
vs. ApoE KO mice and there was the same trend (1.9-fold) in
the male mice. Conversely, LDL plus VLDL concentrations were
significantly 1.8-fold lower in male DKO than ApoE KO mice
and same trend (1.4-fold) was seen for the females. We noted
that total cholesterol levels were significantly higher in males
than females of either genotype. Furthermore, VLDL + LDL levels

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Foamy macrophage activation without lymphocytes

were significantly higher in the male than female ApoE KO mice
(Table 4). Based on increased HDL and decreased VLDL + LDL,
DKO mice of either gender might be expected to be protected from
atherosclerosis compared with ApoE KO mice.
M1/M2 MARKER AND MMP mRNA LEVELS IN GRANULOMA FCMs
FROM SUBCUTANEOUS SPONGES

We sought a ready source of foam cells generated in vivo to
investigate expression of the M1/M2 markers and MMPs and
TIMPs measured in our in vitro study of non-foamy macrophages.
Atherosclerotic plaques are small and difficult to disrupt but
hypercholesterolemia promotes the accumulation of foam cells
at several more accessible sites in human beings and mice. For
example, foam cells accumulate in the peritoneum (28) or in granulomas that form in sterile sponges implanted subcutaneously into
atherosclerosis-prone mice (23). In this study, FCMs were isolated
from subcutaneous granulomas. They were purified based on their
decreased buoyant density by flotation over a density gradient.
The yield of foam cells from subcutaneous sponges implanted for
6 weeks into ApoE KO or DKO mice was 4.07 ± 0.61 × 106 and
5.87 ± 1.21 × 106 cells, respectively, and did not significantly vary
between genders. Foam cells from ApoE KO mice were 95.5 ± 1.6%
macrophages (using MOMA-2 as a marker). FCMs had detectable
levels of the same M1 and M2 markers seen in non-foamy BMDM,
irrespective of whether they came from ApoE KO or DKO mice

(Table 5). We concluded that FCMs acquired M1 marker genes
in the absence of T and B lymphocytes. Indeed, there was a trend
toward higher levels of mRNA expression of M1 markers, iNOS
and COX-2, in DKO animals, although this was not significant
(Table 5). Given this somewhat surprising conclusion, we measured additional M1 markers, namely IL-12 p35 and p40 and
SOCS3, which were also detectable and showed no significant difference between ApoE-KO and DKO mice (Table 5). The mRNAs
for M2 markers arg-1, and Ym-1 were also expressed at similar
levels in granuloma FCMs from ApoE KO or DKO mice (Table 5),
whereas CD206 was slightly elevated in male DKO compared to
ApoE KO mice. The data suggested that M2 polarization also
occurred efficiently in the absence of lymphocytes. To confirm this,
we added measurements of FIZZ1 and IL-10, which also showed
no difference between ApoE KO and DKO FCMs (Table 5). We
noted a few significant gender differences. The mRNA levels of
the M1 marker, IL-12p40, and the M2 markers, CD206 and Ym-1,
were approximately 50% lower in granuloma FCMs from male
compared to female ApoE KO mice.
The MMPs that were increased by classical activation in blood
or BMDM, that is MMP-2, MMP-9, MMP-13, MMP-14, and
MMP-25, were all expressed in granuloma FCMs, irrespective of
Rag-1 genotype (Table 5). TIMP-2 (that was decreased by classical
activation) and MMP-19 (that was decreased by classical activation and increased by alternative activation) were also expressed

Table 5 | Characteristics of foam cell macrophages obtained from subcutaneous sponges.
Male KO

Male DKO

P-value (males)

Female ApoE

Mean

SEM

Mean

SEM

COX2

915

135

2053

729

iNOS

56

7

474

305

IL-12p35

1085

88

904

IL-12p40

6a

1

9

SOCS3

1797a

204

4117

1060

0.1191

2679

327

Arg-1

79646

5095

87511

7825

0.3953

87172

14259

CD206

1317a

198

2150

285

0.0224

2420

212

FIZZ1

377

58

518

82

0.1640

365

IL-10

10262

1115

10234a

780

0.9852

12984

Female DKO

P-value (females)

Mean

SEM

Mean

SEM

0.3692

931

134

1306

243

0.1981

0.3793

69

13

331

242

0.3357

130

0.2451

1563

299

1154

276

0.3316

3

0.7304

13

2

11

4

0.6432

3038

587

0.5896

90946

10539

0.8345

2293

372

0.7707

65

594

136

0.2810

2289

19852

4366

0.1853

Ym-1

35a

5

78

31

0.3311

103

20

124

29

0.5530

MMP2

387

77

614

136

0.1462

535

97

681

173

0.4586

MMP9

104a

18

217

63

0.0720

182

22

155

14

0.3537

MMP12

176264

11743

150381

15260

0.1860

215896

28916

160661

26809

0.1831

MMP13

43800

3821

33103

3990

0.0664

48821

8573

34538

4202

0.1770

MMP14

4888

570

5402

462

0.4963

5214

917

5856

942

0.6332

MMP19

8486

1684

7893

930

0.7723

8756

998

7814

1268

0.5699

MMP25

23

3

112

47

0.6049

27

6

84

42

0.9307

TIMP-1

1674a

237

2321

399

0.1625

2543

330

1771

401

0.1571

TIMP-2

6624a

533

6984

730

0.6884

10456

682

6455

606

0.0008

TIMP-3

1203a

160

2728

689

0.0352

2228

356

2452

586

0.7434

FCMs were isolated by density gradient centrifugation and differential adhesion from sponges implanted under the skin of ApoE KO or ApoE/Rag-1 DKO mice, as
indicated. Levels of mRNAs of M1/M2 markers and MMPs/TIMPs are expressed as copies/ng total RNA (mean and SEM, n = 7–13). Bold type indicates P < 0.05 for
ApoE KO vs DKO.
a

Indicates significant gender difference for that genotype.

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October 2014 | Volume 5 | Article 537 | 153

Hayes et al.

at high levels in both genotypes (Table 5). Female ApoE KO mice
had 1.6-fold more TIMP-2 mRNA than the corresponding males.
MMP-12 and TIMP-1 [that showed no relationship with classical/alternative phenotype under the conditions of our in vitro
studies (Figure 1B)] were expressed irrespective of genotype, but
TIMP-3 was significantly increased 2.3-fold in male DKO compared to ApoE KO mice. The proportion of granuloma FCMs
able to degrade gelatin was determined using in situ zymography. A significantly higher percentage of granuloma FCMs had
gelatinase activity from female DKO (91%) compared with ApoE
KO animals (76%), and the same trend was evident in the in the
males (86 vs. 74%) (Figure 2A), Pooling the data, the 88% of
the DKO mice had gelatinase activity compared to 75% in the
ApoE KO mice (P = 0.0026). The proliferative capacity of FCMs
was assessed by measuring BrdU incorporation (Figure 2B). Proliferation was not different in male mice of either genotype but
was increased almost twofold in female ApoE KO mice compared
to DKO mice (P < 0.0001). Migration through a matrigel layer
in a modified Boyden chamber assay (n = 3–5) was not significantly different amongst granuloma FCMs from the two genotypes
(results not shown).
M1/M2 MARKER PROTEIN EXPRESSION AND PATHWAY ACTIVATION IN
GRANULOMA FCMs FROM SUBCUTANEOUS SPONGES

We used immunohistochemistry to confirm the protein expression of M1 and M2 markers in vivo by using sections taken
from excised subcutaneous sponges. A subpopulation of granuloma cells in sponge sections stained for iNOS (Figures 3A,B) or
COX-2 protein (Figures 3C,D) in either Rag-1 genotype. Control
sections stained with isotype matched non-immune immunoglobulins had no staining (Figures 3E,F). Furthermore, a fraction of
the granuloma cells in sponge sections stained for nuclear localized
NF-κB (Figures 4A,B) or phosphorylated STAT-1 (Figures 4C,D),
indicating that these cells had undergone activation of the signaling pathways that are associated with M1 activation. Sections
stained with isotype matched non-immune immunoglobulins had
no staining (Figures 4E,F). Some FCMs in sections also stained for
arg-1 (Figures 5A,B), Ym-1 (Figures 5C,D), or phosphorylated
STAT-6 (Figures 5E,F), demonstrating the presence of marker

FIGURE 2 | Characteristics of foam cell macrophages (FCMs) obtained
from subcutaneous granulomas. Granuloma FCMs were isolated by
flotation over a density gradient and differential adhesion from sponges
implanted under the skin of ApoE KO or ApoE/Rag-1 DKO mice, as

Frontiers in Immunology | Inflammation

Foamy macrophage activation without lymphocytes

proteins and active signaling pathways that are associated with
M2 macrophages. Sections stained with isotype matched nonimmune immunoglobulins again had no staining (Figures 5G,H).
Similar results to those with sponge sections were found by
immunocytochemistry of FCM isolated from sponges (data not
shown).
PLAQUE SIZE, COMPOSITION, M1/M2 MARKERS, AND MMP PROTEIN
EXPRESSION IN FCMs IN ATHEROSCLEROTIC PLAQUES FROM ApoE KO
AND DKO MICE

Plaques in the aortic sinus tended to be smaller in DKO compared
with ApoE KO mice but this did not reach statistical significance
(Figure 6A). However, in agreement with previous reports (18,
19), significant gender differences were observed (Figures 6A,B).
Aortic sinus plaques were significantly bigger in females compared with males in ApoE KO (twofold: P = 0.0057) and DKO
(2.5-fold: P = 0.0043) mice. Plaques in the BCA were 2.1 times
smaller in the DKO compared to ApoE KO males (P = 0.004)
but there was no significant difference between the two genotypes in the females (Figure 6B). Interestingly, ApoE KO males had
2.9 times larger plaques when compared with ApoE KO females
(P < 0.0001) (Figure 6B), which was the opposite of the relationship observed in the aortic sinus. This difference has also been
noted before (29). These differences reinforced our decision to
stratify our data by gender.
Buried fibrous layers in BCA plaques have been suggested as
a marker of plaque complexity or instability (30). However, we
observed no linear regression between the number of buried layers and plaque size (P-values range from 0.20 to 0.68, n = 15–17) or
macrophage content (P-values range from 0.29 to 0.99, n = 15–17),
which are other measures that have been associated with plaque
vulnerability. In any event, there were no significant differences in
the number of buried layers in plaques from DKO compared to
ApoE KO mice (Figure 6C). The area occupied by plaques in the
aorta (as demonstrated by en face staining with Oil-Red-O) gives
another measure of the extent of plaque progression in these mice.
This did not change with phenotype or gender (Figure 6D).
The comparable plaque size in the aortic root of ApoE KO and
DKO mice has been reported previously (20), and was consistent

indicated. (A) In situ zymography (gelatinase activity, mean and SEM,
n = 9–13). (B) Proliferative capacity (8 h BrdU pulse, mean and SEM,
n = 8–14). *P < 0.05 vs. DKO, $ P < 0.05 indicates gender differences for
that genotype.

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Foamy macrophage activation without lymphocytes

FIGURE 3 | M1 marker staining of sponge sections from ApoE KO or
ApoE/Rag-1 DKO mice. Sections from ApoE KO (A,C,E) or DKO mice
(B,D,F) were stained to show immunolocalization of inducible nitric oxide
synthase (iNOS) (A,B) or cyclooxygenase-2 (COX-2) (C,D). Control sections
(E,F) were exposed to non-immune IgG. Positive staining appears brown
(DAB), nuclei blue-purple (hematoxylin), and sponge spicules light blue.
Sections from either mouse strain could contain cells with positive
cytoplasmic staining (arrowheads). Some cells did not stain, and some
appeared to have COX-2 in their nucleus (arrows) (Magnification: ×600).

FIGURE 4 | Signal pathway activation in sponge granuloma sections
from ApoE KO or ApoE/Rag-1 DKO mice. Sections from ApoE KO
(A,C,E) or DKO (B,D,F) mice were stained to show immunolocalization of
NF-κB (A,B) or phosphorylated STAT1 (C,D). Control sections (E,F) were
exposed to non-immune IgG. Positive staining appears brown (DAB), nuclei
blue-purple (hematoxylin), and sponge spicules light blue. Sections from
either mouse strain contained many cells with positive cytoplasmic staining
(arrowheads). Some cells did not contain detectable levels of NF-κB or
phospho-STAT-1, while others appeared to have NF-κB or phospho-STAT-1
present in their nucleus (arrows) (Magnification: ×600).

with the objective of our experimental design. It ensured that any
difference in foam cell phenotype in the aortic sinus would be
independent of plaque size. However, the larger plaque size in
male BCA plaques in ApoE KO than DKO mice could complicate
the interpretation of data relating to phenotypes in these mice.

Immunohistochemistry for α-smooth-muscle-cell-actin (αSM-actin) was used to quantify the presence of vascular SMC
and staining for GSL to quantify macrophages. As expected
SMC were mainly found in the media and fibrous cap of
plaques (Figures 7A–D). Most GSL positive had a foamy

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Hayes et al.

FIGURE 5 | M2 marker staining and signal pathway activity in
sponge granuloma sections from ApoE KO or ApoE/Rag-1 DKO
mice. Sections from ApoE KO (A,C,E) or DKO (B,D,F) mice were stained
to show immunolocalization of arginase-1 (A,B), Ym-1 (C,D), or
phosphorylated STAT6 (E,F). Control sections from ApoE KO (G) or DKO
(H) mice were exposed to non-immune IgG. Positive staining appears

appearance on close examination and were therefore mainly FCMs
(Figures 7A–D). However, some medial cells, presumably synthetic state SMC or SMC transdifferentiating toward macrophages
(31) also stained with GSL (Figures 7A–D). Given the small size
of atherosclerotic plaques in the AS and BCA, it was impractical to extract mRNA for qRT-PCR, or total protein for Western blotting. We therefore used immunohistological methods
to quantify the presence of M1 and M2 markers as well as
selected MMPs and TIMP-3. Our in vitro studies together with
the availability of suitable antibodies (Table 2) guided our choice
of iNOS as suitable M1 marker and arg-1 and Ym-1 as suitable M2 markers. Based again on our in vitro studies and the
availability of suitable antibodies (Table 2), we chose to study
MMP-13 and MMP-14 as potentially related to classical activation. MMP-12 and TIMP-3 were also chosen for comparison because they are abundantly expressed in vitro, irrespective
of classical and alternative activation. For each of the antibodies and lectin used the staining was specific, both in the aortic
sinus (Figures 7A,B) and the BCA (Figures 7C,D) of either
genotype. Interestingly, phenotypic markers, MMPs and TIMP-3

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Foamy macrophage activation without lymphocytes

brown (DAB), nuclei blue-purple (hematoxylin), and sponge spicules light
blue. Sections from either mouse strain contained cells with positive
cytoplasmic staining (arrowheads), but some cells did not contain
detectible levels of arginase-1, Ym-1, or phospho-STAT-6. Some cells
were observed with nuclear phospho-STAT-6 staining (arrows)
(Magnification: ×600).

were mainly associated with GSL-positive areas rather than αactin (Figures 7A–D). The percentage of the total plaque area
stained with each antibody was measured using image analysis. By confining measurements to the plaque, we avoided any
influence of staining from the media layer. Furthermore, some
of the antibodies stained cardiac myocytes surrounding the aortic root (Figures 7A,B) but this did not distort our subsequent measurements because these areas were excluded from the
quantification.
The area stained with α-SM actin in the aortic sinus (Figure 8A)
was less than 20% under all conditions, consistent with the lipidrich nature of plaques in this model at this time point. Nevertheless, SMC area was 1.5 times higher in the female DKO compared
with ApoE KO mice (P = 0.0051) and the same trend was seen in
the male mice. The pooled data for both genders were also significant (P = 0.0058) There were no differences between genotypes
in the BCA (Figure 8B), although male DKO mice had 3.8-times
more SMC staining than females (Figure 8B). Plaque areas stained
with GSL were 30–40% in AS or BCA, consistent with highly
inflamed nature of these plaques. GSL areas in the AS (Figure 8A)

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FIGURE 6 | Histological appearances of aortic sinus, brachiocephalic
artery and aortic plaques in ApoE KO and ApoE/Rag-1 DKO mice. (A) Area
of plaque in the aortic sinus. The first section showing leaflet of the aortic
valve was stained using Miller’s elastin/van Gieson (EVG) and the area of
plaque in each section calculated using computer aided planimetry,
n = 16–22. (B) A section taken 3 µm after the bifurcation of the
brachiocephalic artery (BCA) from the aorta was treated and stained and

and BCA (Figure 8B) did not significantly differ in the two mouse
genotypes or genders.
Quantitative measurements of staining for iNOS, arg-1, or Ym1 showed that these areas were similar to or less extensive than
GSL (Figures 8A,B), consistent with the concept of their being
restricted to subpopulations of the GSL-positive cells (Figures 7A–
D). Approximately, the same proportion of cells stained for iNOS
and Ym-1, implying that approximately half of the FCMs had M1
or M2 markers, consistent with previous literature (12). Staining
for arg-1 was more extensive and appeared to overlap with that for
iNOS in some cells (Figures 7A–D and 8A,B). Staining for iNOS
tended to be higher in DKO than ApoE KO mice in the aortic sinus
(Figure 8A) and BCA (Figure 8B), although this was not significant. Based on this evidence, M1 activation of plaque FCMs occurs
and may even be increased in the absence of T and B lymphocytes,
irrespective of gender, and consistent with the PCR data on granuloma FCMs described above. Female DKO mice had 1.4-fold more
arg-1 staining than ApoE KO mice in the aortic sinus (Figure 8A)
and the same trend was seen in the BCA (Figure 8B). There were
no differences in arg-1 staining for male DKO and ApoE KO mice
at either site (Figures 8A,B). Staining for Ym-1 was also not different between genotypes in the aortic sinus (Figure 8A) or BCA
(Figure 8B). Based on this evidence, M2 activation of FCM was
present in the absence of T and B lymphocytes in mice of both
sexes, and may be increased in the DKO females.

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Foamy macrophage activation without lymphocytes

examined in a similar fashion to A, n = 13–19. (C) The number of buried layers
in plaque from BCA were assessed in EVG and α-SM-actin stained sections,
n = 11–14. (D) Whole aorta were opened longitudinally and stained en face for
the presence of neutral lipid using Oil-Red-O. The area of lipid-rich plaque in
each aorta was calculated using computer aided planimetry, n = 10–11. Values
are expressed mean and SEM. *P < 0.05 vs. DKO, $ P < 0.05 indicates gender
differences for that genotype.

Additional gender differences were also noted. Staining for
iNOS in the aortic sinus was 2.4 times greater in the male than
female mice of both ApoE KO and DKO mice (Figure 8A). Staining
for arg-1 was 2.1 and 1.4-fold higher in male compared with female
ApoE and DKO mice, respectively in the aortic sinus (Figure 8A)
although not the BCA (Figure 8B). Staining for Ym-1 was similar
between males and females of either genotype in the aortic sinus,
and in ApoE KO in the BCA (Figures 8A,B).
The areas stained for MMPs-12, -13, -14, and TIMP-3 were
extensive in the aortic root and BCA plaques, which shows that
these proteins are widely expressed in plaques (Figures 8A,B).
The area of staining for MMP-13 and MMP-14 was similar in
ApoE KO and DKO mice of either gender in the AS (Figure 8A)
or BCA (Figure 8B). Clearly, the absence of T and B lymphocytes had little impact on extent of MMP-13 or MMP-14
staining, consistent with the PCR data obtained from granuloma
FCMs. MMP-12, however, showed fourfold increased staining in
male DKO compared to ApoE KO mice (P = 0.0261), although
this was not replicated in the BCA (Figure 8B) or in female
mice. The areas of TIMP-3 staining in the BCA were in all
cases similar irrespective of genotype or gender (Figures 8A,B).
Male mice of either genotype had approximately twice as much
staining for MMP-13, MMP-14, or TIMP-3 than females in the
AS (Figure 8A), although there was no difference in the BCA
(Figure 8B).

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FIGURE 7 | Immunohistochemical staining for cell type, M1 and M2
markers and for MMPs and TIMPs. Near consecutive sections to the section
stained with elastin/van Gieson (EVG) were subjected to
immunohistochemistry for macrophages [Griffonia simplicifolia lectin II (GSL)],
smooth muscle cells (α-smooth muscle actin; actin), iNOS, arg-1, Ym-1,
MMP-12, MMP-13, MMP-14, and TIMP-3, using the antibodies detailed in

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Foamy macrophage activation without lymphocytes

Table 2. Controls were performed with non-immune IgG or normal serum
replacing the primary antibody. (A) Aortic sinus plaques from ApoE KO mice
(Magnification: ×4). (B) Aortic sinus plaques from ApoE/Rag1 DKO mice
(Magnification: ×4). (C) Brachiocephalic artery plaques from ApoE KO mice
(Magnification: ×10). (D) Brachiocephalic artery plaques from ApoE/Rag1
DKO mice (Magnification: ×10).

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Foamy macrophage activation without lymphocytes

FIGURE 8 | The proportion of staining for cell type, M1/M2 markers, MMPs, and TIMPs in plaques from ApoE KO or ApoE/Rag1 DKO mice. (A) Aortic
sinus plaques. (B) Brachiocephalic artery plaques. Values are expressed as mean and SEM. *P < 0.05 vs. DKO, $ P < 0.05 indicates gender differences for that
genotype, n = 11–19.

DISCUSSION
The clear conclusion from our experiments is that FCMs in
subcutaneous granulomas and atherosclerotic plaques acquired
markers of classical (M1) and alternative (M2) activation even in
the absence of lymphocytes. This conclusion was valid in male
and female mice, despite differences in the size and composition of plaques between the two genders. Furthermore, since
nuclear localized NF-κB, phospho-STAT-1 and phospho-STAT6 were detected in granuloma FCMs, activators of the pathways
leading to M1 and M2 polarization also existed in adequate quantities in the presence or absence of T or B lymphocytes. We also
observed that expression of several MMPs and TIMPs occurred
in granuloma and plaque FCMs in vivo independently of T and B
lymphocytes.
Recent work demonstrated that foam cell formation did not,
in itself, lead to M1 or M2 activation (28). By contrast, peritoneal
FCM generated in vivo were resistant to M1 activation, owing
to stimulation of the LXR pathway. Nevertheless, several studies
showed that there are FCMs in mouse atherosclerotic plaques that

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express both M1 and M2 markers either in situ (12, 13) or after
collagenase isolation (32). M1 and M2 markers were associated
with distinct cell populations even though they had overlapping
distributions in the intima of advanced mouse plaques (32). Our
histological observations confirmed these findings but showed,
in addition, that FCMs produced in the context of the foreign
body reaction caused by a polyurethane sponge implantation
also prominently expressed M1 and M2 markers. Granuloma
FCMs mimicked the gene expression pattern of plaque FCMs
and may therefore be useful as a more-easily isolated surrogate.
Functionally, iNOS and arg-1 appear to compete for substrate.
Indeed, in a previous study, we demonstrated this directly in
rabbit FCMs where down-regulation of arg-1 compared to nonfoamy macrophages led to decreased urea and increased nitrate
release (24).
In previous papers dealing with atherosclerosis, it has been
acknowledged that Th1- and Th2-lymphocyte related cytokines
can polarize macrophages toward M1 and M2 phenotypes,
respectively (11–13). This suggests that lymphocytes and hence,

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by implication, adaptive immunity plays an essential role in
macrophage polarization, albeit in concert with other factors (11).
On the contrary, our new data show clearly that M1/M2 polarization of FCMs can take place efficiently in the absence of T and
B cells. To do this, we recreated a previously-characterized cross
(20) between the well-established ApoE KO mouse and the equally
well-characterized Rag1 KO mouse. We obtained similar development of plaques in these mice as previously observed (20) and then
went on to make novel observations of macrophage phenotypes
in subcutaneous granulomas and plaques.
It is worth noting that the absolute copy numbers of the M1
and M2 marker mRNAs measured in granuloma FCMs were of
the same order of magnitude as those in measured in classical and
alternatively activated macrophages in vitro (compare Table 5 to
Figure 1A). Despite the obvious limitations of comparing data
across such different experimental conditions, it is hard to escape
the conclusion that the FCMs produced in vivo express M1 and M2
marker genes to a substantial degree, irrespective of the presence of
lymphocytes. Moreover, the canonical pathways of classical activation, NF-κB and STAT-1, and alternative activation, STAT-6, are
also triggered in T and B cell deficient mice. One possible explanation is that TLR-4 mediated induction of so-called interferon
response factors (IRFs, specifically IRF-3 and IRF-7) can lead to
the secretion of IFNα and IFNβ, thereby bypassing the requirement for IFNγ (9). Plaques contain several potential activators of
TLRs (33); and these could well be the sources of classical activation. In addition to TLR agonists, other stimulators of the NF-κB
pathway, including TNFα and IL-1, that are known to occur in
mouse atherosclerotic plaques (3) could also act as classical activators independently of IFNγ, as shown in many previous studies
(3, 14). The presence of these alternative mediators therefore provides a rationale for M1 activation in T and B cell depleted mice,
although additional experiments beyond the present scope would
be needed to identify the specific mediators. M2 polarization can
also occur in response to a variety of mediators, although activation of STAT-6 appears to indicate the mediation of IL-4 and/or
IL-13 in our mice even in the absence of lymphocytes. An additional, non-exclusive explanation for our findings is that there are
sources other than lymphocytes for the cytokines associated with
M1 and M2 activation in mice. For example, natural killer cells
were shown to be an active source of IFNγ in Rag-1 KO mice (34)
and could therefore account for the residual levels of IFNγ we
observed in the blood of ApoE/Rag-1 DKO mice (Table 3). Likewise, mast cells (35) and neutrophils (36) are plausible sources of
IL-4 and IL-13 in lymphocyte-depleted mice.
MMPs have been strongly implicated in the progression of atherosclerosis, and more particularly in ECM degradation as well
migration, proliferation and apoptosis of vascular cells (5, 37).
The results in Figure 1B showing that MMP-2, MMP-9, MMP-13,
MMP-14, and MMP-25 were up-regulated and TIMP-2 downregulated in mouse macrophages during classical activation are
consistent with previously reviewed data for MMP-9 and MMP13 (38). However, IFNγ had no effect on MMP or TIMP mRNA
expression in mouse macrophages at the 18-h time point we used
(Figures 1C,D), which was chosen to allow time for priming effects
to be observed. Up-regulation of MMP-19 and down-regulation
of MMP-14 were the only changes that we observed in response

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Foamy macrophage activation without lymphocytes

to IL-4 (Figure 1B). Furthermore, there was no difference in the
expression level of any of the MMPs or TIMPs in FCMs isolated
from sponge granulomas in ApoE KO and DKO mice. Despite
this, there was a small increase in the proportion of cells able
to degrade gelatin in DKO mice, which might be explained by the
trend toward increases in MMP-2 and MMP-9 mRNA expressions.
Further experiments would be needed to verify this. Recent work
has placed increased emphasis on macrophage proliferation in
mouse atherosclerosis (39). Specifically, it has been suggested that
proliferation, rather than recruitment, is the major factor leading
to accumulation of FCMs into atherosclerotic plaques of ApoE
null mice at early time points (40). We found a relative decrease in
FCMs from female DKO mice, but this did not appear to be associated with differences in M1/M2 polarization or MMP expression.
Turning to our immunohistochemical studies of atherosclerotic
plaques, few changes were noted in the extent of MMP-12, -13,
-14, or TIMP-3 staining. Only MMP-12 staining appeared to be
increased in male DKO mice in the AS but not BCA. Since, we
found no effect of classical or alternative activation on MMP-12
expression in vitro, this isolated observation might be explained
by another mediator such as GM-CSF, which has been shown
to up-regulate MMP-12 in several, previously reviewed studies
(38). However, the plasma cytokine levels (Table 3) provide no
corroboration for this contention.
During the course of our studies, we noted significant differences in cytokine and lipid levels, plaque sizes, and content of SMC
between male and female mice that obliged us to consider these
data separately. We found smaller AS plaques in male than female
ApoE KO mice, similar to what has been previously noted and
attributed to the effects of estrogens (19) and prostaglandins (41).
On the other hand, we found that males develop larger lesions
in the BCA, confirming what we previously published in thesis
form (29) and consonant with findings in the aorta at longer time
points (18). This is most likely related, at least in part, to the higher
total cholesterol and VLDL + LDL levels, we observed in male mice
(Table 4). The important fact to stress, however, is that acquisition
of M1 and M2 markers and expression of MMPs and TIMP was
independent of T and B lymphocytes, irrespective of the gender of
mice we analyzed.
Human atherosclerotic plaques also have prominent populations of FCMs that show M1 markers (42, 43) and have been
known for many years to have nuclear localized NF-κB (44). Consistent with our present results, work on cells isolated from human
atherosclerotic plaques, placed emphasis on innate immune mechanisms, by showing that TLR-2 activation plays an important role
in M1 polarization and MMP secretion (45). There are also foci of
FCMs in the intima that express M2 markers, which are distinct
from FCMs carrying M1 markers (11). Non-foamy macrophages
carrying M2 markers are even more prevalent in the adventitia (43). Hence, the distribution of cells carrying M2 markers in
human plaques appears to be more restricted than in the mouse
plaques in our study (Figures 7A–D) and in those previously
published (12, 13).
With respect to MMPs and TIMPs, comparison of our in vitro
results with published data from human macrophages isolated and
incubated under very similar conditions (38, 42, 46) demonstrates
many differences. For example, MMP-1 is absent but MMP-13

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is abundant in mouse macrophages, whereas MMP-13 is absent
and MMP-1 is abundant in human macrophages (42, 47), consistent with the limited distribution of MMP-13 in human tissues
(48). Furthermore, MMP-12 is apparently much more abundant
in mouse macrophages (Figure 1B) than human macrophages
(42). Conversely, mRNAs for MMP-2, MMP-3, MMP-7, MMP-9,
MMP-10, MMP-11, MMP-17, TIMP-1, and TIMP-3 appear much
less abundant in mouse macrophages (Figure 1B) than human
macrophages (42). Only MMP-8, MMP-14, MMP-19, MMP-25,
and TIMP-2 show similar (within 10-fold) abundance in both
species at the mRNA level. Responses of MMPs and TIMPs to
classical and alternative also present stark contrasts between mice
and men, under the conditions of our experiments. For example, expression of MMP-1, MMP-3, MMP-7, MMP-10, MMP12, and TIMP-1 was increased by classical activation of human
macrophages, whereas MMP-9 was constitutive (42), in contrast to what we observed here (Figure 1B). Furthermore, IFNγ
increased MMP-1, MMP-10, MMP-12, and MMP-14 expression
and decreased TIMP-3 expression in human macrophages (42),
none of which we observed in mouse macrophages at the same
time point (Figure 1C). Finally, IL-4 treatment increased MMP11, MMP-12, and TIMP-3 expression in human macrophages (42)
but MMP-19 expression in mice (Figure 1B). Similar disparity has
been previously noted with respect to the expression levels of M1
and M2 markers in human and mouse macrophages (15). Moreover, the overall transcriptomic response to several in vivo models
of inflammation appears highly divergent in mice and men (49).
These limitations therefore caution against over-extrapolating our
present results from a mouse model to human atherosclerosis.
In conclusion, our results definitively counter the hypothesis that lymphocytes are necessary for M1 or M2 polarization
in mouse atherosclerosis, although more work will be needed to
define the mediators responsible. Lymphocytes are also not needed
for MMP and TIMP expression in FCMs in vivo. However, our
studies do not rule out a modulatory role for T or B lymphocytes on either macrophage polarization or MMP production. It is
conceivable that deletion of different lymphocyte populations has
opposing effects of macrophage and foam cell activation, leading
to a neutral effect overall. Subsequent studies using more selective interventions will be needed to investigate the role of specific
lymphocyte subsets in mice.

ACKNOWLEDGMENTS
This work was supported by funding from by the British Heart
Foundation. The authors would like to thank Mrs. Kristina Baker
for her help with some tissue harvests and KWS BioTest Ltd. for
measuring the cytokine levels.

REFERENCES
1. Smith JD, Trogan E, Ginsberg M, Grigaux C, Tian J, Miyata M. Decreased
atherosclerosis in mice deficient in both macrophage colony-stimulating factor (op) and apolipoprotein E. Proc Natl Acad Sci U S A (1995) 92:8264–8.
doi:10.1073/pnas.92.18.8264
2. Paulson KE, Zhu SN, Chen M, Nurmohamed S, Jongstra-Bilen J, Cybulsky MI.
Resident intimal dendritic cells accumulate lipid and contribute to the initiation
of atherosclerosis. Circ Res (2010) 106:383–90. doi:10.1161/CIRCRESAHA.109.
210781
3. Tedgui A, Mallat Z. Cytokines in atherosclerosis: pathogenic and regulatory
pathways. Physiol Rev (2006) 86:515–81. doi:10.1152/physrev.00024.2005

www.frontiersin.org

Foamy macrophage activation without lymphocytes

4. Bennett M, Yu H, Clarke M. Signalling from dead cells drives inflammation
and vessel remodelling. Vascul Pharmacol (2012) 56:187–92. doi:10.1016/j.vph.
2012.01.006
5. Libby P. Collagenases and cracks in the plaque. J Clin Invest (2013) 123:3201–3.
doi:10.1172/JCI67526
6. Adams DO, Hamilton TA. The cell biology of macrophage activation. Annu Rev
Immunol (1984) 2:283–318. doi:10.1146/annurev.iy.02.040184.001435
7. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8:958–69. doi:10.1038/nri2448
8. Martinez FO, Gordon S, Locati M, Mantovani A. Transcriptional profiling
of the human monocyte-to-macrophage differentiation and polarization: new
molecules and patterns of gene expression. J Immunol (2006) 177:7303–11.
doi:10.4049/jimmunol.177.10.7303
9. Hu X, Chakravarty SD, Ivashkiv LB. Regulation of interferon and toll-like receptor signaling during macrophage activation by opposing feedforward and feedback inhibition mechanisms. Immunol Rev (2008) 226:41–56. doi:10.1111/j.
1600-065X.2008.00707.x
10. Platanias LC. Mechanisms of type-I- and type-II-interferon-mediated signalling. Nat Rev Immunol (2005) 5:375–86. doi:10.1038/nri1604
11. Bouhlel MA, Derudas B, Rigamonti E, Dievart R, Brozek J, Haulon S,
et al. PPARgamma activation primes human monocytes into alternative M2
macrophages with anti-inflammatory properties. Cell Metab (2007) 6:137–43.
doi:10.1016/j.cmet.2007.06.010
12. Kadl A, Meher AK, Sharma PR, Lee MY, Doran AC, Johnstone SR, et al.
Identification of a novel macrophage phenotype that develops in response to
atherogenic phospholipids via Nrf2. Circ Res (2010) 107:737–46. doi:10.1161/
CIRCRESAHA.109.215715
13. Cardilo-Reis L, Gruber S, Schreier SM, Drechsler M, Papac-Milicevic N, Weber
C, et al. Interleukin-13 protects from atherosclerosis and modulates plaque
composition by skewing the macrophage phenotype. EMBO Mol Med (2012)
4:1072–86. doi:10.1002/emmm.201201374
14. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine
system in diverse forms of macrophage activation and polarization. Trends
Immunol (2004) 25:677–86. doi:10.1016/j.it.2004.09.015
15. Martinez FO, Helming L, Gordon S. Alternative activation of macrophages:
an immunologic functional perspective. Annu Rev Immunol (2009) 27:451–83.
doi:10.1146/annurev.immunol.021908.132532
16. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages and
the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.4049/jimmunol.
164.12.6166
17. Libby P, Lichtman AH, Hansson GK. Immune effector mechanisms implicated in atherosclerosis: from mice to humans. Immunity (2013) 38:1092–104.
doi:10.1016/j.immuni.2013.06.009
18. Tangirala RK, Rubin EM, Palinski W. Quantitation of atherosclerosis in
murine models: correlation between lesions in the aortic origin and in the
entire aorta, and differences in the extent of lesions between sexes in LDL
receptor-deficient and apolipoprotein E-deficient mice. J Lipid Res (1995) 36:
2320–8.
19. Caligiuri G, Nicoletti A, Zhou X, Tornberg I, Hansson GK. Effects of sex and
age on atherosclerosis and autoimmunity in apoE-deficient mice. Atherosclerosis
(1999) 145:301–8. doi:10.1016/S0021-9150(99)00081-7
20. Daugherty A, Pure E, Delfel-Butteiger D, Chen S, Leferovich J, Roselaar SE,
et al. The effects of total lymphocyte deficiency on the extent of atherosclerosis in apolipoprotein E−/− mice. J Clin Invest (1997) 100:1575–80.
doi:10.1172/JCI119681
21. Di Gregoli K, Jenkins N, Salter R, White S, Newby AC, Johnson JL. MicroRNA-24
regulates macrophage behavior and retards atherosclerosis. Arterioscler Thromb
Vasc Biol (2014) 34:1990–2000. doi:10.1161/ATVBAHA.114.304088
22. Chase A, Bond M, Crook MF, Newby AC. Role of nuclear factor-kB activation
in metalloproteinase-1, -3 and -9 secretion by human macrophages in vitro
and rabbit foam cells produced in vivo. Arterioscler Thromb Vasc Biol (2002)
22:765–71. doi:10.1161/01.ATV.0000015078.09208.92
23. Johnson JL, Devel L, Czarny B, George SJ, Jackson CL, Rogakos V, et al. A selective matrix metalloproteinase-12 inhibitor retards atherosclerotic plaque development in apolipoprotein E-knockout aice. Arterioscler Thromb Vasc Biol (2011)
31:528–35. doi:10.1161/ATVBAHA.110.219147
24. Thomas AC, Sala-Newby GB, Ismail Y, Johnson JL, Pasterkamp G, Newby
AC. Genomics of foam cells and nonfoamy macrophages from rabbits
identifies arginase-I as a differential regulator of nitric oxide production.

October 2014 | Volume 5 | Article 537 | 161

Hayes et al.

25.

26.

27.

28.

29.
30.

31.

32.

33.
34.

35.

36.
37.
38.

39.

40.

41.

Arterioscler Thromb Vasc Biol (2007) 27:571–7. doi:10.1161/01.ATV.0000256470.
23842.94
Hadler-Olsen E, Kanapathippillai P, Berg E, Svineng G, Winberg JO, UhlinHansen L. Gelatin in situ zymography on fixed, paraffin-embedded tissue: zinc
and ethanol fixation preserve enzyme activity. J Histochem Cytochem (2010)
58:29–39. doi:10.1369/jhc.2009.954354
Paigen B, Ishida BY, Verstuyft J, Winters RB, Albee D. Atherosclerosis
susceptibility differences among progenitors of recombinant inbred strains
of mice. Arterioscler Thromb Vasc Biol (1990) 10:316–23. doi:10.1161/01.ATV.
10.2.316
Newby AC, George SJ, Ismail Y, Johnson JL, Sala-Newby GB, Thomas AC. Vulnerable atherosclerotic plaque metalloproteinases and foam cell phenotypes.
Thromb Haemost (2009) 101:1006–11. doi:10.1160/TH08-07-0469
Spann NJ, Garmire LX, Mcdonald JG, Myers DS, Milne SB, Shibata N, et al. Regulated accumulation of desmosterol integrates macrophage lipid metabolism and
inflammatory responses. Cell (2012) 151:138–52. doi:10.1016/j.cell.2012.06.054
Johnson JL. The Role of Matrix Metalloproteinases in an Animal Model of Atherosclerotic Plaque Rupture. PhD. Bristol: University of Bristol (2005).
Johnson JL, Carson K, Williams HM, Karanam S, Newby AC, Angelini GD,
et al. Plaque rupture after short periods of fat-feeding in the apolipoprotein E knockout mouse: model characterisation, and effects of pravastatin
treatment. Circulation (2005) 111:1422–30. doi:10.1161/01.CIR.0000158435.
98035.8D
Rong JX, Shapiro M, Trogan E, Fisher EA. Transdifferentiation of mouse aortic
smooth muscle cells to a macrophage-like state after cholesterol loading. Proc
Natl Acad Sci U S A (2003) 100:13531–6. doi:10.1073/pnas.1735526100
Koltsova EK, Hedrick CC, Ley K. Myeloid cells in atherosclerosis: a delicate
balance of anti-inflammatory and proinflammatory mechanisms. Curr Opin
Lipidol (2013) 24:371–80. doi:10.1097/MOL.0b013e328363d298
Erridge C. The roles of toll-like receptors in atherosclerosis. J Innate Immun
(2009) 1:340–9. doi:10.1159/000191413
Thale C, Kiderlen AF. Sources of interferon-gamma (IFN-gamma) in
early immune response to listeria monocytogenes. Immunobiology (2005)
210:673–83. doi:10.1016/j.imbio.2005.07.003
Bradding P, Feather IH, Howarth PH, Mueller R, Roberts JA, Britten K, et al.
Interleukin 4 is localized to and released by human mast cells. J Exp Med (1992)
176:1381–6. doi:10.1084/jem.176.5.1381
Brandt E, Woerly G,Younes AB, Loiseau S, Capron M. IL-4 production by human
polymorphonuclear neutrophils. J Leukoc Biol (2000) 68:125–30.
Newby AC. Matrix metalloproteinase inhibition therapy for vascular diseases.
Vascl Pharmacol (2012) 56:232–44. doi:10.1016/j.vph.2012.01.007
Newby AC. Metalloproteinase expression in monocytes and macrophages and
its relationship to atherosclerotic plaque instability. Arterioscler Thromb Vasc Biol
(2008) 28:2108–14. doi:10.1161/ATVBAHA.108.173898
Andrés V, Pello OM, Silvestre-Roig C. Macrophage proliferation and apoptosis
in atherosclerosis. Current Opin Lipidol (2012) 23:429–38. doi:10.1097/MOL.
0b013e328357a379
Robbins CS, Hilgendorf I, Weber GF, Theurl I, Iwamoto Y, Figueiredo JL, et al.
Local proliferation dominates lesional macrophage accumulation in atherosclerosis. Nat Med (2013) 19:1166–72. doi:10.1038/nm.3258
Smith DD, Tan X, Tawfik O, Milne G, Stechschulte DJ, Dileepan KN. Increased
aortic atherosclerotic plaque development in female apolipoprotein E-null mice

Frontiers in Immunology | Inflammation

Foamy macrophage activation without lymphocytes

is associated with elevated thromboxane A2 and decreased prostacyclin production. J Physiol Pharmacol (2010) 61:309–16.
42. Huang WC, Sala-Newby GB, Susana A, Johnson JL, Newby AC. Classical
macrophage activation up-regulates several matrix metalloproteinases through
mitogen activated protein kinases and nuclear factor-kappaB. PLoS One (2012)
7:e42507. doi:10.1371/journal.pone.0042507
43. Stoger JL, Gijbels MJ,Van Der Velden S, Manca M,Van Der Loos CM, Biessen EA,
et al. Distribution of macrophage polarization markers in human atherosclerosis. Atherosclerosis (2012) 225:461–8. doi:10.1016/j.atherosclerosis.2012.09.013
44. Brand K, Page S, Rogler G, Bartsch A, Brandl R, Knuechel R, et al. Activated transcription factor nuclear factor-kappa B is present in the atherosclerotic lesion.
J Clin Invest (1996) 97:1715–22. doi:10.1172/JCI118598
45. Monaco C, Gregan SM, Navin TJ, Foxwell BM, Davies AH, Feldmann M. Tolllike receptor-2 mediates inflammation and matrix degradation in human atherosclerosis. Circulation (2009) 120:2462–9. doi:10.1161/CIRCULATIONAHA.
109.851881
46. Reel B, Sala-Newby GB, Huang W-C, Newby AC. Diverse patterns of
cyclooxygenase-independent metalloproteinase gene regulation in human
monocytes. Br J Pharmacol (2011) 163:1679–90. doi:10.1111/j.1476-5381.2011.
01298.x
47. Bar-Or A, Nuttall RK, Duddy M, Alter A, Kim HJ, Ifergan I, et al. Analyses of
all matrix metalloproteinase members in leukocytes emphasize monocytes as
major inflammatory mediators in multiple sclerosis. Brain (2003) 126:2738–49.
doi:10.1093/brain/awg285
48. Vincenti MP, Brinckerhoff CE. Transcriptional regulation of collagenase (MMP1, MMP-13) genes in arthritis: integration of complex signaling pathways for the
recruitment of gene-specific transcription factors. Arthritis Res (2002) 4:157–64.
doi:10.1186/ar401
49. Seok J, Warren HS, Cuenca AG, Mindrinos MN, Baker HV, Xu WH, et al.
Genomic responses in mouse models poorly mimic human inflammatory
diseases. Proc Natl Acad Sci U S A (2013) 110:3507–12. doi:10.1073/pnas.
1222878110
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 29 July 2014; accepted: 10 October 2014; published online: 28 October 2014.
Citation: Hayes EM, Tsaousi A, Di Gregoli K, Jenkinson SR, Bond AR, Johnson JL,
Bevan L, Thomas AC and Newby AC (2014) Classical and alternative activation
and metalloproteinase expression occurs in foam cell macrophages in male and female
ApoE null mice in the absence of T and B lymphocytes. Front. Immunol. 5:537. doi:
10.3389/fimmu.2014.00537
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Hayes, Tsaousi, Di Gregoli, Jenkinson, Bond, Johnson, Bevan,
Thomas and Newby. This is an open-access article distributed under the terms of
the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are
credited and that the original publication in this journal is cited, in accordance with
accepted academic practice. No use, distribution or reproduction is permitted which
does not comply with these terms.

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MINI REVIEW ARTICLE
published: 13 October 2014
doi: 10.3389/fimmu.2014.00490

Cholesterol efflux pathways regulate myelopoiesis: a
potential link to altered macrophage function in
atherosclerosis
Andrew James Murphy 1,2,3 *, Dragana Dragoljevic 1 and Alan Richard Tall 4
1
2
3
4

Haematopoiesis and Leukocyte Biology, Baker IDI, Melbourne, VIC, Australia
Department of Immunology, Monash University, Melbourne, VIC, Australia
University of New South Wales, Sydney, NSW, Australia
Division of Molecular Medicine, Department of Medicine, Columbia University Medical Center, New York, NY, USA

Edited by:
Klaus Ley, La Jolla Institute for Allergy
and Immunology, USA
Reviewed by:
Janos G. Filep, University of
Montreal, Canada
Clinton S. Robbins, University of
Toronto, Canada
Kevin Woollard, Imperial College
London, UK
*Correspondence:
Andrew James Murphy , Baker IDI
Heart and Diabetes Institute,
Melbourne, Australia
e-mail: andrew.murphy@
bakeridi.edu.au

Atherosclerotic cardiovascular disease is a chronic inflammatory disease of the blood vessels that can lead to myocardial infarction or stroke. The major cell in the atherosclerotic
lesion, the macrophage, is thought to be an important contributor to the production of
inflammatory mediators that exacerbate this disease. Macrophages are generally derived
from circulating monocytes, which are in turn produced by hematopoietic stem and multipotential progenitor cells (HSPCs) in the bone marrow and other medullary organs.
Recent studies suggest that disruption in cholesterol homeostasis or prolonged exposure
to a hypercholesterolemic environment can influence HSPCs to over-produce monocytes,
resulting in monocytosis. These monocytes may carry a pre-programed ability to become
M1-like macrophages once they enter the atherosclerotic lesion. Future studies may help
to differentiate the role of such pre-programing versus responses to local environmental
cues in determining M1, M2, or other macrophage phenotypes in atherosclerotic lesions.
Keywords: atherosclerosis, hematopoiesis, cholesterol efflux, monocytes, macrophages, HDL

Innate immunity has long been considered a driving force in
the initiation and progression of atherosclerotic cardiovascular
disease (CVD) (1). Indeed, inflammation is a process that has
attracted considerable attention as a potential therapeutic target
in atherosclerosis. It has also become well accepted that cholesterol metabolism is intimately linked to inflammation and innate
immune processes. This close relationship is not only important
in the effector cells of atherosclerotic disease such as monocytes
(2) and macrophages (3) but cholesterol metabolism has also been
shown to play a central role in their hematopoietic precursors (4,
5). This is important as changes in cholesterol homeostasis in the
hematopoietic stem and multipotential progenitor cells (HSPCs)
control the rate of production of monocytes/macrophages, and
possibly have an influence on their function (4–6). Increased numbers of circulating monocytes are a predictor of cardiovascular
risk (7–14) and studies in mice have shown a causal role (4–6,
15, 16). These key studies in mice have also revealed that HSPCs
can mobilize from the bone marrow (BM) to extramedullary sites
such as the spleen (4–6, 17), where they can also produce monocytes that contribute to atherogenesis (6). In this article, we will
review these topics and also explore the hypothesis that the mechanisms contributing to monocyte production from HSPCs could
also influence the type and function of lesional macrophages.

INNATE IMMUNE CELL PRODUCTION AND
CARDIOVASCULAR RISK
Monocytosis is associated with CVD and atherosclerotic plaque
severity in prospective and cross-sectional human studies (7, 11–
13). Monocytosis is also closely linked to plasma lipids, where

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a positive correlation is observed with total cholesterol levels
(15, 18–20), and an inverse correlation with plasma high-density
lipoprotein (HDL) levels (11, 14, 20, 21). Gerrity et al. first suggested that excessive monocyte production contributed to atherogenesis in rabbit and pig l models of hypercholesterolemia and
atherosclerosis (18, 19). These studies also made the link between
hypercholesterolemia and enhanced monocyte production from
the BM using colony-forming assays and suggested that this could
be driving the atherogenic phenotype (18). Through the use of
mouse models, a causal relationship between monocyte levels and
severity of atherosclerotic lesions has been shown (4, 5, 15). Studies employing the op/op mouse that carries a mutation in the gene
encoding macrophage colony-stimulating factor (CSF-1; M-CSF)
have a gene dose-dependent decrease in monocyte levels that is
reflected by smaller atherosclerotic lesions (22). Conversely, western diet (WTD)-fed Apoe−/− mice display monocytosis that is
proportionate to the length of feeding and reflects the size of the
atherosclerotic lesion (15). We have also shown that monocytosis, largely independent of activation, accelerates atherosclerosis
in mouse models (4). In addition to the abundance of monocytes
that circulate, the site of production may play an important role,
as monocytes produced in the spleen appear to have an atherogenic phenotype (6). While monocytes may directly contribute
to atherogenesis by secreting inflammatory cytokines, ROS, and
proteases, their most important role is probably to act as precursors lesional macrophages. Macrophages are a heterogeneous
population of cells and have been categorized into two main
groups known as M1 and M2. This classification is based on function and the expression on a number of genes. M1 macrophages

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are thought to be inflammatory cells, expressing a gene signature including iNos, IL-6, Tnf-α, and IL-1β, while M2 cells are
thought to play a resolving role and expression genes such as IL10, Tgf-β, and Arg1. However, these cells due retain plasticity and
can sit at various points along the scale [see recent reviews on
suggested nomenclature (23, 24)]. In Apoe −/− mice, CCR2+ Ly6Chi monocytes preferentially enter the atherosclerotic lesion (15,
16), and this monocyte subset has been suggested to differentiate
into a macrophage with an inflammatory phenotype. Interestingly,
lesional macrophages can also undergo local proliferation to sustain their population within the advanced atherosclerotic plaque
(25). The phenotype of proliferating macrophage or its product
cells has not yet been studied in detail; however, as the proliferation of these cells is dependent on SR-A (25), it could perhaps be
of the M2 variety (26). Below we will discuss how defects in cholesterol metabolism pathways influence the HSPCs, monocytes,
and macrophages to promote atherosclerosis, and will make the
speculative suggestion that events in the hematopoietic stem and
progenitor populations may influence the ultimate functions of
the macrophage.

CHOLESTEROL EFFLUX PATHWAYS LINK HSPC
PROLIFERATION, MONOCYTE PRODUCTION, AND
ATHEROSCLEROSIS
Impaired cholesterol efflux has long been associated with atherosclerosis, and more recently, the ability of HDL to promote efflux
from cholesterol loaded cells was shown to be a stronger predictor
or atherosclerotic burden than HDL cholesterol or apoA-I levels
(27). In line with this is the experimental evidence in animal models of atherosclerosis where increasing HDL levels either therapeutically (rHDL infusions) (28) or genetically (ApoA-I transgene) (5,
29) is protective. This is thought to be due to the ability of HDL or
ApoA-I to prevent foam cell formation, inhibit leukocyte adhesion,
and protect the endothelium from activation (30, 31). However,
recent studies have shown that HDL via cholesterol removal from
the cell membrane can regulate the production of innate immune
cells (4, 5, 32), particularly monocytes, by acting on HSPCs (4,
5). In respect to the anti-atherogenic properties of HDL, this may
be an important function that could affect the types and/or functions of the downstream cells that eventually mature into lesional
macrophages.
The removal of cholesterol from HSPCs can be facilitated
by a number of pathways. We discovered that HSPCs express
Abca1, Abcg1, and Apoe at high levels and these key efflux genes
could further be induced in vivo by the administration of LiverX-Receptor (LXR) agonists (4). Co-deletion of two key cholesterol efflux genes ATP bind cassette transporter (ABC) A1 and
Abcg1, in the hematopoietic compartment and transplantation
into Ldlr +/− mice resulted in prominent monocytosis and neutrophilia, which was accompanied by a dramatic acceleration in
atherosclerotic lesion formation (5). A myeloproliferative phenotype was suggested, as myeloid cells infiltrated many major organs,
including the spleen, liver, and intestine. Mice with Abca1/g1 KO
BM had a dramatic expansion of the HSPCs, which were proliferating at higher rates compared to mice that received WT
BM. The enhanced proliferation in the Abca1/g1 KO HSPCs was
found to be due to an increase in the expression of the common

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Cholesterol efflux, myelopoiesis, and macrophages

β subunit of the IL-3/GM-CSF receptor (IL-3Rβ; aka CD131),
making these cells more sensitive to these cytokines. Promoting
cholesterol efflux with an apoA-I transgene reversed the proliferative defects and reduced the severity of the atherosclerosis.
Abca1−/− , Abcg1−/− , and Apoe−/− HSPCs also mobilized into the
circulation in increased amounts and established extramedullary
hematopoiesis in the spleen and other organs (17). These sites
of extramedullary hematopoiesis provide an important reservoir
for monocytes in acute coronary disease (33), highlighting the
multiple links between hypercholesterolemia, defective cholesterol efflux pathways, and the over-production of monocytes and
neutrophils that contribute to atherosclerosis.
As mentioned above, we and others have also reported that
WTD-fed Apoe−/− mice display prominent monocytosis (4, 15,
16). We found that this was also due to expansion and proliferation of the HSPCs as a result of increased expression of the IL-3Rβ.
Treating Apoe−/− mice with reconstituted HDL (rHDL; CSL-111)
to promote cholesterol efflux normalized this proliferative defect
(4). The role for the IL-3Rβ in promoting HSPC proliferation
and monocytosis in Apoe−/− mice was confirmed in mice with
deficiency of both genes (34). Through the use of competitive
BM transplant (cBMT) studies, we found that these efflux pathways at least partly functioned in a cell intrinsic manner (4). For
example, deletion of Apoe in cells marked by CD45.2 produced
more monocytes and lesional macrophages compared to WT cells
marked by CD45.1 that were transplanted into the same recipients. We also found that the Ldlr−/− mice that received the mix
of Apoe−/− (CD45.2)/WT(CD45.1) had larger lesions compared
to those that received WT(CD45.2)/WT(CD45.1). The increase in
lesion size was independent of monocyte activation and supports
the idea that increased production of monocytes directly impacts
lesion monocyte/macrophage content, size, and severity. However,
we speculate that other explanations may be involved, including
that increased entry of Apoe−/− monocytes results in macrophages
that have an altered phenotype/function, or that alterations in
cholesterol metabolism in HSPCs pre-program their daughter
cells (i.e., monocytes and macrophages) into an inflammatory
phenotype (Figure 1).
To further explore the contribution of cholesterol efflux in
macrophages versus HSPCs, cell specific knockouts of Abca1/Abcg1
have been employed to examine the role of these transporters
in cells down stream of HSPCs (35). Using the Lysozyme M
Cre mouse crossed with Abca1flox/flox Abcg1flox/flox mice (MacDKO), Westerterp et al. (35) were able to reduce the expression of Abca1/Abcg1 by approximately half in the granulocytemacrophage progenitors (GMPs) and monocytes, and by ~90% in
macrophages. Transplantation of the MacABCDKO BM into Ldlr−/−
mice resulted in a ~1.7-fold increase in atherosclerosis plaque
area compared to the mice that received the control BM. However, the lesions in the mice that received Abca1/g1 KO BM has
significantly larger lesions (~3-fold) compared to control and
MacABCDKO mice (~1.7-fold). This implies that there is a major
effect of cholesterol efflux in cells upstream of macrophages in
controlling atherosclerosis, likely HSPCs.
The studies of Westerterp et al., in the MacABCDKO mice also
provided a novel insight on effects of altered cholesterol homeostasis in regulating the production of monocytes from the BM.

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FIGURE 1 | How alterations in cholesterol metabolism and myeloid
skewing contribute to atherosclerosis. In the setting of
hypercholesterolemia, inflammatory signals could be sensed by
receptors such as TLR4 on HSPCs to trigger a number of downstream
signaling events. This could (1) inhibit key cholesterol efflux pathways
(ABCA1, ABCG1, ApoE), which would result in cellular accumulation. The
increase in membrane cholesterol could lead to (2) increased cell surface
expression of cytokine receptors such as IL-3Rβ and the M-CSFR due to
a failure to activate E3-ubiquitin ligases (E3-UL). (3) Sustained signaling
from myeloid cytokines (IL-3, GM-CSF, M-CSF) along with the
hypercholesterolemic environment could pre-program the HSPC via PU.1

Macrophages deficient in Abca1/Abcg1 produced more M-CSF,
G-CSF, and MCP-1, key cytokines involved in myelopoiesis and
monocyte emigration from the BM (36). Interestingly, there
was also a significant number of lipid-laden foam cells in the
BM and spleen of the MacABCDKO mice, probably representing BM monocytes and macrophages. Consistent with this idea,
there was a significant portion of the blood monocytes from
the MacABCDKO mice that were loaded with cholesterol. Thus,
deletion of Abca1/Abcg1 in hematopoietic progenitors (i.e., GMPs)

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Cholesterol efflux, myelopoiesis, and macrophages

to produce more myeloid cells. As these cells mature in to CMPs and
GMPs, they have the potential to carry more cellular cholesterol if their
cholesterol efflux pathways are suppressed. (4) Once the blood
monocyte is circulating, more lipid is acquired and it can carry this into
the atherosclerotic plaque. (5) These lipid-laden monocytes could then
differentiate into an M1-like macrophage that can also undergo local
proliferation, which enhance inflammation by producing a number of
cytokines and chemokines. These M-1 cells may also have a defect in
Nr4a1 and lack the ability to convert into M-2 resolving macrophages.
Triangle indicates cellular cholesterol accumulation as the myeloid cells
mature (orange to yellow).

could predispose these and their daughter cells (monocytes) to
accumulate cholesterol. This could result in monocytes carrying
lipid into the atheroma and perhaps facilitating their maturation
into M1 macrophages as the macrophages from the MacABCDKO
mice also displayed enhanced inflammatory gene expression
(Figure 1). Additionally, in vitro migration studies revealed a
severe migratory defect of cultured macrophages deficient in
Abca1/Abcg1 (37), how this translates into the in vivo setting is
unknown.

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Murphy et al.

MONOCYTE TO MACROPHAGE DIFFERENTIATION
The origin of the macrophage itself is not always from a blood
monocyte (38). Yolk-sac derived tissue macrophages can sustain
their population under steady-state conditions, without recruitment of blood monocytes (39–41). After an inflammatory insult,
blood monocytes can be recruited to increase the macrophage pool
and to enhance the inflammatory response (39–41). In the heart,
an organ with abundant resident macrophages that are established during embryonic development, it was found that CCR2+
Ly6-Chi monocyte-derived macrophages coordinate the inflammatory response after cardiac injury by AngII infusion (39) or
myocardial infarction (MI) (39). These Ly6-Chi monocytes express
Nr4a1, a transcription factor critical in the development of Ly6Clo monocytes (42), at low levels (43). However, in the healing
phase after a MI, Nr4a1 (Nur77) levels are increased permitting
the maturation and differentiation of Ly6-Chi monocytes into Ly6Clo monocyte/macrophages (43). These macrophages contribute
to healing and tissue remodeling by producing factors such as
TGF-β, IL-10, and VEGF-α. In essence, these studies revealed that
the Ly6-Chi monocyte orchestrate the initial inflammatory event,
likely by forming M1 macrophages and then also develop into the
reparative, M2-like macrophage (43).
Extending these key findings to the atherosclerotic lesion, as
M1 macrophages can develop into M2 macrophages after Nr4a1
induction (43), and deletion of Nr4a1 results in M1 polarized
macrophages and increased atherosclerosis (44), it is possible that
the environment of the atherosclerotic lesion could affect the M1
macrophages resulting in a failure to upregulate Nr4a1 and prevents the differentiation into M2 cells (Figure 1). It should also
be noted that Ly6-Clo monocytes do enter the lesion (45), and
while these cells could become M2-like macrophages, they may not
frequent the lesion in large enough numbers to make an impact.
Another newly discovered macrophage subset is the Mox
macrophage. These macrophages are distinct to the classical M1
or M2 macrophage, as these cells display a unique gene expression profile with induction of redox-related genes including heme
oxygenase-1 under the control of the transcription factor Nrf2
(46). Mox macrophages also display a decrease in phagocytic and
chemotactic capacity. Interestingly, both M1 and M2 macrophages
can differentiate into the Mox macrophage when incubated with
oxidized phospholipids. The in vivo relevance of these cells is noted
as approximately 30% of all lesional macrophages are of the Mox
phenotype.

HYPERCHOLESTEROLEMIA INFLUENCES HSPCs TO
PRODUCE ATHEROGENIC MACROPHAGES
It is clear from animal studies that a hypercholesterolemic environment enhances the production of myeloid cells, namely monocytes, which contribute to atherogenesis. However, a hypercholesterolemic environment could also induce a “memory” effect in
the HSPCs, which could also alter the function of their daughter cells. This hypothesis was recently explored by Seijkens and
co-workers (47). Similar to our studies (4), they found that hypercholesterolemic Ldlr−/− mice had an expanded pool of HSPCs in
the BM. Interestingly, when they harvested the BM from hypercholesterolemic Ldlr−/− mice and transplanted it competition with
BM from normocholesterolemic mice, they found that the BM

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Cholesterol efflux, myelopoiesis, and macrophages

from the hypercholesterolemic mice had an enhanced propensity
to produce myeloid cells (47). This was even observed in a normocholesterolemic environment. Evidence was provided to support
the hypothesis that the hypercholesterolemic-primed HSPCs produced atherogenic (i.e., M1) macrophages as the macrophages
from these HSPCs produced higher amounts of TNF-α, IL-6, and
MCP-1. It was also found in the subsequent atherogenesis studies that hypercholesterolemic-primed HSPCs produced leukocytes
that more readily entered the atherosclerotic lesion. This resulted
in larger more macrophage-rich lesions.
The cBMT studies into hypercholesterolemic and normocholesterolemic mice suggest that there is a memory effect in the
HSPCs (47). This idea was recently brought to light by Kampen
et al., who discovered that BM harvested from WTD-fed mice has
a loss of epigenetic control of key myeloid genes such as PU.1
and IRF8 (48). Transplantation of the BM from the WTD-fed
mice into Ldlr −/− recipients, like the studies of Seijkens et al.,
also resulted in larger lesion compared to recipient mice that
received BM from chow fed donors. Consistent with the changes in
PU.1 and IRF8, the WTD-conditioned BM produced more leukocytes, particularly of the myeloid variety. There were also signs
of extramedullary hematopoiesis as the WTD-conditioned BMT
mice had splenomegaly. However, one caveat of this study was the
mice that received the WTD-conditioned BM-developed hyperglycemia, which has been shown to have independent effects on
BM progenitors to induce monocyte production and contribute
to atherosclerosis (49).
Another important point to note is that these studies either
performed BMTs using total BM or the total pool of HSPCs and
not just the long-term repopulating cells. Thus, as we have also
noted, a predominant expansion of the multipotential progenitor
2 (MMP2) HSPCs that is thought to give rise to myeloid cells in
Apoe−/− mice (4), it is possible that hypercholesterolemia-priming
promotes the expansion of a subset of HSPCs that preferentially
produces atherogenic myeloid cells.

EARLY MYELOID LINEAGE SKEWING IN ATHEROSCLEROSIS:
EMERGING CONCEPTS
The idea is emerging that signaling events in hematopoietic stem
cells (HSCs) are able to influence lineage selection in these cells.
Recently, it was discovered that HSCs express the M-CSF receptor and the engagement with M-CSF activated the myeloid master
regulator, PU.1 (50). Injection of mice with LPS increased MCSF levels and PU.1 expression in HSCs, which is likely to be
the initiating step of myeloid lineage skewing in response to an
infection. However, the LPS receptor TLR4 is also expressed on
BM stem and progenitor cells (51) and could have been an additional contributor to the early lineage selection in these studies.
Linking these findings with cholesterol metabolism, macrophages
deficient in either Abca1 and/or Abcg1 express more TLR4 on their
surface and like Apoe−/− macrophages are more responsive to
TLR4 ligands (3, 52, 53). Thus, it is also conceivable that defective cholesterol efflux pathways in HSCs could lead to enhanced
expression of TLR4 that could sense endogenous ligands, priming these cells to sense myeloid promoting cytokines. Whether
ligands of pattern recognition receptors (PRRs) such as damage
associated molecular pattern (DAMPs) (including S100A8/A9 and

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HMGB1), heat shock proteins, and modified LDL particles (54),
some of which are increased in people with CVD, are present
in the BM and bind to TLR4 on HSPCs is unknown. Assuming TLR4 ligands are present within the stem cell niche, it is
possible that their interaction with TLR4 on HSPCs could downregulate Abca1, Abcg1 (55), and Apoe (56) by the activation of
IRF3, preventing LXR activating these target genes (57). This lead
to increased cholesterol in the cell membrane and increased levels of cytokine receptors (4, 5). This could occur through the
prevention of key feedback loops, such as activation of the E3ubiquitin ligase c-CBL, which we recently reported was perturbed
in progenitor cells lacking ABCG4 (32), and is also reported to
downregulate the M-CSFR (58). Taken together, it is conceivable that defective cholesterol efflux and a hypercholesterolemic
environment could influence the HSCs to respond to myeloid
promoting cytokines to produce more monocytes that may have
an altered function, which could ultimately contribute to the
pool of inflammatory lesional macrophages in the atherosclerotic
plaque.
Dissecting out the contribution of changes in the HSPCs to the
function of the macrophage will be critical in further understanding the mechanisms contributing to not only atherogenesis but
also lesion regression. The lesion milieu is also critically important,
and is a dynamic environment with the newly recruited cells also
contributing to and being influenced by the environment. However, taken together, the emerging theme from recent literature
suggests that therapeutic interventions aimed at targeting HSPCs
(i.e., cholesterol efflux pathways) may be an effective strategy to
treat atherosclerosis by not only inhibiting monocyte production
and entry into lesions but also to change the function/phenotype
of the mature macrophage.

ACKNOWLEDGMENTS
Andrew James Murphy was supported by a Viertel award from
Diabetes Australia Research Trust Australia, and a National Health
and Medical Research Council program grant (APP10363652).
Dragana Dragoljevic was supported by an Australian Postgraduate Award. Alan Richard Tall was supported by a grant from the
NIH (HL107653).

Cholesterol efflux, myelopoiesis, and macrophages

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.
19.

REFERENCES
1. Libby P. Inflammation in atherosclerosis. Nature (2002) 420:868–74. doi:10.
1038/nature01323
2. Murphy AJ, Woollard KJ, Hoang A, Mukhamedova N, Stirzaker RA, McCormick
SP, et al. High-density lipoprotein reduces the human monocyte inflammatory response. Arterioscler Thromb Vasc Biol (2008) 28:2071–7. doi:10.1161/
ATVBAHA.108.168690
3. Yvan-Charvet L, Welch C, Pagler TA, Ranalletta M, Lamkanfi M, Han S,
et al. Increased inflammatory gene expression in ABC transporter-deficient
macrophages: free cholesterol accumulation, increased signaling via toll-like
receptors, and neutrophil infiltration of atherosclerotic lesions. Circulation
(2008) 118:1837–47. doi:10.1161/CIRCULATIONAHA.108.793869
4. Murphy AJ, Akhtari M, Tolani S, Pagler T, Bijl N, Kuo CL, et al. ApoE regulates hematopoietic stem cell proliferation, monocytosis, and monocyte accumulation in atherosclerotic lesions in mice. J Clin Invest (2011) 121:4138–49.
doi:10.1172/JCI57559
5. Yvan-Charvet L, Pagler T, Gautier EL, Avagyan S, Siry RL, Han S, et al. ATPbinding cassette transporters and HDL suppress hematopoietic stem cell proliferation. Science (2010) 328:1689–93. doi:10.1126/science.1189731
6. Robbins CS, Chudnovskiy A, Rauch PJ, Figueiredo JL, Iwamoto Y, Gorbatov
R, et al. Extramedullary hematopoiesis generates Ly-6C(high) monocytes that

www.frontiersin.org

20.

21.

22.

23.

24.

25.

infiltrate atherosclerotic lesions. Circulation (2012) 125:364–74. doi:10.1161/
CIRCULATIONAHA.111.061986
Kocaman SA, Sahinarslan A, Kunak T, Balcioğlu S, Cetin M, Cemri M, et al. The
particular interactions of the traditional cardiovascular risk factors with different circulating specific leukocyte subtype counts in blood: an observational
study. Anadolu Kardiyol Derg (2011) 11:573–81. doi:10.5152/akd.2011.158
Lee CD, Folsom AR, Nieto FJ, Chambless LE, Shahar E, Wolfe DA. White
blood cell count and incidence of coronary heart disease and ischemic stroke
and mortality from cardiovascular disease in African-American and white men
and women: atherosclerosis risk in communities study. Am J Epidemiol (2001)
154:758–64. doi:10.1093/aje/154.8.758
Mochizuki K, Miyauchi R, Misaki Y, Kasezawa N, Tohyama K, Goda T. Associations between leukocyte counts and cardiovascular disease risk factors
in apparently healthy Japanese men. J Nutr Sci Vitaminol (2012) 58:181–6.
doi:10.3177/jnsv.58.181
Sweetnam PM, Thomas HF, Yarnell JW, Baker IA, Elwood PC. Total and differential leukocyte counts as predictors of ischemic heart disease: the Caerphilly and Speedwell studies. Am J Epidemiol (1997) 145:416–21. doi:10.1093/
oxfordjournals.aje.a009123
Waterhouse DF, Cahill RA, Sheehan F, McCreery C. Prediction of calculated
future cardiovascular disease by monocyte count in an asymptomatic population. Vasc Health Risk Manag (2008) 4:177–87. doi:10.2147/vhrm.2008.04.01.
177
Rogacev KS, Seiler S, Zawada AM, Reichart B, Herath E, Roth D, et al.
CD14++CD16+ monocytes and cardiovascular outcome in patients with
chronic kidney disease. Eur Heart J (2011) 32:84–92. doi:10.1093/eurheartj/
ehq371
Rogacev KS, Ulrich C, Blömer L, Hornof F, Oster K, Ziegelin M, et al. Monocyte heterogeneity in obesity and subclinical atherosclerosis. Eur Heart J (2010)
31:369–76. doi:10.1093/eurheartj/ehp308
Rogacev KS, Zawada AM, Emrich I, Seiler S, Böhm M, Fliser D, et al. Lower
apo A-I and lower HDL-C levels are associated with higher intermediate
CD14++CD16+ monocyte counts that predict cardiovascular events in CKD.
Arterioscler Thromb Vasc Biol (2014) 34:2120–7. doi:10.1161/ATVBAHA.114.
304172
Swirski FK, Libby P, Aikawa E, Alcaide P, Luscinskas FW, Weissleder R, et al. Ly6Chi monocytes dominate hypercholesterolemia-associated monocytosis and
give rise to macrophages in atheromata. J Clin Invest (2007) 117:195–205.
doi:10.1172/JCI29950
Tacke F,Alvarez D, Kaplan TJ, Jakubzick C, Spanbroek R, Llodra J, et al. Monocyte
subsets differentially employ CCR2, CCR5, and CX3CR1 to accumulate within
atherosclerotic plaques. J Clin Invest (2007) 117:185–94. doi:10.1172/JCI28549
Westerterp M, Gourion-Arsiquaud S, Murphy AJ, Shih A, Cremers S, Levine
RL, et al. Regulation of hematopoietic stem and progenitor cell mobilization
by cholesterol efflux pathways. Cell Stem Cell (2012) 11:195–206. doi:10.1016/j.
stem.2012.04.024
Averill LE, Meagher RC, Gerrity RG. Enhanced monocyte progenitor cell proliferation in bone marrow of hyperlipemic swine. Am J Pathol (1989) 135:369–77.
Feldman DL, Mogelesky TC, Liptak BF, Gerrity RG. Leukocytosis in rabbits with
diet-induced atherosclerosis. Arterioscler Thromb Vasc Biol (1991) 11:985–94.
doi:10.1161/01.ATV.11.4.985
Huang ZS, Chiang BL. Correlation between serum lipid profiles and the ratio
and count of the CD16+ monocyte subset in peripheral blood of apparently
healthy adults. J Formos Med Assoc (2002) 101:11–7.
Tolani S, Pagler TA, Murphy AJ, Bochem AE, Abramowicz S, Welch C, et al.
Hypercholesterolemia and reduced HDL-C promote hematopoietic stem cell
proliferation and monocytosis: studies in mice and FH children. Atherosclerosis
(2013) 229:79–85. doi:10.1016/j.atherosclerosis.2013.03.031
Rajavashisth T, Qiao JH, Tripathi S, Tripathi J, Mishra N, Hua M, et al. Heterozygous osteopetrotic (op) mutation reduces atherosclerosis in LDL receptordeficient mice. J Clin Invest (1998) 101:2702–10. doi:10.1172/JCI119891
Guilliams M, Ginhoux F, Jakubzick C, Naik SH, Onai N, Schraml BU, et al.
Dendritic cells, monocytes and macrophages: a unified nomenclature based on
ontogeny. Nat Rev Immunol (2014) 14:571–8. doi:10.1038/nri3712
Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41:14–20. doi:10.1016/j.immuni.2014.06.008
Robbins CS, Hilgendorf I, Weber GF, Theurl I, Iwamoto Y, Figueiredo JL, et al.
Local proliferation dominates lesional macrophage accumulation in atherosclerosis. Nat Med (2013) 19:1166–72. doi:10.1038/nm.3258

October 2014 | Volume 5 | Article 490 | 167

Murphy et al.

26. Canton J, Neculai D, Grinstein S. Scavenger receptors in homeostasis and immunity. Nat Rev Immunol (2013) 13:621–34. doi:10.1038/nri3515
27. Khera AV, Cuchel M, de laLlera-Moya M, Rodrigues A, Burke MF, Jafri K, et al.
Cholesterol efflux capacity, high-density lipoprotein function, and atherosclerosis. N Engl J Med (2011) 364:127–35. doi:10.1056/NEJMoa1001689
28. Murphy AJ, Funt S, Gorman D, Tall AR, Wang N. Pegylation of high-density
lipoprotein decreases plasma clearance and enhances antiatherogenic activity.
Circ Res (2013) 113:e1–9. doi:10.1161/CIRCRESAHA.113.301112
29. Plump AS, Scott CJ, Breslow JL. Human apolipoprotein A-I gene expression increases high density lipoprotein and suppresses atherosclerosis in the
apolipoprotein E-deficient mouse. Proc Natl Acad Sci USA (1994) 91:9607–11.
doi:10.1073/pnas.91.20.9607
30. Tall AR, Yvan-Charvet L, Westerterp M, Murphy AJ. Cholesterol efflux: a novel
regulator of myelopoiesis and atherogenesis. Arterioscler Thromb Vasc Biol
(2012) 32:2547–52. doi:10.1161/ATVBAHA.112.300134
31. Murphy AJ, Westerterp M, Yvan-Charvet L, Tall AR. Anti-atherogenic mechanisms of high density lipoprotein: effects on myeloid cells. Biochim Biophys Acta
(2012) 1821:513–21. doi:10.1016/j.bbalip.2011.08.003
32. Murphy AJ, Bijl N, Yvan-Charvet L, Welch CB, Bhagwat N, Reheman A, et al.
Cholesterol efflux in megakaryocyte progenitors suppresses platelet production
and thrombocytosis. Nat Med (2013) 19:586–94. doi:10.1038/nm.3150
33. Swirski FK, Nahrendorf M, Etzrodt M, Wildgruber M, Cortez-Retamozo V,
Panizzi P, et al. Identification of splenic reservoir monocytes and their deployment to inflammatory sites. Science (2009) 325:612–6. doi:10.1126/science.
1175202
34. Wang M, Subramanian M, Abramowicz S, Murphy AJ, Gonen A, Witztum J, et al.
Interleukin-3/granulocyte macrophage colony-stimulating factor receptor promotes stem cell expansion, monocytosis, and atheroma macrophage burden in
mice with hematopoietic ApoE deficiency. Arterioscler Thromb Vasc Biol (2014)
34:976–84. doi:10.1161/ATVBAHA.113.303097
35. Westerterp M, Murphy AJ, Wang M, Pagler TA, Vengrenyuk Y, Kappus MS, et al.
Deficiency of ATP-binding cassette transporters A1 and G1 in macrophages
increases inflammation and accelerates atherosclerosis in mice. Circ Res (2013)
112:1456–65. doi:10.1161/CIRCRESAHA.113.301086
36. Metcalf D. Hematopoietic cytokines. Blood (2008) 111:485–91. doi:10.1182/
blood-2007-03-079681
37. Pagler TA, Wang M, Mondal M, Murphy AJ, Westerterp M, Moore KJ, et al. Deletion of ABCA1 and ABCG1 impairs macrophage migration because of increased
Rac1 signaling. Circ Res (2011) 108:194–200. doi:10.1161/CIRCRESAHA.110.
228619
38. Schulz C, Gomez Perdiguero E, Chorro L, Szabo-Rogers H, Cagnard N, Kierdorf
K, et al. A lineage of myeloid cells independent of Myb and hematopoietic stem
cells. Science (2012) 336:86–90. doi:10.1126/science.1219179
39. Epelman S, Lavine KJ, Beaudin AE, Sojka DK, Carrero JA, Calderon B, et al.
Embryonic and adult-derived resident cardiac macrophages are maintained
through distinct mechanisms at steady state and during inflammation. Immunity (2014) 40:91–104. doi:10.1016/j.immuni.2013.11.019
40. Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, et al. Tissueresident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity (2013) 38:792–804.
doi:10.1016/j.immuni.2013.04.004
41. Heidt T, Courties G, Dutta P, Sager HB, Sebas M, Iwamoto Y, et al. Differential contribution of monocytes to heart macrophages in steady-state and after
myocardial infarction. Circ Res (2014) 115:284–95. doi:10.1161/CIRCRESAHA.
115.303567
42. Hanna RN, Carlin LM, Hubbeling HG, Nackiewicz D, Green AM, Punt JA,
et al. The transcription factor NR4A1 (Nur77) controls bone marrow differentiation and the survival of Ly6C-monocytes. Nat Immunol (2011) 12:778–85.
doi:10.1038/ni.2063
43. Hilgendorf I, Gerhardt LM, Tan TC, Winter C, Holderried TA, Chousterman BG,
et al. Ly-6Chigh monocytes depend on Nr4a1 to balance both inflammatory and
reparative phases in the infarcted myocardium. Circ Res (2014) 114:1611–22.
doi:10.1161/CIRCRESAHA.114.303204
44. Hanna RN, Shaked I, Hubbeling HG, Punt JA, Wu R, Herrley E, et al. NR4A1
(Nur77) deletion polarizes macrophages toward an inflammatory phenotype and increases atherosclerosis. Circ Res (2012) 110:416–27. doi:10.1161/
CIRCRESAHA.111.253377

Frontiers in Immunology | Molecular Innate Immunity

Cholesterol efflux, myelopoiesis, and macrophages

45. Potteaux S, Gautier EL, Hutchison SB, van Rooijen N, Rader DJ, Thomas MJ,
et al. Suppressed monocyte recruitment drives macrophage removal from atherosclerotic plaques of Apoe−/− mice during disease regression. J Clin Invest
(2011) 121:2025–36. doi:10.1172/JCI43802
46. Kadl A, Meher AK, Sharma PR, Lee MY, Doran AC, Johnstone SR, et al.
Identification of a novel macrophage phenotype that develops in response to
atherogenic phospholipids via Nrf2. Circ Res (2010) 107:737–46. doi:10.1161/
CIRCRESAHA.109.215715
47. Seijkens T, Hoeksema MA, Beckers L, Smeets E, Meiler S, Levels J, et al.
Hypercholesterolemia-induced priming of hematopoietic stem and progenitor
cells aggravates atherosclerosis. FASEB J (2014) 28(5):2202–13. doi:10.1096/fj.
13-243105
48. van Kampen E, Jaminon A, van Berkel TJC, Van Eck M. Diet-induced (epigenetic) changes in bone marrow augment atherosclerosis. J Leukoc Biol (2014).
doi:10.1189/jlb.1A0114-017R
49. Nagareddy PR, Murphy AJ, Stirzaker RA, Hu Y, Yu S, Miller RG, et al. Hyperglycemia promotes myelopoiesis and impairs the resolution of atherosclerosis.
Cell Metab (2013) 17:695–708. doi:10.1016/j.cmet.2013.04.001
50. Mossadegh-Keller N, Sarrazin S, Kandalla PK, Espinosa L, Stanley ER, Nutt SL,
et al. M-CSF instructs myeloid lineage fate in single haematopoietic stem cells.
Nature (2013) 497:239–43. doi:10.1038/nature12026
51. Nagai Y, Garrett KP, Ohta S, Bahrun U, Kouro T, Akira S, et al. Toll-like receptors
on hematopoietic progenitor cells stimulate innate immune system replenishment. Immunity (2006) 24:801–12. doi:10.1016/j.immuni.2006.04.008
52. Ali K, Middleton M, Pure E, Rader DJ. Apolipoprotein E suppresses the type I
inflammatory response in vivo. Circ Res (2005) 97:922–7. doi:10.1161/01.RES.
0000187467.67684.43
53. Van Oosten M, Rensen PC, Van Amersfoort ES, Van Eck M, Van Dam AM, Breve
JJ, et al. Apolipoprotein E protects against bacterial lipopolysaccharide-induced
lethality. A new therapeutic approach to treat gram-negative sepsis. J Biol Chem
(2001) 276:8820–4. doi:10.1074/jbc.M009915200
54. Erridge C. Endogenous ligands of TLR2 and TLR4: agonists or assistants?
J Leukoc Biol (2010) 87:989–99. doi:10.1189/jlb.1209775
55. Maitra U, Li L. Molecular mechanisms responsible for the reduced expression of cholesterol transporters from macrophages by low-dose endotoxin.
Arterioscler Thromb Vasc Biol (2013) 33:24–33. doi:10.1161/ATVBAHA.112.
300049
56. Gafencu AV, Robciuc MR, Fuior E, Zannis VI, Kardassis D, Simionescu
M. Inflammatory signaling pathways regulating ApoE gene expression in
macrophages. J Biol Chem (2007) 282:21776–85. doi:10.1074/jbc.M611422200
57. Castrillo A, Joseph SB, Vaidya SA, Haberland M, Fogelman AM, Cheng G,
et al. Crosstalk between LXR and toll-like receptor signaling mediates bacterial and viral antagonism of cholesterol metabolism. Mol Cell (2003) 12:805–16.
doi:10.1016/S1097-2765(03)00384-8
58. Mancini A, Koch A, Wilms R, Tamura T. c-Cbl associates directly with the Cterminal tail of the receptor for the macrophage colony-stimulating factor, cFms, and down-modulates this receptor but not the viral oncogene v-Fms. J Biol
Chem (2002) 277:14635–40. doi:10.1074/jbc.M109214200

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 04 August 2014; paper pending published: 19 August 2014; accepted: 22
September 2014; published online: 13 October 2014.
Citation: Murphy AJ, Dragoljevic D and Tall AR (2014) Cholesterol efflux pathways
regulate myelopoiesis: a potential link to altered macrophage function in atherosclerosis.
Front. Immunol. 5:490. doi: 10.3389/fimmu.2014.00490
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Murphy, Dragoljevic and Tall. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The
use, distribution or reproduction in other forums is permitted, provided the original
author(s) or licensor are credited and that the original publication in this journal is cited,
in accordance with accepted academic practice. No use, distribution or reproduction is
permitted which does not comply with these terms.

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REVIEW ARTICLE
published: 12 November 2014
doi: 10.3389/fimmu.2014.00579

Dynamic aspects of macrophage polarization during
atherosclerosis progression and regression
Michael Peled * and Edward A. Fisher *
The Marc and Ruti Bell Program in Vascular Biology, Department of Medicine, Leon H. Charney Division of Cardiology, New York University School of Medicine,
New York, NY, USA

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Klaus Ley, La Jolla Institute for Allergy
& Immunology, USA
Janos G. Filep, University of
Montreal, Canada
*Correspondence:
Michael Peled and Edward A. Fisher ,
The Marc and Ruti Bell Program in
Vascular Biology, Department of
Medicine, Leon H. Charney Division
of Cardiology, New York University
School of Medicine, Smilow 7, 522
First Avenue, New York, NY 10016,
USA
e-mail: michael.peled@nyumc.org;
edward.fisher@nyumc.org

It is well recognized that macrophages in many contexts in vitro and in vivo display a spectrum of inflammatory features and functional properties. A convenient system to group
together different subsets of macrophages has been the M1 (inflammatory)/M2 (antiinflammatory) classification. In addition to other sites of inflammation, it is now established
that atherosclerotic plaques contain both M1 and M2 macrophages. We review results
made possible by a number of recent mouse models of atherosclerotic regression that,
taken with other literature, have shown the M1/M2 balance in plaques to be dynamic,
with M1 predominating in disease progression and M2 in regression. The regulation of the
macrophage phenotype in plaques and the functional consequences of the M1 and M2
states in atherosclerosis will also be discussed.
Keywords: macrophages, atherosclerosis, regression, cholesterol, polarization

INTRODUCTION
Atherosclerotic cardiovascular diseases, which include myocardial
infraction and stroke, are the most common causes of morbidity and mortality in western society and will soon be the
same world-wide. Atherosclerosis represents a failure to resolve
the inflammatory response in the arterial wall initiated by the
retention of apolipoprotein B (apoB)-containing lipoproteins (1).
These lipoproteins are taken up by tissue macrophages, which
ultimately become engorged with cholesterol (foam cells) and
activated. The continuing stimulus of the entry and retention of
apoB-lipoproteins fuels not only the accumulation of foam cells
to form a plaque, but also the chronicity and amplification of the
inflammatory response, which contribute to the vulnerability of
some plaques to rupture and cause acute tissue ischemia. The central role of macrophages in atherosclerosis pathophysiology has,
therefore, focused attention on their properties in plaque initiation
and progression, and more recently, in regression (2–5).
The study of macrophages over the past decade is characterized
by a remarkable expansion of knowledge concerning their origin,
functional properties, and potential to both protect from and contribute to disease [e.g., see in Ref. (6, 7)]. Admitting the complexity
of macrophage biology, for the purposes of this review, we have
restricted ourselves to considering how aspects of macrophage
polarization in the M1/M2 classification system (8) relate to atherosclerosis progression and regression. In this system, which is
influenced by the Th1 and Th2 classification of lymphocytes,
macrophages can be grossly divided into pro-inflammatory, M1
cells and anti-inflammatory, M2 cells based mainly on in vitro
criteria (9). It is important to note, however, that while the classification of lymphocytes into Th1 and Th2 preceded the classification

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scheme of macrophages as M1/M2, the Th1 and Th2-like responses
result from polarization of macrophages to M1 and M2 states,
respectively. Furthermore, M1/M2 polarization is not dependent
on T cells, as has been demonstrated in Rag KO and other immune
deficient mice (8).
Polarization toward the M1 state is induced by several stimuli in vitro, including Toll-like receptor (TLR) ligands (such
as lipopolysaccharide, LPS) and interferon γ (potential endogenous stimuli in atherosclerosis will be discussed below). M1
macrophages express several pro-inflammatory mediators, such as
inducible nitric oxide synthase, tumor necrosis factor-a (TNF-a),
interleukin-1b (IL-1b), IL-6, IL-12, and proteolytic enzymes. M1
macrophages have been found in both human and mouse atherosclerosis, and their secretion of the pro-inflammatory mediators
is thought to maintain local inflammation and the degradation of
extracellular matrix components, resulting in disease progression
[e.g., Ref. (10)], and ultimately, unstable plaques in humans. As
alluded to earlier, unstable plaques are at increased risk of rupture
and causing thrombosis, resulting in myocardial infarction and
stroke (11–14).
On the other side of the polarization spectrum, M2
macrophages are induced in vitro by Th2-type cytokines, such
as IL-4 and IL-13. T regulatory cells (Tregs) have also been implicated in the induction of M2 polarization (15), possibly through
IL-10. M2 polarized macrophages have been characterized by their
expression of CD163, mannose receptor 1 (also known as CD206),
FIZZ1, and high levels of arginase 1. In addition, they secrete antiinflammatory cytokines, such as transforming growth factorβ, IL-1
receptor antagonist, IL-10, and have increased secretion of collagen. Based particularly on the role of M2 macrophages in wound

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Peled and Fisher

healing, the combination of the factors they express are thought
to be particularly suited for tissue repair, which is consistent with
their increased presence in regressing plaques in mouse models of
atherosclerosis (16, 17).
In addition to the M1 and M2 macrophages, oxidized phospholipids present in oxidized LDL induce a macrophage phenotype
that is distinct from M1 or M2 phenotypes and that has been
termed Mox; these macrophages are characterized by the increases
in the expression of nuclear factor erythroid 2-related factor 2
(NRF2)-dependent genes and in reactive oxygen species, and are
found in the progressing plaques (18). Their role in atherosclerosis
regression, however, has not been established.
With this background, we will now turn to a more complete consideration of the inflammatory states of macrophages
in atherosclerosis progression and regression.

ATHEROSCLEROSIS PROGRESSION
DRIVERS OF MACROPHAGE INFLAMMATION IN PROGRESSING
PLAQUES

As mentioned above, M1 macrophages are thought to have significant roles in progressing and vulnerable plaques (19). As noted
above, a potent inducer of the M1 state in vitro is LPS, which is
part of the outer membrane of Gram-negative bacteria, and which
binds and activates TLR4 (20). Dozens of studies have demonstrated associations among high-fat diet, the metabolic syndrome,
and endotoxemia (i.e., increased levels of LPS in the plasma) (21–
23). It appears that a high-fat diet, the consumption of which leads
to metabolic syndrome in mouse models, also induces a change
in the gut bacterial flora, which, in turn, causes an increase in the
LPS plasma levels. Circumstantial considerations that support a
link between the metabolic syndrome and LPS signaling include
the increased cardiovascular risk in patients with the metabolic
syndrome and the positive association of TLR4 activity with atherosclerosis progression in mice and human beings (24–28). Despite
the evidence for association between LPS, atherosclerosis progression, and M1 polarization, a study in germ-free apoE-deficient
mice on low-fat chow diet showed increased atherosclerosis progression (29), suggesting that some bacteria in the gut flora have
an anti-inflammatory effect (30).
Cholesterol and macrophage activation

The most accepted and robust risk factor for atherosclerosis is
low-density lipoprotein-cholesterol (LDL-C). Thus, several studies have tried to understand how cholesterol can induce inflammation in general, and specifically to an activated state. The different
mechanisms by which cholesterol can drive macrophage activation could be divided into those direct – how cholesterol affects
macrophages, and indirect – how cholesterol affects other cell types
through which activation could be induced, for example by the
secretion of pro-inflammatory cytokines from T cells (Figure 1).
Direct mechanisms that link cholesterol to macrophage inflammation. Accumulation of cholesterol leads to the formation of
crystals that are both intra- and extracellular. The presence of
cholesterol crystals in early lesions in Apoe-/- mice was recently
demonstrated (31, 32); in addition, it was shown that both
macrophage engulfment of cholesterol crystals and de novo formation of intracellular cholesterol crystals activate the NLRP3

Frontiers in Immunology | Molecular Innate Immunity

Macrophage polarization in atherosclerosis

(NOD-, LRR- and pyrin domain-containing 3) inflammasome
(33). Activation of NLRP3, in turn, results in the secretion of
the pro-inflammatory cytokine IL-1β.This pathway appears to be
necessary for atherosclerosis progression, as LDL receptor (Ldlr)/- mice, transplanted with bone marrow cells deficient in IL-1β or
in components of the NLRP3 inflammasome, had reduced plaque
progression (31, 34).
A second direct mechanism that can explain macrophage activation by cholesterol is mediated through oxidized LDL. Oxidized
LDL is present in both human and mouse atheromas. LDL oxidation is thought to be mediated by enzymes (such as 12/15lipoxygenase and myeloperoxidase) and by free radicals that are
abundant in the atherosclerotic plaque (35). Several studies have
demonstrated that oxidized LDL can act as a ligand for both the
scavenger receptors and TLRs on macrophages. Depending on the
extent of oxidation, species of oxidized LDL were found to be
agonists of CD14–TLR4–MD2 or CD36–TLR4–TLR6 complexes
in vitro; these complexes can induce a pro-inflammatory signaling cascade involving IL-1 receptor-associated kinase 4 (IRAK4)
(36, 37), myeloid differentiation primary-response protein 88
(MYD88) (28, 38) and other signaling proteins, culminating in
activation of Nf-kB targets. Whole-body knock out mice of TLR2,
TLR4 and some of the signaling proteins mentioned here were
demonstrated to have reduced atherosclerosis progression (28, 39).
A third direct mechanism points toward the increase in plasma
membrane cholesterol. The change in the microenvironment of
TLRs (40, 41) renders them more sensitive to ligands, thereby
heightening the inflammatory responses.
Despite the predominance of data that show a direct link
between cholesterol and macrophage inflammation, there are a
number of studies that show the opposite – an anti-inflammatory
phenotype induced by intracellular cholesterol. The basis for this is
rooted in the LXRs and PPARs, which are important nuclear receptors. The ligands for these receptors include lipids, and in addition
to regulating many steps of their metabolism, LXRs and PPARs
can also suppress inflammatory signaling in macrophages (42).
For example, a recent study demonstrated that cholesterol-loading
induced LXR-dependent downregulation of inflammatory gene
expression in macrophages as a consequence of the accumulation
of the cholesterol precursor desmosterol (43).
Indirect mechanisms that link cholesterol to macrophage inflammation. It has been long known that T cells participate in plaque
inflammation (4). For example, using flow cytometry of whole
aortic digests, Galkina et al. could demonstrate the presence of
diverse cell-types of the adaptive immune system in the atherosclerotic plaque and the surrounding adventitia (44). Moreover,
the same group showed that antigen presentation to CD4+ T cells
in the arterial wall causes local T cell activation and the secretion
of pro-inflammatory cytokines, which promote atherosclerosis by
maintaining chronic inflammation and inducing foam cell formation (45). Other studies have shown a direct pro-inflammatory
role of intracellular cholesterol in T cells, mediated by cholesterolinduced nano-clustering of T cell receptors (46). In addition, mice
lacking T cells can have a significant reduction in atherosclerosis
progression (47). Some studies, however, have demonstrated only
a minor role for T cells in atherosclerosis progression (48).

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Peled and Fisher

FIGURE 1 | Summary of how changes in plasma cholesterol level and
in cellular cholesterol content affect macrophage polarization and
kinetics in atherosclerotic plaque progression and regression. Left
panel: An increase in non-HDL cholesterol (mainly VLDL cholesterol and
LDL cholesterol) in mouse models has been linked to an increase in
monocyte recruitment into atherosclerotic plaques, with their subsequent
polarization to M1 macrophages, which are retained. This ultimately leads
to atherosclerotic plaque progression, as evident by plaque enlargement.
The failure to clear dead macrophages by efferocytosis results in the
appearance of the necrotic core. Right panel: An opposite effect has been
demonstrated in atherosclerosis regression models, where a reduction in
non-HDL-C or a selective increase in HDL-C (representing an increase in
functional HDL particles) induces a decrease in plaque size and

Another indirect effect on macrophage polarization/activation
by cholesterol is related to its being a major structural component of progressing plaques, so its ongoing accumulation will
contribute to enlargement of the atheroma. This, in turn, is likely
to contribute to a hypoxic environment because as the atheroma
grows, the distance between the intimal cells from an oxygen supply will increase, particularly in mice, which have little capacity
to form vasa vasorum. Hypoxia triggers a heavy reliance on glycolysis for energy production and it has been recently recognized
that M1 cells are more glycolytic and M2 cells are more fatty acid
oxidizing, and that factors that promote one pathway of energy
generation over the other will promote the polarization state corresponding to the favored pathway (49). In addition, hypoxia can
contribute to the formation of the necrotic core, a characteristic
feature of advanced plaques that in humans increases the possibility of rupture (50). In response to the hypoxic conditions in
the arterial wall, the development of vasa vasorum is enhanced,

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Macrophage polarization in atherosclerosis

macrophage content (from decreased monocyte recruitment and
macrophage retention), as well as enrichment in the expression of
markers of the M2 state. Improved efferocytosis is also expected under
these conditions, with shrinkage of the necrotic core. There is an increase
in collagen content, likely from decreased MMP production by the
macrophages. It is also likely that in a regression environment there are
decreases in the secretion of inflammatory cytokines and chemokines by
the macrophages as a result of the polarization of macrophages toward a
M2-like state. The different mechanisms by which cholesterol can drive
macrophage activation and polarization are divided into those direct – how
cholesterol affects macrophages, and indirect – how cholesterol affects
other cell types, for example by the secretion of pro-inflammatory
cytokines from T cells.

and this has also been related to atherosclerosis progression, possibly by the recruitment of monocytes to the plaque through this
vascular route (51).
MACROPHAGE POLARIZATION AND LOCATION IN PROGRESSING
PLAQUES

Recent studies have found that although both M1 and M2
macrophage numbers are increased during human plaque progression, M1 macrophages were the predominant phenotype in
rupture-prone shoulder regions, whereas M2 markers were predominant in the adventitia and in stable cell-rich areas the of
plaque (19). In another study of human plaques, using the mannose receptor as a marker, M2 macrophages were found located
far from the lipid core of the plaque (52).
In mouse plaques, M1 macrophages tend to be diffusely distributed in, and characteristic of, progressing plaques at the usual age
that atherosclerotic mice are examined in detail (typically after

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12–16 of the consumption of a high-fat, high-cholesterol diet,
or ~16–20 weeks of age). Khallou-Laschet et al. have found in
apoE-deficient mice of a similar age, but with less advanced atherosclerosis because they were maintained on a low-fat, chow diet
that these early plaques were infiltrated by M2 macrophages, with
M1 macrophages appearing later. Disease progression correlated
with the dominance of M1 over the M2 phenotype (53). Based
on serial histologic examination, they further propose that the
M2 → M1 shift in balance was due to a phenotypic switch of the
infiltrated cells, but the data to support this were indirect and
did not exclude the possibility of replacement of macrophages by
new ones with a different phenotype, or the local proliferation
of M2-like tissue-resident cells, which has been reported in other
contexts as well (54, 55). In regard to the location of the different macrophage phenotypes in the plaque, they show that M2
macrophages accumulated in the luminal side of the plaques in
young mice, while at 55 weeks of age, both M1 and M2 labeling
was evenly distributed across the plaque.
FUNCTIONAL CONSEQUENCES OF M1 POLARIZATION IN PROGRESSING
PLAQUES

The secretion of a wide range of cytokines and chemokines
(e.g., IL-1, TNFalpha, MCP-1) by M1 macrophages serves to
further activate macrophages, as well as other cell types in the
atheroma, such as endothelial and smooth muscle cells. There are
also effects on cellular lipid metabolism. In one study of human
plaques, macrophages with a marker of M2 macrophages had
small lipid droplets and in studies in vitro, this was associated
with a decreased ability to efflux cholesterol (52). In a study of
murine macrophages, when the cells were polarized to the M1
state, there was downregulation of ABCA1 and reduced cholesterol efflux (56). If this were to happen in vivo, a vicious cycle
would be formed – cholesterol accumulation would contribute
to macrophage activation and M1 polarization, which would further induces cholesterol accumulation. Before concluding that this
happens, given the divergent results on the effects of the M1 and
M2 phenotypes on cholesterol efflux, more research in this area is
clearly needed.
M1 macrophages also secrete chemokines (such as MCP-1) and
cytokines (such as IL-12) that induce chemotaxis of other white
blood cells (57, 58). In addition, M1 macrophages secrete several
matrix metallo-proteinases (MMPs), such as MMP2 and MMP9,
that can degrade the extracellular matrix in the plaque, which
is thought to lead to destabilization and rupture. Indeed, MMPs
were shown to co-localize with M1 macrophages in atherosclerotic
plaques (59).

ATHEROSCLEROSIS REGRESSION
Fatty streaks, the initial phase in the development of plaques,
were found in children as young as 3 years of age (60), with wellestablished plaques developing by adolescence (61). Thus, while
most studies are focused on the progression of atherosclerosis and
finding means to delay it, the more frequent clinical scenario is that
by the time the patient comes for treatment of cardiovascular risk
factors, as a frequently middle-aged adult, he or she may already
have a significant burden of atherosclerosis, making the optimal
goal of therapy the induction of plaque regression.

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Macrophage polarization in atherosclerosis

Plaque regression can be defined in various ways, such as
a reduction in plaque size, plaque cholesterol content, plaque
macrophage number/percentage, or a decreased inflammatory
state. Of course, multiple changes can occur simultaneously, but
not in every case. For example, if the plaque macrophage content
decreases, while collagen content increases, as we have observed
experimentally in some models of regression [e.g., Ref. (16)], the
size may not change, but there will be less inflammation and more
stabilizing material. Nevertheless, though size changes may vary,
one consistent finding in various mouse models of atherosclerosis regression in which the issue of macrophage polarization was
examined, as will be summarized below, is that the plaque content
of M1 markers decreased, while those of M2 markers increased
[e.g., Ref. (16, 17, 62)].
ATHEROSCLEROSIS REGRESSION MODELS AND CHANGES IN PLAQUE
MACROPHAGES

Some of the currently available regression models include aortic
arch transplantation model (17, 63), Reversa mouse model (16,
64), adenoviral gene transfer of the LDL receptor, apoA1 or apoE,
a “hypomorphic apoE” model, administration of an inhibitor of
MTP, and infusion of apoA-I (the major protein in HDL) or apoA-I
mimetics (5, 65–69). By necessity, all models begin with a progression phase, in which the total plasma cholesterol, and in particular,
LDL-C and VLDL-C, are very high. After a certain period of time,
preferably at least 12–16 weeks of western diet (rich in saturated
fat and cholesterol) in order to accelerate in apoE-deficient-based
or enable in Ldl receptor-deficient-based models the development
of a complex atherosclerotic plaque, regression is typically induced
by a major change in the plasma lipid profile. This change is either
a reduction of LDL/VLDL-C or an increase in apoA1/HDL, both
of which would decrease the plaque content of cholesterol.
In the transplantation model, the plaque-containing aortic arch
from a donor apoE-deficient (apoE −/− ) or Ldl receptor-deficient
(ldlr −/− ) mouse fed a high-fat diet for 16 weeks is transferred into
the abdominal aorta of a normo-lipidemic wild-type recipient
mouse or an apoE −/− mouse made transgenic in human apoA1
(“hA1/EKO”) (17, 63, 70, 71). In either case, the regression of the
plaque in the transplanted arch occurs within a few days. The
advantage of the transplant model is that it can be used to test
the effects of specific genes on regression by using knock out or
transgenic strains, either for the donor or recipient mice, as well
as to conveniently study leukocyte trafficking in and out of the
plaques if the donors and recipients are mismatched in isoforms
of the pan-leukocyte marker for CD45. The disadvantage lies in
the technical difficulty in performing such a surgery in the mouse,
thereby limiting the throughput. In addition, there is inherent
inflammation induced by the surgery itself, which theoretically
can affect the process of regression, though control transplants
into apoE −/− mice are used as a standard control for these and
other effects related to the surgical procedures.
As noted above, HDL and its major protein, apoA1, can be
increased by injections of apoA1, apoA1 mimetics, or an adenoviral vector expressing apoA1 (69, 72). In advanced atherosclerotic
plaques of apoE −/− mice, every other day injections of native
human apoA-I over only 1 week led to atherosclerosis regression,
as demonstrated by significant decreases in plaque lipid content,

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macrophage number, and an increase in collagen content; moreover, apoA1 injections led to a significant reduction in the plaques
of inflammatory M1 and an increase in anti-inflammatory M2
macrophage markers, mannose receptor 1 and arginase 1 (73).
Treating mice with a recombinant adenovirus encoding human
apoA1 with relatively early atherosclerotic plaques resulted in a
70% reduction in aortic lesion area characterized by a significant
decrease in the fraction of lesions occupied by macrophages and
macrophage-derived foam cells. The inflammatory status of this
population of cells was not reported (69).
Another example for plaque regression induced by an increase
in HDL was shown with our collaborators using an inhibitor of
microRNA-33 (miR-33). miR-33 suppresses HDL formation in
the liver and its ability to efflux cholesterol from macrophages by
suppressing the expression of cholesterol transporter ATP-binding
cassette transporter 1 (ABCA1) (74). It was hypothesized that
inhibiting it by an antagomir (anti-miR-33) would promote atherosclerosis regression. Ldlr −/− mice with established plaques were
treated with anti-miR-33 over 4 weeks. As expected, anti-miR-33
treatment led to increased reverse cholesterol transport through
an increase in HDL levels and expression of ABCA1 in the liver
and macrophages. Consistent with that, and consistent with the
apoA1 injection study, atherosclerotic lesions regressed by antimiR-33 treatment, as shown by reduced plaque size, lipid and
macrophage content, increased collagen content and a diminished
inflammatory state of the macrophages in the plaque (75).
The Reversa (Ldlr −/− ApoB100/100Mttpfl/flMx1Cre +/+ ) mouse
is a non-surgical regression model, based on the Ldlr −/− mouse,
in which the hyperlipidemia can be reversed by inducing the conditional knock out of the microsomal triglyceride transfer protein
(MTTP) gene (64). MTTP is required for the proper assembly of
VLDL, the precursor of LDL (76). The reversal of hyperlipidemia
by inactivation of MTTP leads to regression of atherosclerosis over
a few weeks accompanied by favorable changes in the composition
of the atherosclerotic plaque. Again, plaque lipid content decreases,
collagen content increases, and M1 markers are decreased while
M2 markers are increased in the plaque macrophages (16). The
advantage of the Reversa model is that it does not require any
surgery in order to get extreme reduction in LDL/VLDL-C. In
addition, it is important to note that unlike the transplant models, there is no increase in HDL after inducing the conditional
knock out of the MTTP gene. This might be part of the reason
for the reduced regression rate that can be seen in the Reversa
model compared with the transplant model. The disadvantage
of the model lies in the complicated genetic manipulations that
were performed to create it – there are four different gene insertions/deletions in the Reversa mouse, and thus breeding it with
another transgenic/knock out mouse to test the importance of
a specific gene for regression is extremely time-intensive, making bone marrow transfer for myeloid-specific factors a more
convenient manipulation.
The hepatic overexpression of apoE in apoE −/− or ldlr in
ldlr −/− mice is two gene transfer strategies to induce regression,
again by normalization of the lipid profile (5). The drawback of
this method is a potential immune response of the host after
the adenoviral gene transfer (77), which might complicate the
interpretation of the inflammatory state of cells in the plaque.

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Macrophage polarization in atherosclerosis

Also, especially with early versions of viral vectors, there can be
limitations related to the duration or amplitude of expression.
Inducing regression just by a diet change, from western diet to
chow, in the ldlr −/− mouse model has also been tried. Many times,
no significant changes have been observed, presumably because the
plasma cholesterol levels remain elevated, and perhaps, the experiments were not continued long enough. In one recent report,
we and our collaborators have observed a reduction in plaque
macrophage content and inflammatory state over 4 weeks (78).
Notably, these favorable changes were impaired by hyperglycemia,
consistent with our previous report using Reversa mice (79).
Mechanisms for M2 macrophage enrichment in atherosclerosis
regression

There are two major questions we will consider in this section –
the origin of the M2 macrophages in regressing plaques and the
mechanisms for their increase.
One possibility is that the re-balancing from enrichment in M1
to M2 markers in regressing plaques represents either a change
in an individual cell, as can be accomplished in vitro by changing the cytokine environment, or as proposed to happen in the
“early to advanced” plaque transition in apoE −/− mice. Indeed, it
is generally accepted that the phenotype of a macrophage is quite
“plastic” and responsive to microenvironmental changes (9). It
is also possible that M1-like macrophages leave and are replaced
by M2-like cells, as occurs in wound healing. Support for this
scenario is our demonstration, particularly in the aortic transplantation model, of emigration of macrophages from, and the ongoing
recruitment of monocytes to, plaques in the regression environment (80). A third possibility is the induction of the proliferation
of a resident population of yolk-sac-derived M2 macrophages,
as observed in other settings (54). Support for this possibility is
the recent demonstration in progressing plaques for macrophage
proliferation (55).
It is tempting to speculate that the pro-activating direct and
indirect effects of cholesterol described above are reversed under
conditions of regression, under which the plaque content of cellular and extracellular cholesterol is typically reduced. Little experimental evidence, however, is available in vivo to prove this. Even
for factors with known cholesterol-removing function, such as
HDL, there could be lipid-independent reasons for their effects.
For example, we have recently reported that murine bone marrow derived-macrophages (BMDM) not loaded with cholesterol
will increase their M2 marker expression when incubated with
HDL (81) [though in human monocytes this was not found to
be the case (82)], and Latz and colleagues have found that HDL
will induce in macrophages the transcriptional regulator ATF3, a
repressor of a number of inflammatory factors (83). There is also
a study in which injection of apoA1 reduced skin inflammation
in ldlr −/− , apoA1−/− DKO mice (which show signs of autoimmunity) by augmenting the effectiveness of the lymph node Treg cells
(84); Tregs were shown to induce an M2 polarization in yet another
study (15). In this example, there may have been a lipid-removal
aspect in that apoA1−/− mice are deficient in cholesterol efflux
because of reduced numbers of HDL particles. Although unrelated directly to macrophage polarization, it has been shown that
defects in cholesterol efflux can enrich the plasma membranes of

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monocyte precursors in the bone marrow with cholesterol, which
results in their greater proliferation, circulating monocytosis, and
increased entry into plaques of monocytes, which subsequently
promotes atherogenesis (85). Importantly, we have recently found
that this mechanism might also be related to the impairment of
atherosclerosis regression in diabetes (7).
Turning to the issue of what regulates the enrichment in M2
macrophages in regressing plaques, there are no results to discuss
at this time, but there are a number of possibilities. As mentioned
earlier, potent cytokines that polarize macrophages are IL-4 and
IL-13. These can be derived from a variety of leukocytes, namely
Th2 lymphocytes, eosinophils, and basophils. Even if one or more
of these types of leukocytes were the source of polarizing signals,
there is still the mystery of how the change in the lipoprotein/lipid
environment causes either the recruitment of the cells to the plaque
or the stimulation of secretion from either pre-existing or newly
recruited cells.
Functional contribution of M2 polarized cells to atherosclerosis
regression

Another major and incompletely understood area is the requirement for, and the function of, the enrichment in M2 macrophages
in atherosclerosis regression. Because their properties include tissue remodeling and inflammation resolution, it is tempting to
attribute such changes in regressing plaques to M2 macrophages.
This would be consistent with studies in which treatment of
ldlr −/− mice with IL-13 resulted in pre-existing plaques the
following: an increase in collagen, a reduction in vascular cell
adhesion molecule-1 (VCAM-1)-dependent monocyte recruitment, decreased macrophage content, and the induction of
M2 macrophages, despite ongoing hyperlipidemia (86). Another
reflection of the importance of the M2 macrophages for regression comes from our studies in diabetic Reversa mice, in which
hyperglycemia impaired their enrichment in plaques despite lipid
lowering and limited the favorable changes in macrophage content
and inflammatory state (79).
The promotion of the resolution of inflammation by M2
macrophages in regressing plaques is likely a consequence of
their secretion of IL-10. The plaque remodeling may represent
at least two other properties of M2 macrophages, namely their
secretion of collagen (87) and their enhanced capacity for efferocytosis (88). Efferocytosis is the clearance of apoptotic cells by
macrophages. During atherosclerosis progression, the ability to
store cholesterol in plaque macrophages in the form of cholesteryl ester lipid droplets wanes and free cholesterol accumulates,
causing ER stress and apoptosis (89). If efferocytosis of the dying
cells does not keep up with their formation, the cellular debris
that are not cleared would be expected to accumulate and become
a necrotic core. Indeed, in both mouse and human plaques, As
plaques advance, the efferocytotic activity of their macrophages
decreases (50). Thus, an enrichment in M2 macrophages with
enhanced efferocytosis would be expected to clear apoptotic cells
and thereby stop, and even reverse, necrotic core expansion.

CONCLUDING REMARKS
During both the progression and the regression of atherosclerosis,
macrophages have central roles. While macrophage phenotypes

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Macrophage polarization in atherosclerosis

are diverse and form a continuum (90), we chose a simple and
dichotomous approach in order to emphasize the differences
between macrophage properties in atherosclerosis progression and
regression. M1 macrophages predominate in progression and contribute to the inflammatory state, whereas M2 macrophages are
enriched in many models of regression and appear to participate
in inflammation resolution and plaque remodeling. Interestingly,
M2 macrophages seem to have a beneficial role even when plasma
cholesterol levels remain high [e.g., Ref. (17, 86)], and the opposite is also true – M1 macrophages, though activated by direct and
indirect effects of cholesterol, have also been linked to an increase
in atherosclerosis progression despite similar levels of plasma cholesterol, for example, in diabetes (91). Thus, there are complex
interactions between macrophage phenotypes and plasma cholesterol levels, a situation likely to also exist with other known (and
yet to be discovered) risk factors.
As noted throughout this review, there are many areas in which
our knowledge of macrophage biology in plaques is inadequate.
Yet, it is already clear that the inflammatory state of these cells is
dynamically influenced by multiple metabolic, genetic, and pharmacologic factors. Deeper understanding of how these factors
effect changes in plaque macrophages will likely advance the development of new strategies to reduce the huge burden of cardiovascular morbidity and mortality that persists with existing therapies.

ACKNOWLEDGMENTS
We thank all of our lab members, past and present, as well as our
collaborators, who have contributed to our studies cited in this
review. Support for these studies comes from the following NIH
grants: HL098055, HL084312, DK95684, and HL117226.

REFERENCES
1. Williams KJ, Tabas I. The response-to-retention hypothesis of early
atherogenesis. Arterioscler Thromb Vasc Biol (1995) 15(5):551–61. doi:10.1161/
01.ATV.15.5.551
2. Ross R. Atherosclerosis – an inflammatory disease. N Engl J Med (1999)
340(2):115–26. doi:10.1056/NEJM199901143400207
3. Weber C, Noels H. Atherosclerosis: current pathogenesis and therapeutic
options. Nat Med (2011) 17(11):1410–22. doi:10.1038/nm.2538
4. Hansson GK, Hermansson A. The immune system in atherosclerosis. Nat
Immunol (2011) 12(3):204–12. doi:10.1038/ni.2001
5. Williams KJ, Feig JE, Fisher EA. Rapid regression of atherosclerosis: insights
from the clinical and experimental literature. Nat Clin Pract Cardiovasc Med
(2008) 5(2):91–102. doi:10.1038/ncpcardio1086
6. Epelman S, Lavine KJ, Randolph GJ. Origin and functions of tissue macrophages.
Immunity (2014) 41(1):21–35. doi:10.1016/j.immuni.2014.06.013
7. Distel E, Barrett TJ, Chung K, Girgis NM, Parathath S, Essau CC, et al. miR33
inhibition overcomes deleterious effects of diabetes mellitus on atherosclerosis plaque regression in mice. Circ Res (2014) 115(9):759–69. doi:10.1161/
CIRCRESAHA.115.304164
8. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
9. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism and
functions. Immunity (2010) 32(5):593–604. doi:10.1016/j.immuni.2010.05.007
10. Hanna RN, Shaked I, Hubbeling HG, Punt JA, Wu R, Herrley E, et al. NR4A1
(Nur77) deletion polarizes macrophages toward an inflammatory phenotype
and increases atherosclerosis. Circ Res (2012) 110(3):416–27. doi:10.1161/
CIRCRESAHA.111.253377
11. Davies MJ. Stability and instability: two faces of coronary atherosclerosis. The
Paul Dudley White lecture 1995. Circulation (1996) 94(8):2013–20. doi:10.1161/
01.CIR.94.8.2013

November 2014 | Volume 5 | Article 579 | 174

Peled and Fisher

12. Stary HC, Chandler AB, Dinsmore RE, Fuster V, Glagov S, Insull W Jr, et al. A
definition of advanced types of atherosclerotic lesions and a histological classification of atherosclerosis. A report from the committee on vascular lesions of
the council on arteriosclerosis, American heart association. Arterioscler Thromb
Vasc Biol (1995) 15(9):1512–31. doi:10.1161/01.ATV.15.9.1512
13. Virmani R, Kolodgie FD, Burke AP, Farb A, Schwartz SM. Lessons from sudden coronary death: a comprehensive morphological classification scheme for
atherosclerotic lesions. Arterioscler Thromb Vasc Biol (2000) 20(5):1262–75.
doi:10.1161/01.ATV.20.5.1262
14. Kolodgie FD, Narula J, Burke AP, Haider N, Farb A, Hui-Liang Y, et al.
Localization of apoptotic macrophages at the site of plaque rupture in sudden coronary death. Am J Pathol (2000) 157(4):1259–68. doi:10.1016/S00029440(10)64641-X
15. Tiemessen MM, Jagger AL, Evans HG, van Herwijnen MJ, John S, Taams LS.
CD4+CD25+Foxp3+ regulatory T cells induce alternative activation of human
monocytes/macrophages. Proc Natl Acad Sci U S A (2007) 104(49):19446–51.
doi:10.1073/pnas.0706832104
16. Feig JE, Parathath S, Rong JX, Mick SL, Vengrenyuk Y, Grauer L, et al. Reversal of
hyperlipidemia with a genetic switch favorably affects the content and inflammatory state of macrophages in atherosclerotic plaques. Circulation (2011)
123(9):989–98. doi:10.1161/CIRCULATIONAHA.110.984146
17. Feig JE, Rong JX, Shamir R, Sanson M, Vengrenyuk Y, Liu J, et al. HDL promotes rapid atherosclerosis regression in mice and alters inflammatory properties of plaque monocyte-derived cells. Proc Natl Acad Sci U S A (2011)
108(17):7166–71. doi:10.1073/pnas.1016086108
18. Kadl A, Meher AK, Sharma PR, Lee MY, Doran AC, Johnstone SR, et al. Identification of a novel macrophage phenotype that develops in response to atherogenic phospholipids via Nrf2. Circ Res (2010) 107(6):737–46. doi:10.1161/
CIRCRESAHA.109.215715
19. Stöger JL, Gijbels MJ, van der Velden S, Manca M, der Loos CM, Biessen EA, et al.
Distribution of macrophage polarization markers in human atherosclerosis.
Atherosclerosis (2012) 225(2):461–8. doi:10.1016/j.atherosclerosis.2012.09.013
20. Lu YC, Yeh WC, Ohashi PS. LPS/TLR4 signal transduction pathway. Cytokine
(2008) 42(2):145–51. doi:10.1016/j.cyto.2008.01.006
21. Cani PD, Amar J, Iglesias MA, Poggi M, Knauf C, Bastelica D, et al. Metabolic endotoxemia initiates obesity and insulin resistance. Diabetes (2007)
56(7):1761–72. doi:10.2337/db06-1491
22. Amar J, Burcelin R, Ruidavets JB, Cani PD, Fauvel J, Alessi MC, et al. Energy
intake is associated with endotoxemia in apparently healthy men. Am J Clin
Nutr (2008) 87(5):1219–23.
23. Cani PD, Bibiloni R, Knauf C, Waget A, Neyrinck AM, Delzenne NM, et al.
Changes in gut microbiota control metabolic endotoxemia-induced inflammation in high-fat diet-induced obesity and diabetes in mice. Diabetes (2008)
57(6):1470–81. doi:10.2337/db07-1403
24. Arbour NC, Lorenz E, Schutte BC, Zabner J, Kline JN, Jones M, et al. TLR4
mutations are associated with endotoxin hyporesponsiveness in humans. Nat
Genet (2000) 25(2):187–91. doi:10.1038/76048
25. Kiechl S, Lorenz E, Reindl M, Wiedermann CJ, Oberhollenzer F, Bonora E, et al.
Toll-like receptor 4 polymorphisms and atherogenesis. N Engl J Med (2002)
347(3):185–92. doi:10.1056/NEJMoa012673
26. Kiechl S, Wiedermann CJ, Willeit J. Toll-like receptor 4 and atherogenesis. Ann
Med (2003) 35(3):164–71. doi:10.1080/07853890310008215
27. Ameziane N, Beillat T, Verpillat P, Chollet-Martin S, Aumont MC, Seknadji
P, et al. Association of the toll-like receptor 4 gene Asp299Gly polymorphism
with acute coronary events. Arterioscler Thromb Vasc Biol (2003) 23(12):e61–4.
doi:10.1161/01.ATV.0000101191.92392.1D
28. Michelsen KS, Wong MH, Shah PK, Zhang W, Yano J, Doherty TM, et al. Lack of
toll-like receptor 4 or myeloid differentiation factor 88 reduces atherosclerosis
and alters plaque phenotype in mice deficient in apolipoprotein E. Proc Natl
Acad Sci U S A (2004) 101(29):10679–84. doi:10.1073/pnas.0403249101
29. Stepankova R, Tonar Z, Bartova J, Nedorost L, Rossman P, Poledne R, et al.
Absence of microbiota (germ-free conditions) accelerates the atherosclerosis
in ApoE-deficient mice fed standard low cholesterol diet. J Atheroscler Thromb
(2010) 17(8):796–804. doi:10.5551/jat.3285
30. Sokol H, Pigneur B, Watterlot L, Lakhdari O, Bermúdez-Humarán LG, Gratadoux JJ, et al. Faecalibacterium prausnitzii is an anti-inflammatory commensal
bacterium identified by gut microbiota analysis of Crohn disease patients. Proc
Natl Acad Sci U S A (2008) 105(43):16731–6. doi:10.1073/pnas.0804812105

www.frontiersin.org

Macrophage polarization in atherosclerosis

31. Duewell P, Kono H, Rayner KJ, Sirois CM, Vladimer G, Bauernfeind
FG, et al. NLRP3 inflammasomes are required for atherogenesis and activated by cholesterol crystals. Nature (2010) 464(7293):1357–61. doi:10.1038/
nature08938
32. Lim RS, Suhalim JL, Miyazaki-Anzai S, Miyazaki M, Levi M, Potma EO, et al.
Identification of cholesterol crystals in plaques of atherosclerotic mice using
hyperspectral CARS imaging. J Lipid Res (2011) 52(12):2177–86. doi:10.1194/
jlr.M018077
33. Sheedy FJ, Grebe A, Rayner KJ, Kalantari P, Ramkhelawon B, Carpenter SB, et al.
CD36 coordinates NLRP3 inflammasome activation by facilitating intracellular
nucleation of soluble ligands into particulate ligands in sterile inflammation.
Nat Immunol (2013) 14(8):812–20. doi:10.1038/ni.2639
34. Kirii H, Niwa T, Yamada Y, Wada H, Saito K, Iwakura Y, et al. Lack of
interleukin-1beta decreases the severity of atherosclerosis in ApoE-deficient
mice. Arterioscler Thromb Vasc Biol (2003) 23(4):656–60. doi:10.1161/01.ATV.
0000064374.15232.C3
35. Miller YI, Choi SH, Wiesner P, Fang L, Harkewicz R, Hartvigsen K, et al.
Oxidation-specific epitopes are danger-associated molecular patterns recognized by pattern recognition receptors of innate immunity. Circ Res (2011)
108(2):235–48. doi:10.1161/CIRCRESAHA.110.223875
36. Kim TW, Febbraio M, Robinet P, Dugar B, Greene D, Cerny A, et al. The critical role of IL-1 receptor-associated kinase 4-mediated NF-kappaB activation
in modified low-density lipoprotein-induced inflammatory gene expression
and atherosclerosis. J Immunol (2011) 186(5):2871–80. doi:10.4049/jimmunol.
1002242
37. Rekhter M, Staschke K, Estridge T, Rutherford P, Jackson N, Gifford-Moore D,
et al. Genetic ablation of IRAK4 kinase activity inhibits vascular lesion formation. Biochem Biophys Res Commun (2008) 367(3):642–8. doi:10.1016/j.bbrc.
2007.12.186
38. Björkbacka H, Kunjathoor VV, Moore KJ, Koehn S, Ordija CM, Lee MA, et al.
Reduced atherosclerosis in MyD88-null mice links elevated serum cholesterol
levels to activation of innate immunity signaling pathways. Nat Med (2004)
10(4):416–21. doi:10.1038/nm1008
39. Mullick AE, Tobias PS, Curtiss LK. Modulation of atherosclerosis in mice by
toll-like receptor 2. J Clin Invest (2005) 115(11):3149–56. doi:10.1172/JCI25482
40. Yvan-Charvet L, Welch C, Pagler TA, Ranalletta M, Lamkanfi M, Han S,
et al. Increased inflammatory gene expression in ABC transporter-deficient
macrophages: free cholesterol accumulation, increased signaling via toll-like
receptors, and neutrophil infiltration of atherosclerotic lesions. Circulation
(2008) 118(18):1837–47. doi:10.1161/CIRCULATIONAHA.108.793869
41. Zhu X, Lee JY, Timmins JM, Brown JM, Boudyguina E, Mulya A, et al.
Increased cellular free cholesterol in macrophage-specific Abca1 knock-out
mice enhances pro-inflammatory response of macrophages. J Biol Chem (2008)
283(34):22930–41. doi:10.1074/jbc.M801408200
42. Chawla A. Control of macrophage activation and function by PPARs. Circ Res
(2010) 106(10):1559–69. doi:10.1161/CIRCRESAHA.110.216523
43. Spann NJ, Garmire LX, McDonald JG, Myers DS, Milne SB, Shibata N, et al.
Regulated accumulation of desmosterol integrates macrophage lipid metabolism and inflammatory responses. Cell (2012) 151(1):138–52. doi:10.1016/j.cell.
2012.06.054
44. Galkina E, Kadl A, Sanders J, Varughese D, Sarembock IJ, Ley K. Lymphocyte
recruitment into the aortic wall before and during development of atherosclerosis is partially L-selectin dependent. J Exp Med (2006) 203(5):1273–82.
doi:10.1084/jem.20052205
45. Koltsova EK, Garcia Z, Chodaczek G, Landau M, McArdle S, Scott SR, et al.
Dynamic T cell-APC interactions sustain chronic inflammation in atherosclerosis. J Clin Invest (2012) 122(9):3114–26. doi:10.1172/JCI61758
46. Molnár E, Swamy M, Holzer M, Beck-García K, Worch R, Thiele C, et al. Cholesterol and sphingomyelin drive ligand-independent T-cell antigen receptor
nanoclustering. J Biol Chem (2012) 287(51):42664–74. doi:10.1074/jbc.M112.
386045
47. Emeson EE, Shen ML, Bell CG, Qureshi A. Inhibition of atherosclerosis in CD4
T-cell-ablated and nude (nu/nu) C57BL/6 hyperlipidemic mice. Am J Pathol
(1996) 149(2):675–85.
48. Dansky HM, Charlton SA, Harper MM, Smith JD. T and B lymphocytes play
a minor role in atherosclerotic plaque formation in the apolipoprotein Edeficient mouse. Proc Natl Acad Sci U S A (1997) 94(9):4642–6. doi:10.1073/
pnas.94.9.4642

November 2014 | Volume 5 | Article 579 | 175

Peled and Fisher

49. Odegaard JI, Chawla A. Alternative macrophage activation and metabolism.
Annu Rev Pathol (2011) 6:275–97. doi:10.1146/annurev-pathol-011110-130138
50. Thorp E, Tabas I. Mechanisms and consequences of efferocytosis in advanced
atherosclerosis. J Leukoc Biol (2009) 86(5):1089–95. doi:10.1189/jlb.0209115
51. Moulton KS, Vakili K, Zurakowski D, Soliman M, Butterfield C, Sylvin E, et al.
Inhibition of plaque neovascularization reduces macrophage accumulation and
progression of advanced atherosclerosis. Proc Natl Acad Sci U S A (2003)
100(8):4736–41. doi:10.1073/pnas.0730843100
52. Chinetti-Gbaguidi G, Baron M, Bouhlel MA, Vanhoutte J, Copin C, Sebti
Y, et al. Human atherosclerotic plaque alternative macrophages display low
cholesterol handling but high phagocytosis because of distinct activities of
the PPARgamma and LXRalpha pathways. Circ Res (2011) 108(8):985–95.
doi:10.1161/CIRCRESAHA.110.233775
53. Khallou-Laschet J,Varthaman A, Fornasa G, Compain C, Gaston AT, Clement M,
et al. Macrophage plasticity in experimental atherosclerosis. PLoS One (2010)
5(1):e8852. doi:10.1371/journal.pone.0008852
54. Davies LC, Jenkins SJ, Allen JE, Taylor PR. Tissue-resident macrophages. Nat
Immunol (2013) 14(10):986–95. doi:10.1038/ni.2705
55. Robbins CS, Hilgendorf I, Weber GF, Theurl I, Iwamoto Y, Figueiredo JL, et al.
Local proliferation dominates lesional macrophage accumulation in atherosclerosis. Nat Med (2013) 19(9):1166–72. doi:10.1038/nm.3258
56. Maitra U, Parks JS, Li L. An innate immunity signaling process suppresses
macrophage ABCA1 expression through IRAK-1-mediated downregulation of
retinoic acid receptor alpha and NFATc2. Mol Cell Biol (2009) 29(22):5989–97.
doi:10.1128/MCB.00541-09
57. Devaraj S, Jialal I. C-reactive protein polarizes human macrophages to an
M1 phenotype and inhibits transformation to the M2 phenotype. Arterioscler
Thromb Vasc Biol (2011) 31(6):1397–402. doi:10.1161/ATVBAHA.111.225508
58. Iwasaki M, Mukai T, Gao P, Park WR, Nakajima C, Tomura M, et al. A critical
role for IL-12 in CCR5 induction on T cell receptor-triggered mouse CD4(+)
and CD8(+) T cells. Eur J Immunol (2001) 31(8):2411–20. doi:10.1002/15214141(200108)31:8<2411::AID-IMMU2411>3.0.CO;2-Y
59. Huang WC, Sala-Newby GB, Susana A, Johnson JL, Newby AC. Classical
macrophage activation up-regulates several matrix metalloproteinases through
mitogen activated protein kinases and nuclear factor-kappaB. PLoS One (2012)
7(8):e42507. doi:10.1371/journal.pone.0042507
60. Holman RL, McGILL HC Jr, STRONG JP, GEER JC. The natural history of atherosclerosis: the early aortic lesions as seen in New Orleans in the middle of the
20th century. Am J Pathol (1958) 34(2):209–35.
61. McMahan CA, Gidding SS, McGill HC Jr. Coronary heart disease risk factors and atherosclerosis in young people. J Clin Lipidol (2008) 2(3):118–26.
doi:10.1016/j.jacl.2008.02.006
62. Trogan E, Feig JE, Dogan S, Rothblat GH, Angeli V, Tacke F, et al. Gene expression
changes in foam cells and the role of chemokine receptor CCR7 during atherosclerosis regression in ApoE-deficient mice. Proc Natl Acad Sci U S A (2006)
103(10):3781–6. doi:10.1073/pnas.0511043103
63. Chereshnev I, Trogan E, Omerhodzic S, Itskovich V, Aguinaldo JG, Fayad ZA,
et al. Mouse model of heterotopic aortic arch transplantation. J Surg Res (2003)
111(2):171–6. doi:10.1016/S0022-4804(03)00039-8
64. Lieu HD, Withycombe SK, Walker Q, Rong JX, Walzem RL, Wong JS, et al. Eliminating atherogenesis in mice by switching off hepatic lipoprotein secretion.
Circulation (2003) 107(9):1315–21. doi:10.1161/01.CIR.0000054781.50889.0C
65. Potteaux S, Gautier EL, Hutchison SB, van Rooijen N, Rader DJ, Thomas MJ,
et al. Suppressed monocyte recruitment drives macrophage removal from atherosclerotic plaques of Apoe−/− mice during disease regression. J Clin Invest
(2011) 121(5):2025–36. doi:10.1172/JCI43802
66. Van Craeyveld E, Gordts SC, Nefyodova E, Jacobs F, De Geest B. Regression
and stabilization of advanced murine atherosclerotic lesions: a comparison of
LDL lowering and HDL raising gene transfer strategies. J Mol Med (Berl) (2011)
89(6):555–67. doi:10.1007/s00109-011-0722-x
67. Raffai RL, Loeb SM, Weisgraber KH. Apolipoprotein E promotes the regression of atherosclerosis independently of lowering plasma cholesterol levels. Arterioscler Thromb Vasc Biol (2005) 25(2):436–41. doi:10.1161/01.ATV.
0000152613.83243.12
68. Hewing B, Parathath S, Mai CK, Fiel MI, Guo L, Fisher EA. Rapid regression of atherosclerosis with MTP inhibitor treatment. Atherosclerosis (2013)
227(1):125–9. doi:10.1016/j.atherosclerosis.2012.12.026
69. Tangirala RK, Tsukamoto K, Chun SH, Usher D, Puré E, Rader DJ.
Regression of atherosclerosis induced by liver-directed gene transfer of

Frontiers in Immunology | Molecular Innate Immunity

Macrophage polarization in atherosclerosis

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

apolipoprotein A-I in mice. Circulation (1999) 100(17):1816–22. doi:10.1161/
01.CIR.100.17.1816
Reis ED, Li J, Fayad ZA, Rong JX, Hansoty D, Aguinaldo JG, et al. Dramatic remodeling of advanced atherosclerotic plaques of the apolipoprotein E-deficient mouse in a novel transplantation model. J Vasc Surg (2001)
34(3):541–7. doi:10.1067/mva.2001.115963
Rong JX, Li J, Reis ED, Choudhury RP, Dansky HM, Elmalem VI, et al.
Elevating high-density lipoprotein cholesterol in apolipoprotein E-deficient
mice remodels advanced atherosclerotic lesions by decreasing macrophage and
increasing smooth muscle cell content. Circulation (2001) 104(20):2447–52.
doi:10.1161/hc4501.098952
Zhao Y, Black AS, Bonnet DJ, Maryanoff BE, Curtiss LK, Leman LJ, et al.
In vivo efficacy of HDL-like nanolipid particles containing multivalent peptide mimetics of apolipoprotein A-I. J Lipid Res (2014) 55(10):2053–63.
doi:10.1194/jlr.M049262
Hewing B, Parathath S, Barrett T, Chung WK, Astudillo YM, Hamada T, et al.
Effects of native and myeloperoxidase-modified apolipoprotein a-I on reverse
cholesterol transport and atherosclerosis in mice. Arterioscler Thromb Vasc Biol
(2014) 34(4):779–89. doi:10.1161/ATVBAHA.113.303044
Rayner KJ, Suárez Y, Dávalos A, Parathath S, Fitzgerald ML, Tamehiro N, et al.
MiR-33 contributes to the regulation of cholesterol homeostasis. Science (2010)
328(5985):1570–3. doi:10.1126/science.1189862
Rayner KJ, Sheedy FJ, Esau CC, Hussain FN, Temel RE, Parathath S, et al. Antagonism of miR-33 in mice promotes reverse cholesterol transport and regression
of atherosclerosis. J Clin Invest (2011) 121(7):2921–31. doi:10.1172/JCI57275
Olofsson SO, Asp L, Boren J. The assembly and secretion of apolipoprotein
B-containing lipoproteins. Curr Opin Lipidol (1999) 10(4):341–6. doi:10.1097/
00041433-199908000-00008
Ritter T, Lehmann M, Volk HD. Improvements in gene therapy: averting
the immune response to adenoviral vectors. BioDrugs (2002) 16(1):3–10.
doi:10.2165/00063030-200216010-00001
Nagareddy PR, Murphy AJ, Stirzaker RA, Hu Y, Yu S, Miller RG, et al. Hyperglycemia promotes myelopoiesis and impairs the resolution of atherosclerosis.
Cell Metab (2013) 17(5):695–708. doi:10.1016/j.cmet.2013.04.001
Parathath S, Grauer L, Huang LS, Sanson M, Distel E, Goldberg IJ, et al. Diabetes
adversely affects macrophages during atherosclerotic plaque regression in mice.
Diabetes (2011) 60(6):1759–69. doi:10.2337/db10-0778
Llodrá J, Angeli V, Liu J, Trogan E, Fisher EA, Randolph GJ. Emigration of
monocyte-derived cells from atherosclerotic lesions characterizes regressive, but
not progressive, plaques. Proc Natl Acad Sci U S A (2004) 101(32):11779–84.
doi:10.1073/pnas.0403259101
Sanson M, Distel E, Fisher EA. HDL induces the expression of the M2
macrophage markers arginase 1 and Fizz-1 in a STAT6-dependent process. PLoS
One (2013) 8(8):e74676. doi:10.1371/journal.pone.0074676
Colin S, Fanchon M, Belloy L, Bochem AE, Copin C, Derudas B, et al. HDL
does not influence the polarization of human monocytes toward an alternative
phenotype. Int J Cardiol (2014) 172(1):179–84. doi:10.1016/j.ijcard.2013.12.168
De Nardo D, Labzin LI, Kono H, Seki R, Schmidt SV, Beyer M, et al. High-density
lipoprotein mediates anti-inflammatory reprogramming of macrophages via the
transcriptional regulator ATF3. Nat Immunol (2014) 15(2):152–60. doi:10.1038/
ni.2784
Wilhelm AJ, Zabalawi M, Owen JS, Shah D, Grayson JM, Major AS, et al.
Apolipoprotein A-I modulates regulatory T cells in autoimmune LDLr−/−,
ApoA-I−/− mice. J Biol Chem (2010) 285(46):36158–69. doi:10.1074/jbc.M110.
134130
Murphy AJ, Westerterp M, Yvan-Charvet L, Tall AR. Anti-atherogenic mechanisms of high density lipoprotein: effects on myeloid cells. Biochim Biophys Acta
(2012) 1821(3):513–21. doi:10.1016/j.bbalip.2011.08.003
Cardilo-Reis L, Gruber S, Schreier SM, Drechsler M, Papac-Milicevic N, Weber
C, et al. Interleukin-13 protects from atherosclerosis and modulates plaque
composition by skewing the macrophage phenotype. EMBO Mol Med (2012)
4(10):1072–86. doi:10.1002/emmm.201201374
Schnoor M, Cullen P, Lorkowski J, Stolle K, Robenek H, Troyer D, et al.
Production of type VI collagen by human macrophages: a new dimension
in macrophage functional heterogeneity. J Immunol (2008) 180(8):5707–19.
doi:10.4049/jimmunol.180.8.5707
Zizzo G, Hilliard BA, Monestier M, Cohen PL. Efficient clearance of early
apoptotic cells by human macrophages requires M2c polarization and MerTK
induction. J Immunol (2012) 189(7):3508–20. doi:10.4049/jimmunol.1200662

November 2014 | Volume 5 | Article 579 | 176

Peled and Fisher

89. Feng B, Yao PM, Li Y, Devlin CM, Zhang D, Harding HP, et al. The endoplasmic
reticulum is the site of cholesterol-induced cytotoxicity in macrophages. Nat
Cell Biol (2003) 5(9):781–92. doi:10.1038/ncb1035
90. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
91. Vikramadithyan RK, Hu Y, Noh HL, Liang CP, Hallam K, Tall AR, et al. Human
aldose reductase expression accelerates diabetic atherosclerosis in transgenic
mice. J Clin Invest (2005) 115(9):2434–43. doi:10.1172/JCI24819

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.

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Macrophage polarization in atherosclerosis

Received: 25 September 2014; accepted: 29 October 2014; published online: 12
November 2014.
Citation: Peled M and Fisher EA (2014) Dynamic aspects of macrophage polarization during atherosclerosis progression and regression. Front. Immunol. 5:579. doi:
10.3389/fimmu.2014.00579
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Peled and Fisher. This is an open-access article distributed under the
terms of the Creative Commons Attribution License (CC BY). The use, distribution or
reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
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REVIEW ARTICLE
published: 27 October 2014
doi: 10.3389/fimmu.2014.00533

Functions of arginase isoforms in macrophage
inflammatory responses: impact on cardiovascular
diseases and metabolic disorders
Zhihong Yang* and Xiu-Fen Ming*
Vascular Biology, Division of Physiology, Department of Medicine, Faculty of Science, University of Fribourg, Fribourg, Switzerland

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Dalil Hannani, APCure SAS, France
Sruti Shiva, University of Pittsburgh,
USA
*Correspondence:
Zhihong Yang and Xiu-Fen Ming,
Vascular Biology, Division of
Physiology, Department of Medicine,
Faculty of Science, University of
Fribourg, Chemin du Musée 5,
Fribourg CH-1700, Switzerland
e-mail: zhihong.yang@unifr.ch;
xiu-fen.ming@unifr.ch

Macrophages play a paramount role in immunity and inflammation-associated diseases,
including infections, cardiovascular diseases, obesity-associated metabolic imbalances,
and cancer. Compelling evidence from studies of recent years demonstrates that
macrophages are heterogeneous and undergo heterogeneous phenotypic changes in
response to microenvironmental stimuli. The M1 killer type response and the M2 repair
type response are best known, and are two extreme examples. Among other markers,
inducible nitric oxide synthase and type-I arginase (Arg-I), the enzymes that are involved
in L-arginine/nitric oxide (NO) metabolism, are associated with the M1 and M2 phenotype, respectively, and therefore widely used as the markers for characterization of the
two macrophage phenotypes. There is also a type-II arginase (Arg-II), which is expressed
in macrophages and prevalently viewed as having the same function as Arg-I in the cells.
In contrast to Arg-I, little information on the role of Arg-II in macrophage inflammatory
responses is available. Emerging evidence, however, suggests that differential roles of Arg-I
and Arg-II in regulating macrophage functions. In this article, we will review recent developments on the functional roles of the two arginase isoforms in regulation of macrophage
inflammatory responses by focusing on their impact on the pathogenesis of cardiovascular
diseases and metabolic disorders.
Keywords: arginase, arginine, macrophages, nitric oxide synthase, cardiovascular diseases

INTRODUCTION
Macrophages are important sentinel cells in our body and
are involved in maintenance of tissue homeostasis, immune
responses, and inflammation-associated diseases. Recent findings
have revised our traditional view on the origin and biological functions of macrophages. We now know that tissue macrophages are
not only recruited from bone marrow-derived monocytes but also
differentiated from yolk sac-derived embryonic stem cells (1–3).
Moreover, tissue macrophages are not terminally differentiated
and are maintained throughout life by local proliferative selfrenewal (4, 5). Importantly, macrophages are highly heterogeneous
and undergo phenotypic changes, i.e., macrophage plasticity, in
response to specific signals as a consequence of adaptation to
local tissue environmental cues (6, 7). The original and the best
known types of macrophage responses are the pro-inflammatory
M1 type (killer cells) and the anti-inflammatory M2 type (repair
type cells) (3, 7). There are convincing evidences from research
of recent years showing that different phenotypic macrophages
are indeed importantly participating in the process of immune
and inflammatory responses, which have been reviewed by many
comprehensive articles (7, 8).

MACROPHAGE POLARIZATION
Macrophage polarization describes acquirement of distinctive
phenotypic and functional characteristics of fully differentiated
macrophages in response to microenvironmental stimuli.
Functional polarization of macrophages and the underlying
Frontiers in Immunology | Molecular Innate Immunity

mechanisms that control the cell phenotypes are complex and
have been extensively investigated in recent years. As mentioned,
the M1 and M2 classifications of macrophages described the two
major and opposing activities committed to killing and repairing
functions of the cells. It is emerging that macrophage polarization is regulated by a broad spectrum of recognition receptors, cytokines, specific signaling pathways, and genetic programs.
Some of them are used as markers or functional repertoire of the
macrophage phenotypes. There are, however, no standard guidelines for classification of macrophage phenotypes. Most importantly, information about functions of these markers in regulation
of macrophage inflammatory responses or phenotypes is either
lacking or controversial. The conclusions are usually based on
association studies. It is generally the view that M1 macrophages
express enhanced genes, which are pro-inflammatory and cytotoxic, typically inducible nitric oxide synthase (iNOS)/NO, IL-12,
class II MHC, and the chemokines IL-8 and CCL2, participating in killing intracellular parasites and tumor development.
In contrast, M2 macrophages produce more anti-inflammatory
cytokines and substances involved in repairing function, typically, arginase/ornithine, EGF, VEGF, and TGF-β, and mannose
receptor (9). This phenotypic cell is mainly participating in resolution of inflammation, tissue repairing, angiogenesis, allergy, and
tumor progression (10). It is, however, to notice that M1 and M2
activation programs display differences, but they may not form
clear-cut activation subsets and reveal overlapping effects. A discussion about the complexity of macrophage phenotype markers,
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Arginase and macrophage functions

differentiation mechanisms, and the roles in human diseases is
beyond the scope of this review article. For these aspects, readers
are kindly asked to refer to several comprehensive review articles
(11, 12). In the following section of this article, we will focus on discussing the role of the enzymes arginase and nitric oxide synthase
(NOS) that are involved in l-arginine metabolism in various cell
types including vascular endothelial cells and macrophages and
widely used as markers to distinguish M1 and M2 macrophage
phenotypes.

encoded by two separate genes. In human beings, Arg-I gene
maps to chromosome 6q23 and encodes a 322 amino acid protein (19–21), while Arg-II gene maps to chromosome 14q24.1
and encodes a 354 amino acid protein (22–24). At the subcellular level, Arg-I is mainly localized in cytoplasm and Arg-II in
mitochondrion (25). The physiological role of the different subcellular compartmentation of the two isoenzymes is not known.
The two isoenzymes, however, share similar structure, reveal more
than 50% of homology of their amino acid residues with 100%
homology in the areas, which are critical for their l-arginine
metabolizing function (22, 23, 26). Although both Arg-I and ArgL-ARGININE METABOLISM, iNOS, AND ARGINASE IN
II are to hydrolyze l-arginine to produce urea and l-ornithine
MACROPHAGE FUNCTIONAL POLARIZATION
The suggestion that l-arginine metabolism could be involved in (25), the functional impact of the two isoenzymes is either simregulation of macrophage phenotypes was from early studies with ilar or different depending on specific organs/cells. For example,
macrophages isolated from the Th1 strain mouse C57B1/6 and increased activity and/or expression of either Arg-I or Arg-II in
Th2 strain BALB/c mouse (13, 14). These studies demonstrate that endothelial cells impair the vasoprotective endothelial NO producisolated macrophages from Th1 strain C57Bl/6 mouse are more tion via eNOS (27). However, in macrophages, Arg-I and Arg-II
readily activated to produce nitric oxide (NO) upon stimulation seem to play an opposite function, which we will discuss later in
with IFN-γ or lipopolysaccharide (LPS) than the macrophages this article. The primary function of Arg-I is to remove excesfrom Th2 strain BALB/c mouse. Later on, it was characterized sive nitrogen produced from amino acid metabolism through
that M1 macrophages are more easily activated by LPS to produce hepatic urea cycle, which is otherwise toxic for our body (28,
cytotoxic NO via iNOS, whereas M2 macrophages generate lit- 29), because Arg-I is constitutively and abundantly expressed as
tle NO but more ornithine from the same substrate l-arginine via a cytosolic enzyme in the liver (30). No Arg-II could be detected
arginase (15). The iNOS and arginase are thought to affect inflam- in hepatocytes. The vital effect of hepatic Arg-I is evidenced by the
matory responses in the opposite way. NO production from iNOS studies showing that Arg-I knockout mice reveal severe sympin M1 macrophages inhibits cell proliferation and kills pathogens, toms of hyperammonemia and die between postnatal days 10
a M1 killing type response (16, 17), while ornithine production and 14 (31). Patients with Arg-I deficiency due to gene mutation
promotes cell proliferation and repairs tissue damage through reveal urea cycle disorder, hyperargininemia, and exhibit progresgeneration of polyamines and collagen in M2 macrophages, a M2 sive neurologic impairment, development retardation, and hepatic
repairing type response (11, 18). Both NO and ornithine are gen- dysfunction associated with cirrhosis and carcinoma in early childerated from the same substrate l-arginine via iNOS and arginase, hood (28, 29). Arg-I has been reported to be expressed also in
respectively (11, 18) (Figure 1). From these studies, one can con- many extrahepatic tissues such as stomach, pancreas, and lung
sider dominant NO production as M1 activity, whereas dominant (32). The functions of Arg-I in these organs are far from clear.
Unlike Arg-I, Arg-II is confined mainly to kidney, brain, prostate,
ornithine production as M2 activity of macrophages.
intestine, and also pancreas (22, 23, 32). The functions of ArgII in these organs are not known. The best characterization of
ARGINASE ISOENZYMES AND L-ARGININE METABOLIZING
Arg-II function is done in vascular endothelial cells in which the
FUNCTIONS
In human beings and mammals, there are two isoforms of arginases, isoenzyme, similar to Arg-I, metabolizes l-arginine to urea and larginase-1 (Arg-I) and arginase-II (Arg-II). Both isoenzymes are ornithine, which limits l-arginine bioavailability for generation of
the vasoprotective NO via eNOS, resulting in vascular endothelial
dysfunction (33, 34). This effect of arginases on endothelial cells is
attributable to eNOS-uncoupling, a situation that eNOS enzyme
produces increased superoxide anion, but decreased NO (34–39),
which is thought to be attributed to l-arginine deficiency, leading
to oxidative stress, and enhanced expression of endothelial inflammatory adhesion molecules such as VCAM-1 and ICAM-1 (39),
since endothelial NO reveals important anti-oxidative and antiinflammatory functions and suppresses expression of the adhesion
molecules (40). These effects of Arg-II are dependent on the enzymatic activity, since loss-of-function point mutation of histidine
to phenylalanine at position 160 in Arg-II abolishes its l-arginine–
urea hydrolase activity and is unable to cause eNOS-uncoupling
and the inflammatory responses in endothelial cells (39). We have
recently reviewed the aspect of arginase in eNOS-uncoupling (41).
FIGURE 1 | L-arginine metabolism by iNOS and arginase and the
Arginase also exerts pleiotropic effects, i.e., l-arginine–urea hydrofunctional consequences in macrophages. ODC, ornithine
lase activity-independent effect, which we will discuss later in this
decarboxylase; OAT, ornithine aminotransferase.
article.

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Yang and Ming

ARGINASE ISOENZYMES AND MACROPHAGE FUNCTIONS
As mentioned above, macrophages are heterogeneous and undergo
phenotypic changes, depending on microenvironmental stimuli.
The expression of Arg-I and Arg-II is inducible in macrophages
depending on external stimuli (42, 43). As discussed, NO from
iNOS in macrophages is linked to M1, whereas ornithine generated
from arginase is associated with M2 phenotype (11). Substantial number of studies demonstrates that Arg-I is dominantly
expressed in M2 cells and reduces NO production from iNOS
through limiting bioavailability of intracellular l-arginine, resulting in dampening of inflammatory tissue damage and suppression
of clearance of intracellular pathogens (44–48). In contrast to ArgI, only very little and even contradictory information is available
about the expression and role of Arg-II in macrophage phenotype
regulation and inflammatory responses. Until we have systematically investigated this specific aspect in macrophage inflammatory
responses (43), the function of Arg-II in macrophages is believed
to be anti-inflammatory, which is extrapolated from its similar
function as Arg-I on l-arginine/NO metabolism. An early study
showed that Arg-II gene is a direct target of liver X receptor that
has been shown to exert inhibitory effects on expression of inflammatory genes in macrophages (49). Based on this association, the
authors suggest that Arg-II is anti-inflammatory. The functional
analysis is, however, not done. It is of particular importance to note
that LPS stimulation exclusively enhances iNOS in macrophages
associated with M1 phenotype (43, 50). We could demonstrate
that iNOS induction in macrophages is paralleled with enhanced
expression of Arg-II, but not Arg-I (43), which suggests that
Arg-I and Arg-II shall have different functions in macrophage
inflammatory responses or phenotype regulation. In line with this
observation, accumulation of Arg-II-expressing macrophages is
associated with advanced atherosclerotic lesions in which proinflammatory cells are dominant (42), suggesting that Arg-II is
associated with pro-inflammatory responses. Because of this contradictory concept about the role of Arg-II and lack of functional
analysis of Arg-II in macrophage inflammatory responses, we
recently systematically characterized the role of Arg-II in regulation of macrophage inflammations at the cellular and whole body
levels in mouse models of chronic inflammatory diseases such
as obesity-linked insulin resistance, type-II diabetes mellitus, and
atherosclerosis (43).
In this study, we demonstrate that M1 activation of
macrophages by LPS exclusively up-regulates iNOS and Arg-II,
but not Arg-I expression in murine and human macrophages
(43). Silencing Arg-II gene in human monocyte/macrophage cell
lines decreases the cell adhesion to endothelial cells with reduced
production of pro-inflammatory cytokines in response to LPS or
ox-LDL at both the mRNA and protein levels. Moreover, LPSinduced up-regulation of numerous pro-inflammatory mediators, including MCP-1, TNF-α, IL-6, MMP14, and iNOS, is significantly suppressed in macrophages isolated from Arg-II−/−
mice as compared with those from wild-type control animals.
Convincingly, introducing Arg-II gene back to the Arg-II−/−
macrophages restores or enhances the LPS-stimulated expression
of the pro-inflammatory genes to much higher levels compared to
the Arg-II+/+ cells from wild-type mice. Importantly, Arg-II−/−
mice are protected from systemic pro-inflammatory macrophage

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Arginase and macrophage functions

infiltration in various organs and expression of pro-inflammatory
mediators in high-fat diet (HFD)-induced obesity. Arg-II−/− mice,
when fed a HFD, although have similar body weight as WT controls, reveal lower fasting plasma glucose concentration, are more
glucose tolerant and insulin sensitive (43) as compared to WT
mice on HFD. Interestingly, Arg-II levels in macrophages are significantly increased in WT mice fed HFD, which is associated
with pro-inflammatory responses. The pro-inflammatory function of Arg-II in macrophages is further demonstrated in another
chronic inflammatory disease model, i.e., atherosclerosis mouse
model (43). Knocking-out Arg-II gene in the atherosclerosisprone ApoE−/− mice (ApoE−/− /Arg-II−/− ) decreases inflammatory cytokine levels and macrophage content in the aortas,
reduces atherosclerotic plaque formation, and reveals more stable plaque features as compared to ApoE−/− Arg-II+/+ control
mice. Since M1 pro-inflammatory macrophages play crucial role
in development of insulin resistance and type-II diabetes and
atherogenesis (51–55), our results demonstrate that Arg-II promotes pro-inflammatory or M1 phenotype of macrophages and
favors chronic inflammatory disease development such as obesityassociated insulin resistance, type-II diabetes, and atherosclerosis.
It is to mention that the pro-inflammatory effect of Arg-II in
macrophages does not seem to be relying on iNOS, since inhibition
of iNOS does not significantly affect expression of several proinflammatory genes in macrophages. The dissociation of arginase
activity from iNOS has been reported by several studies, showing
that alteration of arginase activity in macrophages is not necessarily associated with functional changes in iNOS (56–59). These
iNOS-independent pro-inflammatory responses mediated by ArgII in macrophages are due to enhanced mitochondrial ROS, since
reintroduction of the Arg-II gene into Arg-II−/− macrophages
enhances mitochondrial O−
2 and H2 O2 generation and inhibition of mitochondrial ROS significantly reduces Arg-II-mediated
inflammatory responses. The function of Arg-II in comparison
with Arg-I in macrophage inflammatory responses and chronic
inflammatory diseases, i.e., atherosclerosis and insulin resistance
is summarized in Figure 2. It is not very surprising, since Arg-II is
a mitochondrial enzyme (60). The question remains elusive how
Arg-II affects mitochondrial function leading to mitochondrial
ROS production in macrophages.
Although we have characterized the function of Arg-II in
macrophages, many important questions remain unanswered. In
the following section, we will briefly discuss several important
remaining questions regarding the role of arginase isoenzymes in
macrophage functions.

FUTURE RESEARCH QUESTIONS AND PERSPECTIVES
DOES Arg-I PLAY A CAUSAL ROLE IN M2 MACROPHAGE PHENOTYPE?

As discussed, Arg-I is constitutively expressed in hepatocytes and
is inducible in macrophages, e.g., by Th2 cytokines IL-4 and
IL-13 (44, 61, 62). It is highly upregulated in M2 macrophages
and widely used as a M2 macrophage marker (11). It has been
suggested that Arg-I in macrophages promotes Th2 cytokine production, contributing to resolution of inflammation and tissue
repairing (63). A study in human smooth muscle cells showing
that overexpression of Arg-I gene is able to decrease LPS-induced
pro-inflammatory cytokine production (64), seems to support the

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FIGURE 2 | The distinct role of Arg-I and Arg-II in macrophage
inflammatory responses. Dashed line indicates that the causal role of
Arg-I in M2 phenotype requires determination.

anti-inflammatory function of Arg-I. However, mice with specific Arg-I gene deficiency in macrophages show exacerbated Th2
response and fibrosis in the liver of Schistosoma mansoni infected
mice (65), which does not seem to support previous suggestions
in promotion of Th2 responses (63). Most studies demonstrate
only the positive correlation of Arg-I with M2 macrophage phenotype, the causal relationship of Arg-I with macrophage phenotype
is, however, not fully clear. Importantly, a recent study demonstrates that Arg-I deficient mouse macrophages has even higher
polyamine production and does not impair gene expression in
response to IL-4 (66), which raises the question about the role
of Arg-I in M2 macrophage regulation. Future research shall
elucidate the causal role of Arg-I in regulation of macrophage
functional polarization.
DOES L-ARGININE DEFICIENCY EXPLAIN THE FUNCTIONS OF ARGINASE
ISOENZYMES IN MACROPHAGES?

There is continuing debate about the role of l-arginine deficiency in arginase-induced alterations of cellular functions. It
is generally believed that arginase including Arg-I and Arg-II

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Arginase and macrophage functions

causes l-arginine deficiency, resulting in decreased NO production from eNOS in endothelial cells (endothelial dysfunction) and
from iNOS in macrophages (M2 type function) (41). It has been
demonstrated that the concentration of l-arginine in adult human
and mouse plasma (0.1 mmol/L), as well as intracellular arginine
concentration (0.05–0.2 mmol/L) far exceed the Km of eNOS (2–
20 µmol/L) (67). Even though, acute l-arginine supplementation
in cells, isolated blood vessels, or in animals or in patients is able
to enhance NO production and improve endothelium-dependent
relaxations, a situation called “arginine paradox” (68, 69). This
phenomenon led to doubt whether l-arginine deficiency caused by
arginase is true. Several hypotheses have been proposed to explain
the “arginine paradox.” First, a “relative” intracellular l-arginine
deficiency, resulting from an increased level of endogenous competitors for eNOS substrate l-arginine such as ADMA that binds
to eNOS but could not be metabolized by the enzyme on top of
increased arginase activity either Arg-I or Arg-II in endothelial
cells has been suggested (70). Experiments showed that inhibition
of arginase improves eNOS function and overexpression of Arg-I
or Arg-II causes eNOS-uncoupling, leading to oxidative stress, and
decreased bioavailability of endothelial NO production, which is
associated with only 11–25% reduction in intracellular l-arginine
concentration in the presence of high-extracellular concentration
of l-arginine (0.4 mmol/L) (71). These results seem to support the
“relative l-arginine deficiency” hypothesis. It is worthy of noting
that NO production, particularly, iNOS/NO can be inhibited by
TGF-β, which is a strong Arg-I up-regulator and present in very
high amount during wound healing (72). It is presumable that NO
production is inhibited even under the condition of high-plasma
l-arginine concentration because of high concentration of TGF-β.
Whether this could explain the “arginine paradox” is not known.
Another alternative hypothesis is that a specific intracellular pool
of l-arginine for NO production may exist in endothelial cells
and could be depleted by enhanced arginase (73), yet, it is highly
speculative. It is not known whether enhanced arginase activity,
particularly Arg-II, could also cause iNOS-uncoupling, affecting
macrophage functions. Another puzzling is that why Arg-I and
Arg-II share the same l-arginine metabolizing function but seem
to exert distinct effects on macrophages.
If there is no real l-arginine deficiency caused by arginase either
Arg-I or Arg-II, l-arginine supplementation therapy aiming to
enhance endothelial NO production and to treat vascular disease
shall not work. In accordance, clinical studies in patients with acute
myocardial infarction or with peripheral arterial disease demonstrate that 6-month oral l-arginine supplementation (3 g three
times a day on top of standard medications) increase mortality
and shorten walking distance accompanied with decreased NO
production as compared to the placebo group (74). The underlying
mechanisms are not known and may be related to the induction of arginase, particularly Arg-II in vascular endothelial cells
by chronic l-arginine exposure as demonstrated by recent studies including our own (37, 69). These studies show that acute
supplementation of l-arginine to endothelial cells increases NO
bioavailability, while chronic l-arginine supplementation causes
eNOS-uncoupling mediated by up-regulation of Arg-II levels,
leading to endothelial senescence (69). Similar to this finding,
l-arginine has also been reported to cause iNOS-uncoupling in

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macrophages (75). These studies do not support a role of absolute
l-arginine deficiency caused by arginase, but strongly implicate
that too much l-arginine is even harmful, which is probably due
to production of other undesired metabolites from l-arginine as
speculated (76). Alternatively, a pleiotropic effect may also provide
explanation for the detrimental effects of arginase at least for ArgII under the condition of sufficient l-arginine supply. This point
will be discussed below.
DOES ARGINASE EXERT PLEIOTROPIC EFFECTS: L-ARGININE–UREA
HYDROLASE ACTIVITY-INDEPENDENT EFFECTS?

Any proteins or enzymes may have pleiotropic or off-target effects
that are not necessarily related to their canonical functions. We
have recently discovered that Arg-II exhibits its biological functions in vascular cells through both mechanisms, which are either
dependent or independent on l-arginine metabolizing function
(l-arginine–urea hydrolase activity) (77). We show that the catalytically inactive mouse Arg-II mutant with a point mutation of
histidine to phenylalanine at position 160 (referred to as H160F),
which lost its l-arginine–urea hydrolase activity, although does

Arginase and macrophage functions

not cause eNOS-dysfunction in endothelial cells, promotes cell
apoptosis and senescence to the same extent as the WT Arg-II
in vascular smooth muscle cells (VSMC). In contrast, only the
WT Arg-II (not the H160F inactive mutant) exerts function to
promote VSMC proliferation (Figure 3), which can be attributed
to the production of polyamine from l-arginine/ornithine pathway. This intriguing result provides evidence that Arg-II on one
hand promotes VSMC proliferation and on the other hand causes
VSMC apoptosis and senescence. While the cell proliferation–
stimulating effect of Arg-II is dependent on its l-arginine–urea
hydrolase activity via synthesis of ornithine and polyamines (71,
78), the cell apoptosis/senescence-promoting effect is independent
of its enzymatic activity (Figure 3). Further experiments show that
this l-arginine–urea hydrolase activity-independent effect is mediated through signaling pathways including mTORC1/S6K1, JNK,
and ERK1/2, converging on p66Shc leading to H2 O2 production
and mitochondrial dysfunction leading to cellular apoptosis and
senescence (77) (Figure 3). In parallel to these signaling pathways,
p53 is also activated by Arg-II independently of its l-arginine–
urea hydrolase activity, contributing to the cell senescence of the

FIGURE 3 | Canonical and pleiotropic effects of Arg-II in vascular endothelial and smooth muscle cells and underlying signaling mechanisms in
development of vascular aging and age-associated vascular diseases.

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apoptosis process. Importantly, expression of Arg-II and activities
of S6K1, ERK1/2, p66Shc, and p53 are all augmented in senescent VSMC, and genetic inhibition or ablation of Arg-II not only
reduces these signaling pathways and VSMC senescence/apoptosis
in vitro but also in atherosclerosis-prone ApoE−/− mice in vivo,
which at least in part accounts for the reduced plaque lesion formation and a more stable plaque characteristics in aortic roots in
Arg-II-deficient ApoE−/− mice (43, 77) (Figure 3). Moreover, we
also show that Arg-II negatively regulates autophagy function –
a cell protective mechanism of lysosomal proteolysis aiming to
remove harmful proteins (79) – in endothelial cells, which is also
independent on its l-arginine–urea hydrolase activity [Ref. (69),
Figure 3]. Decreased autophagy function in vascular cells and
macrophages are implicated in vascular aging and atherosclerotic
vascular disease. Indeed, recent studies provide evidence suggesting that adequate induction of autophagy protects against cellular
injury in endothelial and smooth muscle cells and formation of
foam cells, resulting in anti-atherosclerotic effects (80–82). In line
with this finding, genetic ablation of Arg-II in atherosclerotic
ApoE−/− mice preserves endothelial autophagy in aortas, which
associates with reduced atherosclerosis lesion formation (69). In
this study, we also demonstrate that Arg-II impairs endothelial
autophagy independently of the l-arginine–urea hydrolase activity through activation of mTORC1/S6K1 and p53, resulting in
inhibition of AMPK in endothelial cells, which contributes to
development of atherosclerosis (Figure 3). How Arg-II, independently of its l-arginine–urea hydrolase activity, impacts vascular
cell functions, remains to be investigated. Further, whether these
enzymatic dependent and independent effects also exist for ArgI and whether the pleiotropic effects of arginase account for
functional regulations in macrophages are unknown.
WHAT ARE THE MECHANISMS THAT REGULATE Arg-I AND Arg-II IN
MACROPHAGES

Arginase-I gene expression is inducible in macrophages by a variety of stimuli, for example, by elevated cAMP, IL-4, and TGF-β
(50). The regulation of Arg-I gene expression is mainly investigated at the transcriptional levels in murine macrophages, while
it remains to be investigated whether the findings are also true
in human cells. A number of transcription factors and nuclear
receptors such as RXR, PPARγ, PPAR δ, STAT6, C/EBPβ, KLF4,
PU.1, IRF8, and AP-1 have been shown to bind directly to specific sites in the promotor region of Arg-1 gene and in turn to
enhance Arg-1 expression. The complexity of Arg-I gene regulation mechanisms are further complicated by the fact that these
transcription factors can be regulated by post-translational modification mechanisms such as SUMOylation and ubiquitination
that are participating in the regulation of Arg-1 gene [for detailed
description of these mechanisms please refer to the review articles
(83, 84)]. There is, however, little information available regarding
the upstream regulatory mechanisms involved in gene expression
and enzymatic activity of Arg-I in macrophages. Also, very limited
information is provided on regulation of Arg-II in macrophages.
The stress sensor p38mapk has been demonstrated to participate in up-regulation of activity and expression of Arg-I and
Arg-II in macrophages (85, 86). This seems to be also the case
in bovine and rat aortic endothelial cells for Arg-I expression

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Arginase and macrophage functions

(87) and in human endothelial cells and mouse penile tissues
for Arg-II (88, 89). In accordance, in vivo treatment of hypertensive mouse induced by angiotensin-II infusion with a p38mapk
inhibitor prevents elevation of Arg-II expression and activity and
enhances endothelium-dependent relaxation (88). These studies
demonstrate that p38mapk is the upstream regulator of ArgII in endothelial cells and macrophages. Our most recent study
provides evidence showing that p38mapk also functions as downstream effector of Arg-II in endothelial cells, causing oxidative
stress through eNOS-uncoupling, since overexpression of Arg-II in
human endothelial cells causes eNOS-uncoupling and augments
p38mapk activation (90), and inhibition of p38mapk either pharmacologically by SB203580 or genetically by silencing the major
isoform p38mapkα in endothelial cells prevents eNOS-uncoupling
effect by Arg-II gene overexpression (90). Furthermore, mice fed
HFD, an obesity mouse model, exhibit enhanced Arg-II expression/activity and p38mapk activity and eNOS-uncoupling in the
aortas and inhibition of p38mapk recouples eNOS activity in the
obese mice. Moreover, mice deficient in Arg-II (Arg-II−/− ) on
the same obesogenic diet reveal decreased p38mapk activity and
eNOS function is fully preserved. These results demonstrate that
Arg-II causes eNOS-uncoupling through activation of p38mapk
in HFD-induced obesity (90). Together with the experiments discussed above, there might be a positive regulatory circuit between
p38mapk and Arg-II at least in vascular endothelial cells. Whether
this mechanism is also involved in Arg-I and/or Arg-II gene expression in macrophages is not known. A similar positive regulatory
circuit between S6K1 and Arg-II has also been demonstrated by
our recent studies in vascular endothelial cells (39).
In this study, we show that a persistent hyperactive S6K1 activity
is found to play a causal role in eNOS-uncoupling, leading to vascular endothelial aging and senescence (39, 91). Overexpression
of a constitutively active S6K1 mutant up-regulates Arg-II (not
Arg-I) gene expression and arginase activity in non-senescent cells
by stabilizing Arg-II mRNA (39). Conversely, silencing S6K1 in
senescent cells reduces Arg-II gene expression and activity and
genetic or pharmacological inhibition of S6K1 in senescent cells
or in old rat aortas decreases Arg-II gene expression and activity,
demonstrating a critical role of hyperactive S6K1 in up-regulating
Arg-II gene expression, resulting in enhanced arginase activity in
endothelial aging. Interestingly, silencing Arg-II gene in senescent endothelial cells or deficiency in Arg-II gene in mice reduces
S6K1 activity, recouples eNOS function in aging, and inhibits
endothelial expression of adhesion molecules such as ICAM-1
and VCAM-1, resulting in inhibition of monocyte-endothelial
cell interaction, demonstrating a positive vicious cycle between
S6K1 and Arg-II in vascular endothelial aging. These studies provide evidence showing that a mutual positive regulation between
S6K1 and Arg-II gene expression accelerates endothelial aging
through eNOS-uncoupling, leading to oxidative stress and inflammation (39). Further studies will analyze whether S6K1 is also
involved in regulation of Arg-I and/or Arg-II in macrophages,
participating in macrophages phenotype determination. Also, the
relationship between p38mapk, S6K1, and arginase remain to be
analyzed.
Other signaling pathways such as GTPase RhoA and its downstream kinase ROCK have been reported to mediate Arg-I gene

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expression in porcine coronary arterioles in response to hydrogen peroxide (H2 O2 ) and peroxynitrite (92) and Arg-II (but not
Arg-I) expression and/or activity in women with preeclampsia
(93) and in human endothelial cells in response to thrombin
(35), oxidized LDL (94), and hyperglycemia (36). In macrophages,
however, ROCK kinase inhibitor enhances Arg-I expression and
shift M1 to M2 phenotype (95), suggesting that ROCK pathway
may inhibit Arg-I expression. No information is available so far
whether Rho/ROCK pathway is involved in Arg-II regulation in
macrophages. For the detailed regulatory signaling mechanisms
of Arg-I and Arg-II expression/activity in vascular cells, please
refer to the review article (41).

CONCLUSION
The two isoforms of arginase, i.e., Arg-I and Arg-II, although
located in different subcellular compartments, share the same
function on l-arginine metabolism. Both isoenzymes hydrolyze
l-arginine to urea and l-ornithine, resulting in eNOS-uncoupling
in endothelial cells. In macrophages, Arg-I and Arg-II can be differentially induced by external stimuli. Evidence has been provided
that Arg-II plays a causal role in M1 functions, whereas Arg-I
is associated with M2 function in macrophages and widely used
as M2 marker for macrophages. However, the causal relationship
between Arg-I and M2 phenotype warrants further investigation. It remains to be characterized how Arg-I and Arg-II share
the same l-arginine metabolizing effect, but exhibit distinct or
opposite effects in macrophage inflammatory responses. Arg-II
as therapeutic target in chronic inflammatory disorders such as
age-associated vascular dysfunctions, atherosclerosis, and type-II
diabetes and complications has shown promising beneficial effects
in genetic modified mouse models (39, 43, 96). Some studies
implicate that targeting Arg-I is also beneficial for cardiovascular
functions, these studies are solely dependent on the pharmacological inhibitors, which inhibit both isoforms of arginases (97–99),
since systemic Arg-I deficient mouse exhibits severe symptoms
of hyperammonemia, and die between postnatal days 10 and 14
(31), one should consider that these inhibitors could inhibit liver
Arg-I, resulting in hyperammonemia. Taking into account that
Arg-I in macrophages may exhibit opposite effects as Arg-II, this
is another important reason to develop specific Arg-II inhibitors.
Moreover, whether Arg-I and Arg-II exert pleiotropic effects on
macrophage functions as demonstrated in vascular cells shall be
investigated. If this proves to be true, development of therapeutic drugs that target l-arginine–urea hydrolase activity may have
limitation on treatment of inflammatory diseases. Additionally,
signaling pathways that are involved in regulation of gene expression and enzymatic activity of both Arg-I and Arg-II shall be
further elucidated in macrophages. Characterization of these signaling mechanisms will also provide possibilities or rationales to
target arginase isoforms specifically in an indirect way to treat
inflammatory diseases. Finally, we have focused on cardiovascular
and metabolic diseases here. But, functional analysis of arginase
isoenzymes and their roles in macrophage polarization should
also help understanding other diseases, notably cancer. In particular, monocytes and macrophages are recruited into tumors and
regulate tumor growth by changing their functional phenotypes,
which is originally demonstrated by Mills and colleagues (100).

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Arginase and macrophage functions

M1 macrophage has been shown to have antitumor immunity,
whereas the M2 macrophage exerts protumorigenic properties
(101). Regardless of the inflammatory circumstance, it appears
that macrophage arginases are key players in influencing disease
outcomes.

ACKNOWLEDGMENTS
The original studies from own research were supported by
the Swiss National Science Foundation (310000-120435/1 and
310030-141070/1), Swiss Heart Foundation, and National Centre
of Competence in Research Program (NCCR-Kidney.ch).

REFERENCES
1. Hoeffel G, Wang Y, Greter M, See P, Teo P, Malleret B, et al. Adult Langerhans
cells derive predominantly from embryonic fetal liver monocytes with a minor
contribution of yolk sac-derived macrophages. J Exp Med (2012) 209:1167–81.
doi:10.1084/jem.20120340
2. Schulz C, Gomez Perdiguero E, Chorro L, Szabo-Rogers H, Cagnard N, Kierdorf K, et al. A lineage of myeloid cells independent of Myb and hematopoietic
stem cells. Science (2012) 336:86–90. doi:10.1126/science.1219179
3. Ginhoux F, Jung S. Monocytes and macrophages: developmental pathways and
tissue homeostasis. Nat Rev Immunol (2014) 14:392–404. doi:10.1038/nri3671
4. Jenkins SJ, Ruckerl D, Cook PC, Jones LH, Finkelman FD, van Rooijen
N, et al. Local macrophage proliferation, rather than recruitment from the
blood, is a signature of TH2 inflammation. Science (2011) 332:1284–8.
doi:10.1126/science.1204351
5. Robbins CS, Hilgendorf I, Weber GF, Theurl I, Iwamoto Y, Figueiredo JL, et al.
Local proliferation dominates lesional macrophage accumulation in atherosclerosis. Nat Med (2013) 19:1166–72. doi:10.1038/nm.3258
6. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8:958–69. doi:10.1038/nri2448
7. Wynn TA, Chawla A, Pollard JW. Macrophage biology in development, homeostasis and disease. Nature (2013) 496:445–55. doi:10.1038/nature12034
8. Galkina E, Ley K. Immune and inflammatory mechanisms of atherosclerosis (*). Annu Rev Immunol (2009) 27:165–97. doi:10.1146/annurev.immunol.
021908.132620
9. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity. J Innate Immun (2014) 6:716–26. doi:10.1159/000364945
10. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122:787–95. doi:10.1172/JCI59643
11. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32:463–88. doi:10.1615/CritRevImmunol.v32.i6.10
12. Martinez FO, Gordon S. The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000Prime Rep (2014) 6:13. doi:10.12703/P6-13
13. Oswald IP, Afroun S, Bray D, Petit JF, Lemaire G. Low response of BALB/c
macrophages to priming and activating signals. J Leukoc Biol (1992) 52:315–22.
14. Dileepan KN, Page JC, Li Y, Stechschulte DJ. Direct activation of murine peritoneal macrophages for nitric oxide production and tumor cell killing by
interferon-gamma. J Interferon Cytokine Res (1995) 15:387–94. doi:10.1089/
jir.1995.15.387
15. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.4049/
jimmunol.164.12.6166
16. Hibbs JB Jr, Vavrin Z, Taintor RR. L-arginine is required for expression of the
activated macrophage effector mechanism causing selective metabolic inhibition in target cells. J Immunol (1987) 138:550–65.
17. Nathan C, Ding A. Nonresolving inflammation. Cell (2010) 140:871–82.
doi:10.1016/j.cell.2010.02.029
18. Morris SM Jr. Arginine metabolism: boundaries of our knowledge. J Nutr
(2007) 137(6 Suppl 2):1602S–9S.
19. Dizikes GJ, Grody WW, Kern RM, Cederbaum SD. Isolation of human liver
arginase cDNA and demonstration of nonhomology between the two human
arginase genes. Biochem Biophys Res Commun (1986) 141:53–9. doi:10.1016/
S0006-291X(86)80333-3
20. Dizikes GJ, Spector EB, Cederbaum SD. Cloning of rat liver arginase cDNA
and elucidation of regulation of arginase gene expression in H4 rat hepatoma
cells. Somat Cell Mol Genet (1986) 12:375–84. doi:10.1007/BF01570732

October 2014 | Volume 5 | Article 533 | 184

Yang and Ming

21. Sparkes RS, Dizikes GJ, Klisak I, Grody WW, Mohandas T, Heinzmann C, et al.
The gene for human liver arginase (ARG1) is assigned to chromosome band
6q23. Am J Hum Genet (1986) 39:186–93.
22. Gotoh T, Sonoki T, Nagasaki A, Terada K, Takiguchi M, Mori M. Molecular cloning of cDNA for nonhepatic mitochondrial arginase (arginase II)
and comparison of its induction with nitric oxide synthase in a murine
macrophage-like cell line. FEBS Lett (1996) 395:119–22. doi:10.1016/00145793(96)01015-0
23. Vockley JG, Jenkinson CP, Shukla H, Kern RM, Grody WW, Cederbaum SD.
Cloning and characterization of the human type II arginase gene. Genomics
(1996) 38:118–23. doi:10.1006/geno.1996.0606
24. Gotoh T, Araki M, Mori M. Chromosomal localization of the human arginase
II gene and tissue distribution of its mRNA. Biochem Biophys Res Commun
(1997) 233:487–91. doi:10.1006/bbrc.1997.6473
25. Jenkinson CP, Grody WW, Cederbaum SD. Comparative properties of
arginases. Comp Biochem Physiol B Biochem Mol Biol (1996) 114:107–32.
doi:10.1016/0305-0491(95)02138-8
26. Morris SM Jr, Bhamidipati D, Kepka-Lenhart D. Human type II arginase:
sequence analysis and tissue-specific expression. Gene (1997) 193:157–61.
doi:10.1016/S0378-1119(97)00099-1
27. Santhanam L, Lim HK, Lim HK, Miriel V, Brown T, Patel M, et al. Inducible
NO synthase dependent S-nitrosylation and activation of arginase1 contribute
to age-related endothelial dysfunction. Circ Res (2007) 101:692–702.
28. Crombez EA, Cederbaum SD. Hyperargininemia due to liver
arginase deficiency. Mol Genet Metab (2005) 84:243–51. doi:10.1016/j.
ymgme.2004.11.004
29. Tsang JP, Poon WL, Luk HM, Fung CW, Ching CK, Mak CM, et al. Arginase
deficiency with new phenotype and a novel mutation: contemporary summary.
Pediatr Neurol (2012) 47:263–9. doi:10.1016/j.pediatrneurol.2012.06.012
30. Haraguchi Y, Takiguchi M, Amaya Y, Kawamoto S, Matsuda I, Mori M. Molecular cloning and nucleotide sequence of cDNA for human liver arginase. Proc
Natl Acad Sci U S A (1987) 84:412–5. doi:10.1073/pnas.84.2.412
31. Iyer RK, Yoo PK, Kern RM, Rozengurt N, Tsoa R, O’Brien WE, et al. Mouse
model for human arginase deficiency. Mol Cell Biol (2002) 22:4491–8. doi:10.
1128/MCB.22.13.4491-4498.2002
32. Choi S, Park C, Ahn M, Lee JH, Shin T. Immunohistochemical study of
arginase 1 and 2 in various tissues of rats. Acta Histochem (2012) 114:487–94.
doi:10.1016/j.acthis.2011.09.002
33. Xia Y, Dawson VL, Dawson TM, Snyder SH, Zweier JL. Nitric oxide synthase generates superoxide and nitric oxide in arginine-depleted cells leading to peroxynitrite-mediated cellular injury. Proc Natl Acad Sci U S A (1996)
93:6770–4. doi:10.1073/pnas.93.13.6770
34. Kim JH, Bugaj LJ, Oh YJ, Bivalacqua TJ, Ryoo S, Soucy KG, et al. Arginase
inhibition restores NOS coupling and reverses endothelial dysfunction and
vascular stiffness in old rats. J Appl Physiol (2009) 107:1249–57. doi:10.1152/
japplphysiol.91393.2008
35. Ming XF, Barandier C, Viswambharan H, Kwak BR, Mach F, Mazzolai L, et al. Thrombin stimulates human endothelial arginase enzymatic activity via RhoA/ROCK pathway: implications for atherosclerotic
endothelial dysfunction. Circulation (2004) 110:3708–14. doi:10.1161/01.CIR.
0000142867.26182.32
36. Romero MJ, Platt DH, Tawfik HE, Labazi M, El-Remessy AB, Bartoli M, et al.
Diabetes-induced coronary vascular dysfunction involves increased arginase
activity. Circ Res (2008) 102:95–102. doi:10.1161/CIRCRESAHA.107.155028
37. Scalera F, Closs EI, Flick E, Martens-Lobenhoffer J, Boissel JP, Lendeckel U,
et al. Paradoxical effect of L-arginine: acceleration of endothelial cell senescence. Biochem Biophys Res Commun (2009) 386:650–5. doi:10.1016/j.bbrc.
2009.06.091
38. Shin WS, Berkowitz DE, Ryoo SW. Increased arginase II activity contributes to
endothelial dysfunction through endothelial nitric oxide synthase uncoupling
in aged mice. Exp Mol Med (2012) 44:594–602. doi:10.3858/emm.2012.44.10.
068
39. Yepuri G, Velagapudi S, Xiong YY, Rajapakse AG, Montani JP, Ming XF, et al.
Positive crosstalk between arginase-II and S6K1 in vascular endothelial inflammation and aging. Aging Cell (2012) 11:1005–16. doi:10.1111/acel.12001
40. Lee SK, Kim JH, Yang WS, Kim SB, Park SK, Park JS. Exogenous nitric
oxide inhibits VCAM-1 expression in human peritoneal mesothelial cells.
Role of cyclic GMP and NF-kappaB. Nephron (2002) 90:447–54. doi:10.1159/
000054733

www.frontiersin.org

Arginase and macrophage functions

41. Yang Z, Ming XF. Arginase: the emerging therapeutic target for vascular oxidative stress and inflammation. Front Immunol (2013) 4:149. doi:10.3389/fimmu.
2013.00149
42. Khallou-Laschet J, Varthaman A, Fornasa G, Compain C, Gaston AT, Clement
M, et al. Macrophage plasticity in experimental atherosclerosis. PLoS One
(2010) 5:e8852. doi:10.1371/journal.pone.0008852
43. Ming XF, Rajapakse AG, Yepuri G, Xiong YY, Carvas JM, Ruffieux J, et al.
Arginase II promotes macrophage inflammatory responses through mitochondrial reactive oxygen species, contributing to insulin resistance and atherogenesis. J Am Heart Assoc (2012) 1:e000992. doi:10.1161/JAHA.112.000992
44. Munder M, Eichmann K, Modolell M. Alternative metabolic states in murine
macrophages reflected by the nitric oxide synthase/arginase balance: competitive regulation by CD4+ T cells correlates with Th1/Th2 phenotype. J Immunol
(1998) 160:5347–54.
45. Munder M, Eichmann K, Moran JM, Centeno F, Soler G, Modolell M. Th1/Th2regulated expression of arginase isoforms in murine macrophages and dendritic cells. J Immunol (1999) 163:3771–7.
46. Hesse M, Modolell M, La Flamme AC, Schito M, Fuentes JM, Cheever
AW, et al. Differential regulation of nitric oxide synthase-2 and arginase1 by type 1/type 2 cytokines in vivo: granulomatous pathology is shaped
by the pattern of L-arginine metabolism. J Immunol (2001) 167:6533–44.
doi:10.4049/jimmunol.167.11.6533
47. Rutschman R, Lang R, Hesse M, Ihle JN, Wynn TA, Murray PJ. Cutting
edge: Stat6-dependent substrate depletion regulates nitric oxide production.
J Immunol (2001) 166:2173–7. doi:10.4049/jimmunol.166.4.2173
48. El Kasmi KC, Qualls JE, Pesce JT, Smith AM, Thompson RW, Henao-Tamayo
M, et al. Toll-like receptor-induced arginase 1 in macrophages thwarts effective
immunity against intracellular pathogens. Nat Immunol (2008) 9:1399–406.
doi:10.1038/ni.1671
49. Marathe C, Bradley MN, Hong C, Lopez F, de Galarreta CM, Tontonoz P,
et al. The arginase II gene is an anti-inflammatory target of liver X receptor in macrophages. J Biol Chem (2006) 281:32197–206. doi:10.1074/jbc.
M605237200
50. Morris SM Jr. Regulation of arginine availability and its impact on NO synthesis. In: Ignarro LJ, editor. Nitric Oxide Biology and Pathobiology. San Diego,
CA: Academic Press (2000). p. 187–97.
51. Ross R. Atherosclerosis – an inflammatory disease. N Engl J Med (1999)
340:115–26. doi:10.1056/NEJM199901143400207
52. Swirski FK, Libby P, Aikawa E, Alcaide P, Luscinskas FW, Weissleder R, et al.
Ly-6Chi monocytes dominate hypercholesterolemia-associated monocytosis
and give rise to macrophages in atheromata. J Clin Invest (2007) 117:195–205.
doi:10.1172/JCI29950
53. Patsouris D, Li PP, Thapar D, Chapman J, Olefsky JM, Neels JG. Ablation of
CD11c-positive cells normalizes insulin sensitivity in obese insulin resistant
animals. Cell Metab (2008) 8:301–9. doi:10.1016/j.cmet.2008.08.015
54. Olefsky JM, Glass CK. Macrophages, inflammation, and insulin resistance.
Annu Rev Physiol (2010) 72:219–46. doi:10.1146/annurev-physiol-021909135846
55. Wentworth JM, Naselli G, Brown WA, Doyle L, Phipson B, Smyth GK, et al.
Pro-inflammatory CD11c+CD206+ adipose tissue macrophages are associated with insulin resistance in human obesity. Diabetes (2010) 59:1648–56.
doi:10.2337/db09-0287
56. Fligger J, Blum J, Jungi TW. Induction of intracellular arginase activity does
not diminish the capacity of macrophages to produce nitric oxide in vitro.
Immunobiology (1999) 200:169–86. doi:10.1016/S0171-2985(99)80068-0
57. Kepka-Lenhart D, Mistry SK, Wu G, Morris SM Jr. Arginase I: a limiting factor for nitric oxide and polyamine synthesis by activated macrophages? Am
J Physiol Regul Integr Comp Physiol (2000) 279:R2237–42.
58. Kropf P, Fuentes JM, Fahnrich E, Arpa L, Herath S, Weber V, et al. Arginase and
polyamine synthesis are key factors in the regulation of experimental leishmaniasis in vivo. FASEB J (2005) 19:1000–2.
59. Muller I, Hailu A, Choi BS, Abebe T, Fuentes JM, Munder M, et al. Age-related
alteration of arginase activity impacts on severity of leishmaniasis. PLoS Negl
Trop Dis (2008) 2:e235. doi:10.1371/journal.pntd.0000235
60. Skrzypek-Osiecka I, Robin Y, Porembska Z. Purification of rat kidney arginases
A1 and A4 and their subcellular distribution. Acta Biochim Pol (1983) 30:83–92.
61. Pauleau AL, Rutschman R, Lang R, Pernis A, Watowich SS, Murray PJ.
Enhancer-mediated control of macrophage-specific arginase I expression.
J Immunol (2004) 172:7565–73. doi:10.4049/jimmunol.172.12.7565

October 2014 | Volume 5 | Article 533 | 185

Yang and Ming

62. Gray MJ, Poljakovic M, Kepka-Lenhart D, Morris SM Jr. Induction of
arginase I transcription by IL-4 requires a composite DNA response element
for STAT6 and C/EBPbeta. Gene (2005) 353:98–106. doi:10.1016/j.gene.2005.
04.004
63. Martinez FO, Helming L, Gordon S. Alternative activation of macrophages: an
immunologic functional perspective. Annu Rev Immunol (2009) 27:451–83.
doi:10.1146/annurev.immunol.021908.132532
64. Wang XP, Chen YG, Qin WD, Zhang W, Wei SJ, Wang J, et al. Arginase I attenuates inflammatory cytokine secretion induced by lipopolysaccharide in vascular smooth muscle cells. Arterioscler Thromb Vasc Biol (2011) 31:1853–60.
doi:10.1161/ATVBAHA.111.229302
65. Pesce JT, Ramalingam TR, Mentink-Kane MM, Wilson MS, El Kasmi KC,
Smith AM, et al. Arginase-1-expressing macrophages suppress Th2 cytokinedriven inflammation and fibrosis. PLoS Pathog (2009) 5:e1000371. doi:10.
1371/journal.ppat.1000371
66. Van den Bossche J, Lamers WH, Koehler ES, Geuns JM, Alhonen L, Uimari A,
et al. Pivotal advance: arginase-1-independent polyamine production stimulates the expression of IL-4-induced alternatively activated macrophage markers while inhibiting LPS-induced expression of inflammatory genes. J Leukoc
Biol (2012) 91:685–99. doi:10.1189/jlb.0911453
67. Morris SM Jr. Regulation of enzymes of the urea cycle and arginine metabolism.
Annu Rev Nutr (2002) 22:87–105. doi:10.1146/annurev.nutr.22.110801.140547
68. Kurz S, Harrison DG. Insulin and the arginine paradox. J Clin Invest (1997)
99:369–70. doi:10.1172/JCI119166
69. Xiong Y, Fru MF, Yu Y, Montani JP, Ming XF, Yang Z. Long term exposure to Larginine accelerates endothelial cell senescence through arginase-II and S6K1
signaling. Aging (Albany NY) (2014) 6:369–79.
70. Antoniades C, Shirodaria C, Leeson P, Antonopoulos A, Warrick N, Van-Assche
T, et al. Association of plasma asymmetrical dimethylarginine (ADMA) with
elevated vascular superoxide production and endothelial nitric oxide synthase
uncoupling: implications for endothelial function in human atherosclerosis.
Eur Heart J (2009) 30:1142–50. doi:10.1093/eurheartj/ehp061
71. Li H, Meininger CJ, Hawker JR Jr, Haynes TE, Kepka-Lenhart D, Mistry SK, et al.
Regulatory role of arginase I and II in nitric oxide, polyamine, and proline syntheses in endothelial cells. Am J Physiol Endocrinol Metab (2001) 280:E75–82.
72. Vodovotz Y. Control of nitric oxide production by transforming growth factorbeta1: mechanistic insights and potential relevance to human disease. Nitric
Oxide (1997) 1:3–17. doi:10.1006/niox.1996.0105
73. Santhanam L, Christianson DW, Nyhan D, Berkowitz DE. Arginase and vascular
aging. J Appl Physiol (2008) 105:1632–42. doi:10.1152/japplphysiol.90627.2008
74. Wilson AM, Harada R, Nair N, Balasubramanian N, Cooke JP. L-arginine supplementation in peripheral arterial disease: no benefit and possible harm. Circulation (2007) 116:188–95. doi:10.1161/CIRCULATIONAHA.106.683656
75. Pekarova M, Lojek A, Martiskova H, Vasicek O, Bino L, Klinke A, et al. New
role for L-arginine in regulation of inducible nitric-oxide-synthase-derived
superoxide anion production in raw 264.7 macrophages. ScientificWorldJournal (2011) 11:2443–57. doi:10.1100/2011/321979
76. Dioguardi FS. To give or not to give? Lessons from the arginine paradox. J Nutrigenet Nutrigenomics (2011) 4:90–8. doi:10.1159/000327777
77. Xiong Y, Yu Y, Montani JP, Yang Z, Ming XF. Arginase-II induces vascular
smooth muscle cell senescence and apoptosis through p66Shc and p53 independently of its L-arginine ureahydrolase activity: implications for atherosclerotic plaque vulnerability. J Am Heart Assoc (2013) 2:e000096. doi:10.1161/
JAHA.113.000096
78. Chen B, Calvert AE, Cui H, Nelin LD. Hypoxia promotes human pulmonary
artery smooth muscle cell proliferation through induction of arginase. Am
J Physiol Lung Cell Mol Physiol (2009) 297:L1151–9. doi:10.1152/ajplung.
00183.2009
79. Boya P, Reggiori F, Codogno P. Emerging regulation and functions of
autophagy. Nat Cell Biol (2013) 15:713–20. doi:10.1038/ncb2788
80. Schrijvers DM, De Meyer GR, Martinet W. Autophagy in atherosclerosis: a
potential drug target for plaque stabilization. Arterioscler Thromb Vasc Biol
(2011) 31:2787–91. doi:10.1161/ATVBAHA.111.224899
81. LaRocca TJ, Henson GD, Thorburn A, Sindler AL, Pierce GL, Seals DR. Translational evidence that impaired autophagy contributes to arterial ageing. J Physiol
(2012) 590:3305–16. doi:10.1113/jphysiol.2012.229690
82. Ouimet M. Autophagy in obesity and atherosclerosis: interrelationships
between cholesterol homeostasis, lipoprotein metabolism and autophagy in

Frontiers in Immunology | Molecular Innate Immunity

Arginase and macrophage functions

macrophages and other systems. Biochim Biophys Acta (2013) 1831:1124–33.
doi:10.1016/j.bbalip.2013.03.007
83. Gibbings S, Elkins ND, Fitzgerald H, Tiao J, Weyman ME, Shibao G, et al.
Xanthine oxidoreductase promotes the inflammatory state of mononuclear
phagocytes through effects on chemokine expression, peroxisome proliferatoractivated receptor-{gamma} sumoylation, and HIF-1{alpha}. J Biol Chem
(2011) 286:961–75. doi:10.1074/jbc.M110.150847
84. Pourcet B, Pineda-Torra I. Transcriptional regulation of macrophage arginase
1 expression and its role in atherosclerosis. Trends Cardiovasc Med (2013)
23:143–52. doi:10.1016/j.tcm.2012.10.003
85. Stempin CC, Tanos TB, Coso OA, Cerban FM. Arginase induction promotes
Trypanosoma cruzi intracellular replication in cruzipain-treated J774 cells
through the activation of multiple signaling pathways. Eur J Immunol (2004)
34:200–9. doi:10.1002/eji.200324313
86. Liscovsky MV, Ranocchia RP, Gorlino CV, Alignani DO, Moron G, Maletto
BA, et al. Interferon-gamma priming is involved in the activation of arginase
by oligodeoxinucleotides containing CpG motifs in murine macrophages.
Immunology (2009) 128:e159–69. doi:10.1111/j.1365-2567.2008.02938.x
87. Zhu W, Chandrasekharan UM, Bandyopadhyay S, Morris SM Jr, DiCorleto PE, Kashyap VS. Thrombin induces endothelial arginase through AP-1
activation. Am J Physiol Cell Physiol (2010) 298:C952–60. doi:10.1152/ajpcell.
00466.2009
88. Toque HA, Romero MJ, Tostes RC, Shatanawi A, Chandra S, Carneiro ZN, et al.
p38 Mitogen-activated protein kinase (MAPK) increases arginase activity and
contributes to endothelial dysfunction in corpora cavernosa from angiotensinII-treated mice. J Sex Med (2010) 7:3857–67. doi:10.1111/j.1743-6109.2010.
01996.x
89. Giri H, Muthuramu I, Dhar M, Rathnakumar K, Ram U, Dixit M. Protein tyrosine phosphatase SHP2 mediates chronic insulin-induced endothelial
inflammation. Arterioscler Thromb Vasc Biol (2012) 32:1943–50. doi:10.1161/
ATVBAHA.111.239251
90. Yu Y, Rajapakse AG, Montani JP, Yang Z, Ming XF. p38 Mitogen-activated protein kinase is involved in arginase-II-mediated eNOS-uncoupling in obesity.
Cardiovasc Diabetol (2014) 13:113. doi:10.1186/s12933-014-0113-z
91. Rajapakse AG, Yepuri G, Carvas JM, Stein S, Matter CM, Scerri I, et al. Hyperactive S6K1 mediates oxidative stress and endothelial dysfunction in aging: inhibition by resveratrol. PLoS One (2011) 6:e19237. doi:10.1371/journal.pone.
0019237
92. Thengchaisri N, Hein TW, Wang W, Xu X, Li Z, Fossum TW, et al. Upregulation of arginase by H2O2 impairs endothelium-dependent nitric oxidemediated dilation of coronary arterioles. Arterioscler Thromb Vasc Biol (2006)
26:2035–42. doi:10.1161/01.ATV.0000233334.24805.62
93. Sankaralingam S, Xu H, Davidge ST. Arginase contributes to endothelial cell
oxidative stress in response to plasma from women with preeclampsia. Cardiovasc Res (2010) 85:194–203. doi:10.1093/cvr/cvp277
94. Ryoo S, Bhunia A, Chang F, Shoukas A, Berkowitz DE, Romer LH. OxLDLdependent activation of arginase II is dependent on the LOX-1 receptor and
downstream RhoA signaling. Atherosclerosis (2011) 214:279–87. doi:10.1016/
j.atherosclerosis.2010.10.044
95. Liu C, Li Y, Yu J, Feng L, Hou S, Liu Y, et al. Targeting the shift from M1
to M2 macrophages in experimental autoimmune encephalomyelitis mice
treated with fasudil. PLoS One (2013) 8:e54841. doi:10.1371/journal.pone.
0054841
96. Vaisman BL, Andrews KL, Khong SM, Wood KC, Moore XL, Fu Y, et al. Selective
endothelial overexpression of arginase II induces endothelial dysfunction and
hypertension and enhances atherosclerosis in mice. PLoS One (2012) 7:e39487.
doi:10.1371/journal.pone.0039487
97. Holowatz LA, Kenney WL. Up-regulation of arginase activity contributes to
attenuated reflex cutaneous vasodilatation in hypertensive humans. J Physiol
(2007) 581:863–72. doi:10.1113/jphysiol.2007.128959
98. Shemyakin A, Kovamees O, Rafnsson A, Bohm F, Svenarud P, Settergren M, et al.
Arginase inhibition improves endothelial function in patients with coronary
artery disease and type 2 diabetes mellitus. Circulation (2012) 126:2943–50.
doi:10.1161/CIRCULATIONAHA.112.140335
99. Stanhewicz AE, Bruning RS, Smith CJ, Kenney WL, Holowatz LA. Local tetrahydrobiopterin administration augments reflex cutaneous vasodilation through
nitric oxide-dependent mechanisms in aged human skin. J Appl Physiol (2012)
112:791–7. doi:10.1152/japplphysiol.01257.2011

October 2014 | Volume 5 | Article 533 | 186

Yang and Ming

100. Mills CD, Shearer J, Evans R, Caldwell MD. Macrophage arginine metabolism
and the inhibition or stimulation of cancer. J Immunol (1992) 149:2709–14.
101. Noy R, Pollard JW. Tumor-associated macrophages: from mechanisms to therapy. Immunity (2014) 41:49–61. doi:10.1016/j.immuni.2014.06.010
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 02 September 2014; accepted: 09 October 2014; published online: 27 October
2014.

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Arginase and macrophage functions

Citation: Yang Z and Ming X-F (2014) Functions of arginase isoforms in macrophage
inflammatory responses: impact on cardiovascular diseases and metabolic disorders.
Front. Immunol. 5:533. doi: 10.3389/fimmu.2014.00533
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Yang and Ming . This is an open-access article distributed under the
terms of the Creative Commons Attribution License (CC BY). The use, distribution or
reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
accepted academic practice. No use, distribution or reproduction is permitted which
does not comply with these terms.

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REVIEW ARTICLE
published: 10 September 2014
doi: 10.3389/fimmu.2014.00435

Role of alveolar macrophages in chronic obstructive
pulmonary disease
Ross Vlahos*† and Steven Bozinovski *†
Department of Pharmacology and Therapeutics, Lung Health Research Centre, The University of Melbourne, Parkville, VIC, Australia

Edited by:
Klaus Ley, La Jolla Institute for Allergy
and Immunology, USA
Reviewed by:
Andrew Tasman Hutchinson,
University of Technology Sydney,
Australia
Janos G. Filep, University of
Montreal, Canada
*Correspondence:
Ross Vlahos and Steven Bozinovski ,
Department of Pharmacology and
Therapeutics, Lung Health Research
Centre, The University of Melbourne,
Parkville, VIC 3010, Australia
e-mail: rossv@unimelb.edu.au;
bozis@unimelb.edu.au
†

Ross Vlahos and Steven Bozinovski
have contributed equally to this work.

Alveolar macrophages (AMs) represent a unique leukocyte population that responds to airborne irritants and microbes. This distinct microenvironment coordinates the maturation of
long-lived AMs, which originate from fetal blood monocytes and self-renew through mechanisms dependent on GM-CSF and CSF-1 signaling. Peripheral blood monocytes can also
replenish lung macrophages; however, this appears to occur in a stimuli specific manner.
In addition to mounting an appropriate immune response during infection and injury, AMs
actively coordinate the resolution of inflammation through efferocytosis of apoptotic cells.
Any perturbation of this process can lead to deleterious responses. In chronic obstructive pulmonary disease (COPD), there is an accumulation of airway macrophages that
do not conform to the classic M1/M2 dichotomy. There is also a skewed transcriptome
profile that favors expression of wound-healing M2 markers, which is reflective of a deficiency to resolve inflammation. Endogenous mediators that can promote an imbalance in
inhibitory M1 vs. healing M2 macrophages are discussed, as they are the plausible mechanisms underlying why AMs fail to effectively resolve inflammation and restore normal lung
homeostasis in COPD.
Keywords: alveolar macrophage, chronic obstructive pulmonary disease, efferocytosis, lung inflammation,
oxidative stress, resolution

INTRODUCTION
Macrophages are essential for pulmonary host defense through
their capacity to survey the exposed airways and regulate innate
and adaptive immunity. The pulmonary macrophage system consists of several different populations that are found in anatomically distinct compartments, including the airways, alveolar spaces
[alveolar macrophages (AMs)], and resident lung tissue. AMs constitute over 90% of the pulmonary macrophage population (1)
and have traditionally thought to originate from the bone marrow
(2). More recently, it has been shown that although AMs originate from fetal blood monocytes within the first week of life
via GM-CSF dependent mechanisms, maintenance of this population during homeostasis is dependent on their self-renewing
capacity (3). Once, developed, this lung macrophage pool is longlived in humans (2) in the absence of an inflammatory insult.
This finding has been replicated in mice, where there was approximately 40% turnover of AMs over 1 year in the absence of an
insult (4). Hence, the macrophage lung population, particularly
during the steady state, is primarily sustained by the self-renewal
of pulmonary AMs through local proliferation (5). The local proliferation of AMs during homeostatic repletion is dependent on
both GM-CSF and CSF-1, with no dependence on IL-4 signaling
(6). Both GM-CSF and M-CSF also control the proliferation and
survival of AMs (7). The differentiation of AMs is particularly
dependent on GM-CSF, where the leukocyte growth factor regulates essential functions including phagocytosis and surfactant
catabolism through the PU.1 transcription factor (8). This is consistent with a unique airway environment that is associated with
high oxygen tension and high levels of GM-CSF.

Frontiers in Immunology | Molecular Innate Immunity

Resident AMs are constantly encountering inhaled substances
due to their exposed position in the alveolar lumen. AMs are considered to be major effector cells in innate host defense against
inhaled irritants by virtue of their phagocytic ability (9). Therefore,
it is vital that resident AMs are kept in a relatively quiescent state
with active suppression of inflammation in response to harmless
antigens to prevent collateral damage to lung tissue (10). Although
AM exhibit microbicidal, tumoricidal, and parasiticidal activities
(9, 11), they are functionally less responsive than tissue-resident
macrophages. Relative to tissue macrophages, they display reduced
phagocytic capacity, reduced respiratory burst, and a diminished
capacity to present antigen to T cells (12–14). Under homeostatic conditions, AMs are closely associated with alveolar epithelial
cells (AECs), and this in turn induces the expression of epithelial restricted αvβ6 integrin that binds and activates latent TGFβ
(15). TGFβ can inhibit macrophage activation that is implicated
in alveolar wall destruction (15). Upon recognition of antigens
by TLRs, the rapid induction of actin polymerization promotes
AMs detachment from AECs (16). The subsequent production of
proteases by activated AMs then activates latent TGFβ, thereby
reinstating AMs to their resting state (16). This illustrates an
intricate mechanism of microenvironmental macrophage specialization to keep the macrophage response in check. AMs can also
produce TGFβ, which suppresses T cell activation and has been
shown to promote the emergence of T regulatory cells (17).

ROLE OF AMs IN INNATE HOST DEFENSE
In contrast to the self-renewing capacity of AMs during the steady
state, inflammatory and infective insults dramatically change the

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dynamics of local lung macrophages. Using chimeric mice, parabiosis, and adoptive cellular transfer models, it has been established
that bone marrow-derived blood monocytes replenish the AM
pool during lethal irradiation (3). This is context specific, as inoculation with influenza in mice, which dramatically depletes resident
AMs in a strain specific manner, led to restoration of this population through self-renewal proliferative mechanisms (6). This infectious model also promotes the substantial recruitment of blood
monocytes; however, the fate of monocyte-derived macrophages
has yet to be established in this setting. Using a similar approach,
LPS has been shown to restore lung macrophage numbers through
both local proliferation of resident macrophages and maturation
of recruited blood monocytes (18). Mouse blood monocytes can
be subdivided according to differential expression of chemokine
receptors and adhesion molecules that are involved in cell recruitment. CCR2+ Gr1hi CX3 CR1lo is actively recruited to inflamed
tissues by virtue of their recognition of CCL2 (also known as MCP1) (19). Mouse monocytes that are CCR2+ Gr1hi CX3 CR1lo are
classified as pro-inflammatory but share morphological characteristic and chemokine receptor patterns with the classical human
monocytes (CD14hi CD16− CX3 CR1lo ) (19). Monocyte-derived
macrophages can acquire distinct morphological and functional
properties as directed by the immunological microenvironment.
Alveolar macrophages coordinate antimicrobial defenses
through expression of receptors for immunoglobulin (Fc R), complement, β-glucan, mannose, and several types of scavenger receptors that together facilitates phagocytosis (20). AMs generate reactive nitrogen and oxygen intermediates involved in macrophagemediated defense against microbial infection (21, 22). Surfactant protein A augments pathogen killing by AMs by stimulating phagocytosis and production of reactive oxygen–nitrogen
intermediates (23). In addition, AMs can initiate recruitment of
inflammatory cells from pulmonary vasculature into the alveolar space. There are a number of studies that implicate AMs
as central effector cells in the production of pro-inflammatory
cytokines, which initiate the early phase of neutrophil influx in
response to acute lung injury caused by bacterial products (24, 25).
More recently, selective targeting strategies that ablate different
monocyte/macrophage populations have identified an important
role for peripheral blood monocytes. Ablation of CCR2hi monocytes significantly reduced indices of acute lung injury (26). It
is plausible that AMs may actually play a role in limiting neutrophil influx by controlling MCP-1 production through AECs
(27). Furthermore, in a murine model of pneumococcal pneumonia, AMs depletion resulted in a failure to modulate the
inflammatory response with increased levels of pro-inflammatory
cytokines (28). AMs are also central regulators of the resolution of
inflammation through their ability to engulf apoptotic neutrophils
during the resolution phase (28, 29). The active phagocytosis
of dying cells by macrophages may also lead to the induction
of anti-inflammatory or suppressive properties in macrophages
as shown by the inhibition of IL-1β, IL-8, TNFα, and GM-CSF
production (30).
Alveolar macrophages are also indispensable for the clearance
of influenza infection in a viral strain-dependent manner in mice.
Here, it was established that the depletion of airway macrophages
was associated with more severe lung injury following inoculation

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Alveolar macrophages in COPD

with BJx109, which is a viral strain that infects macrophages with
high efficiency (31). This is in contrast to the highly virulent
PR8 strain that poorly replicated in airway macrophages, suggesting that avoidance of AM engagement may contribute to the
virulence of influenza strains (31). There is also the important
clinical complication of secondary bacterial infections following
a significant viral event, where AM function is likely to be compromised. In murine models of secondary bacterial infection, the
initial depletion of AMs as a consequence of influenza infection
rendered the host susceptible to Streptococcus pneumonia (Spn)
colonization and systemic invasion (32). The repletion of resident
AMs occurred 2 weeks after influenza, which resulted in the reestablishment of early innate host protection to Spn (32). This
AM replenishment phase may represent a window of opportunity for opportunistic respiratory pathogens such as Spn that take
advantage of this immunocompromised state. Interferon-γ production during the recovery phase of a viral infection can also
inhibit lung anti-bacterial defenses. Mechanistically, it was shown
that viral-induced production of inteferon-γ caused downregulation of the scavenger receptor MARCO, and neutralization of
interferon-γ prevented secondary pneumococcal infection (33).
Using MARCO-deficient mice, it has been established that expression of this scavenger receptor on AMs is critical for efficient
clearance of Spn from the lungs (34).

ROLE OF AMs IN CHRONIC OBSTRUCTIVE PULMONARY
DISEASE
There is a large body of evidence implicating AMs in the pathogenesis of Chronic Obstructive Pulmonary Disease (COPD). COPD is
a major global health problem and has been predicted to become
the third largest cause of death in the world by 2020 (35). Cigarette
smoking is the major cause of COPD and accounts for more than
95% of cases in industrialized countries (36), but other environmental pollutants are important causes in developing countries
(37). COPD is “a disease state characterized by airflow limitation
that is not fully reversible.” The airflow limitation is usually progressive and associated with an abnormal inflammatory response
of lungs to noxious particles and gases (38). COPD encompasses
chronic obstructive bronchiolitis with fibrosis and obstruction of
small airways and emphysema with enlargement of airspaces and
destruction of lung parenchyma, loss of lung elasticity, and closure
of small airways. Most patients with COPD have all three pathologic conditions (chronic obstructive bronchiolitis, emphysema,
and mucus plugging), but the relative extent of emphysema and
obstructive bronchiolitis within individual patients can vary.
Studies have highlighted that macrophages play a pivotal role
in the pathophysiology of COPD (39). There is a marked increase
(5- to 10-fold) in the numbers of macrophages in airways, lung
parenchyma, bronchoalveolar lavage fluid (BALF), and sputum
in patients with COPD (40, 41). A morphometric analysis of
macrophage numbers in the parenchyma of patients with emphysema showed a 25-fold increase in the numbers of macrophages
in the tissue and alveolar space compared to smokers with normal lung function (42). There is a positive correlation between
macrophage numbers in the airways and the severity of COPD
(43). In addition, a pathological role for macrophages has been
demonstrated, as the depletion of lung macrophages conferred

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protection against the development of emphysema in an experimental model of COPD (44). Macrophages are activated by
cigarette smoke and other irritants to release inflammatory mediators. AMs also secrete elastolytic enzymes (proteases), including
matrix metalloprotease (MMP)-2, MMP-9, MMP-12, cathepsin K,
L, and S in response to irritants and infection, which together are
responsible for destruction of lung parenchyma (36). In patients
with emphysema, there is an increase in BALF concentrations
and macrophage expression of MMP-1 and MMP-9 (45). There
is an increase in activity of MMP-9 in the lung parenchyma of
patients with emphysema (46). AMs from smokers with normal
lung function express more MMP-9 than those from non-smoking
healthy subjects (47), and there is an even greater increase in
cells from patients with COPD, which have enhanced elastolytic
activity (48).
Chronic obstructive pulmonary disease subjects can also be
very susceptible to bacterial colonization (49, 50) and exacerbations that are commonly caused by respiratory infections of viral
and/or bacterial etiology (51). The frequent exacerbator phenotype has been reported, which is associated with a poorer quality of
life and increased systemic inflammation (52). Impaired AM function is central to high colonization rates and increased susceptibility to exacerbations observed in COPD. Chronic cigarette smoke
exposure is a major cause of COPD, which markedly depletes intracellular GSH stores (53, 54). Oxidative stress leads to disruption of
GSH metabolism, which is considered as a key susceptibility feature of lung diseases (55). Excessive oxidative stress is particularly
deleterious to AM function, leading to a deficiency in phagocytosis of bacteria (56) and efferocytosis of apoptotic cells (57).
Treatment with anti-oxidants such as procysteine can significantly
improve efferocytic function of AMs isolated from experimental
models of COPD (58). Impaired AM-mediated efferocytosis in
COPD can be particularly damaging in COPD as neutrophils are
persistently recruited into the airways. Cigarette smoke impairs
clearance of apoptotic cells through oxidant-dependent activation
of RhoA (59) and inhibition of Rac1 (60), leading to defective actin
polymerization normally required for efficient efferocytosis. The
inability to efficiently remove exhausted neutrophils has damaging implications in COPD as accumulation of necrotic neutrophils
can lead to the indiscriminate release of granule protease pools
including neutrophil elastase. Neutrophil elastase localizes to lung
elastic fibers in emphysematic patients and degrades extracellular
matrix components (61) and can promote the release of mucins
through epidermal growth factor receptor (EGFR)-dependent
mechanisms (62). EGFR transactivation can also augment inflammatory responses initiated by rhinovirus infection (63). Reactive
free radicals also impair clearance mechanisms by directly causing
cytoskeletal instability and carbonyl modification of pseudopodia
(64–66). Macrophages also interact with carbonyl-adduct modified extracellular matrix proteins, which impair their ability to
clear apoptotic neutrophils (67).
In addition to oxidative post translational modification of the
host phagocytic machinery, the complex milieu within COPD airways can alter the phenotype of highly plastic airway macrophages.
It is becoming increasing clear that different macrophage subpopulations exist in the inflamed lung. Although the existence of such

Frontiers in Immunology | Molecular Innate Immunity

Alveolar macrophages in COPD

populations is implicated in COPD, the importance of these subpopulations is unknown (68). The ongoing characterization of
disease-associated macrophages clearly demonstrates that they do
not conform into the classic M1/M2 dichotomy and it is likely
that the inflammatory environment of the COPD airways drives
development of both M1 and M2 macrophages (58, 69). Indeed, it
has been shown that iNOS and arginase activity are concurrently
elevated in COPD airways. Specifically, elevated iNOS expression
has been observed in AMs of COPD patients, which increased with
severity of disease and during exacerbations (70–72). Of interest,
levels of exhaled nitric oxide (NO) are not elevated in stable COPD,
which may be consequential to increased production of superoxide that reacts with NO to generate the highly reactive nitrogen
species, peroxynitrite (73). In addition, arginase-1 is increased
in cigarette smoke exposure models (74), which will reduce larginine availability to iNOS. Reduced l-arginine bioavailability
also stimulates iNOS to simultaneously produce NO and superoxide, which facilitates rapid formation of peroxynitrite (73). Consistent with the concurrent existence of iNOS and arginase expressing
macrophages in COPD, nitrotyrosine (product of peroxynitrite)
levels have been shown to be increased in sputum macrophages
of COPD patients, which are negatively correlated with their
lung function (75). Hence, the relative ratio of iNOS expressing M1 macrophages and arginase expressing M2 macrophages
will be particularly important to the oxidative/nitrosative state of
the lung.
The relative balance between these polarization states can in
turn, have a profound impact on disease progression [reviewed
in Ref. (76)]. There is evidence for the reprograming of AMs as a
consequence of chronic smoke exposure that is associated with the
induction of a unique set of genes including MMP12 (77). There
is also evidence for the transcriptional skewing of AMs toward an
M2 gene profile in smokers with normal lung function that was
more evident in smokers who had progressed to develop COPD
(78). In this study, they also demonstrate the progressive downregulation of M1 genes (78), which would appear paradoxical
to the observation of increased expression of pro-inflammatory
mediators in COPD. Hence, there is a need to better define the relative contribution of M1 vs. M2 macrophages as there is emerging
evidence that both populations do concurrently exist in COPD
airways. Another important consideration in COPD is the interaction between macrophages and T cell subsets. Although it has
been broadly stated that T cells control the polarization state of
macrophages, macrophages can also potently regulate T cell biology (79, 80). Since T cell subsets including CD8+ T cells, TH 17, and
iBALT formation are implicated in COPD pathology, the role of
macrophages in regulating T cell biology remains to be elucidated.
The accumulation of M2 skewed airway macrophages may be
reflective of deficient resolution processes that normally switch
off inflammation and restore lung homeostasis. Since the stimulation of non-phlogistic phagocytosis is essential to resolution of
inflammation, the oxidant-dependent impairment of efferocytic
clearance of damaged tissue may maintain M2 macrophages in
COPD. The induction of CD163 is commonly recognized as a
marker for M2-alternatively activated macrophages involved in
wound-healing (81, 82) and CD163 positive macrophages are

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Vlahos and Bozinovski

highly prominent in the BAL compartment of current and exsmokers with COPD (83). CD163 may constitute a major defense
mechanism to protect the lung as it functions as a scavenger
receptor, which promotes degradation of HbHp complexes and
signaling that induces expression of heme-oxygenase-1 (HO-1)
(82). The persistence of HO-1 in COPD airways (84) is consistent with an environment where there is excessive oxidative stress
and a deficiency in the resolution of inflammation. The mediators
that induce expression of M2 markers in COPD have not been
comprehensively characterized. Interleukin-10 (IL-10) is a potent
inducer of CD163 expression in human monocytes (85); however, there is some data to suggest that the level of IL-10 positive
macrophages is reduced in COPD (86). An alternate mediator that
has been shown to potently induce expression of CD163 in human
monocyte-derived macrophages is serum amyloid A (SAA) (87).
SAA is a major acute phase reactant that has now been shown to
be expressed in COPD lungs, where its levels correlated with neutrophilic inflammation (88). SAA is known to target the ALX/FPR2
receptor and oppose the actions of pro-resolving ligands such
as LipoxinA4 (88–90), which normally stimulate non-phlogistic
clearance pathways. In addition to CD163, SAA can also stimulate
the expression of the TH 17 polarizing cytokines, IL-6, and IL-1β
in monocyte-derived macrophages, and neutralization of IL-17A
expression suppressed neutrophil airway inflammation stimulated
by SAA (91). Hence, the persistence of host defense mediators
such as SAA may maintain alternative macrophage populations in
COPD airways that not only express M2 markers of wound repair,
but also markers of acute inflammation.

Alveolar macrophages in COPD

FIGURE 1 | COPD alveolar macrophage. Cigarette smoke, oxidative
stress, and the airway inflammatory microenvironment have a direct effect
on alveolar macrophage (AM) phenotype in COPD that leads to the
emergence of M1 and M2 populations. The ratio of these macrophages will
govern the pathological processes in COPD. M1 macrophages will further
drive inflammation and oxidative stress. Excessive oxidative stress impairs
resolution mechanisms including macrophage-mediated phagocytosis and
efferocytosis, which leads to colonization and exacerbations in COPD. In
addition, the emergence of M2 macrophages can contribute to deleterious
lung remodeling/damage through increased expression of M2-related
genes and excessive protease (MMP-9, -12) production.

CONCLUSION
In this review, the origin and maintenance of AMs and their
essential role in innate immunity to respiratory pathogens are
discussed. There is emerging evidence for the self-renewal of AMs
and lung tissue macrophages through mechanisms dependent on
GM-CSF and CSF-1 signaling. The role of peripheral monocytes
in replenishing lung macrophages appear to be context specific,
as radiation-induced lung injury stimulates monocytic replenishment, whereas influenza infection replenishes AMs through local
proliferation of resident macrophages. In diseases such as COPD
where there is an accumulation of airway macrophages, the relative
contribution of monocyte versus local proliferation of mature AM
populations remains to be determined. Lineage tracing of monocytes/macrophages in experimental models of COPD will inform
on the origin of macrophages associated with disease pathology.
Alveolar macrophages coordinate the efficient clearance of
inhaled irritants and microbes to resolve inflammation. In addition, AMs display efferocytic activity to clear damaged tissue and
cells following injury and infection. In COPD, the persistence of
inflammation and the inability to efficiently clear damaged tissue
and exhausted immune cells such as neutrophils may be due to
excessive oxidative stress that impairs the phagocytic capacity of
AMs. Airway macrophages in COPD also display a unique phenotype that is associated with the induction of M2-related genes,
which are likely to be upregulated in response to local tissue damage (Figure 1). The maintenance of this subpopulation may also
contribute to deleterious remodeling in COPD. Thus, there is a

www.frontiersin.org

need to better characterize distinct AM populations present in
COPD and their relative contribution to disease pathology, as their
highly plastic nature offer a therapeutic opportunity to reprogram
macrophages to facilitate restoration of lung homeostasis.

REFERENCES
1. van oud Alblas AB, van Furth R. Origin, kinetics, and characteristics of
pulmonary macrophages in the normal steady state. J Exp Med (1979)
149(6):1504–18. doi:10.1084/jem.149.6.1504
2. Thomas ED, Ramberg RE, Sale GE, Sparkes RS, Golde DW. Direct evidence
for a bone marrow origin of the alveolar macrophage in man. Science (1976)
192(4243):1016–8. doi:10.1126/science.775638
3. Guilliams M, De Kleer I, Henri S, Post S, Vanhoutte L, De Prijck S, et al. Alveolar macrophages develop from fetal monocytes that differentiate into long-lived
cells in the first week of life via GM-CSF. J Exp Med (2013) 210(10):1977–92.
doi:10.1084/jem.20131199
4. Maus UA, Janzen S, Wall G, Srivastava M, Blackwell TS, Christman JW, et al.
Resident alveolar macrophages are replaced by recruited monocytes in response
to endotoxin-induced lung inflammation. Am J Respir Cell Mol Biol (2006)
35(2):227–35. doi:10.1165/rcmb.2005-0241OC
5. Tarling JD, Lin HS, Hsu S. Self-renewal of pulmonary alveolar macrophages:
evidence from radiation chimera studies. J Leukoc Biol (1987) 42(5):443–6.
6. Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, et al. Tissueresident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity (2013) 38(4):792–804.
doi:10.1016/j.immuni.2013.04.004
7. Lin HS, Lokeshwar BL, Hsu S. Both granulocyte-macrophage CSF and
macrophage CSF control the proliferation and survival of the same subset of
alveolar macrophages. J Immunol (1989) 142(2):515–9.

September 2014 | Volume 5 | Article 435 | 191

Vlahos and Bozinovski

8. Shibata Y, Berclaz PY, Chroneos ZC, Yoshida M, Whitsett JA, Trapnell BC.
GM-CSF regulates alveolar macrophage differentiation and innate immunity
in the lung through PU.1. Immunity (2001) 15(4):557–67. doi:10.1016/S10747613(01)00218-7
9. Franke-Ullmann G, Pfortner C, Walter P, Steinmuller C, Lohmann-Matthes ML,
Kobzik L. Characterization of murine lung interstitial macrophages in comparison with alveolar macrophages in vitro. J Immunol (1996) 157(7):3097–104.
10. Thepen T, Van Rooijen N, Kraal G. Alveolar macrophage elimination in vivo is
associated with an increase in pulmonary immune response in mice. J Exp Med
(1989) 170(2):499–509. doi:10.1084/jem.170.2.499
11. Prokhorova S, Lavnikova N, Laskin DL. Functional characterization of interstitial macrophages and subpopulations of alveolar macrophages from rat lung.
J Leukoc Biol (1994) 55(2):141–6.
12. Holt PG. Inhibitory activity of unstimulated alveolar macrophages on Tlymphocyte blastogenic response. Am Rev Respir Dis (1978) 118(4):791–3.
13. Hoidal JR, Schmeling D, Peterson PK. Phagocytosis, bacterial killing, and metabolism by purified human lung phagocytes. J Infect Dis (1981) 144(1):61–71.
doi:10.1093/infdis/144.1.61
14. Blumenthal RL, Campbell DE, Hwang P, DeKruyff RH, Frankel LR, Umetsu
DT. Human alveolar macrophages induce functional inactivation in antigenspecific CD4 T cells. J Allergy Clin Immunol (2001) 107(2):258–64. doi:10.1067/
mai.2001.112845
15. Morris DG, Huang X, Kaminski N, Wang Y, Shapiro SD, Dolganov G,
et al. Loss of integrin alpha(v)beta6-mediated TGF-beta activation causes
Mmp12-dependent emphysema. Nature (2003) 422(6928):169–73. doi:10.1038/
nature01413
16. Takabayshi K, Corr M, Hayashi T, Redecke V, Beck L, Guiney D, et al. Induction of a homeostatic circuit in lung tissue by microbial compounds. Immunity
(2006) 24(4):475–87. doi:10.1016/j.immuni.2006.02.008
17. Coleman MM, Ruane D, Moran B, Dunne PJ, Keane J, Mills KH. Alveolar
macrophages contribute to respiratory tolerance by inducing FoxP3 expression
in naive T cells. Am J Respir Cell Mol Biol (2013) 48(6):773–80. doi:10.1165/
rcmb.2012-0263OC
18. Duan M, Li WC, Vlahos R, Maxwell MJ, Anderson GP, Hibbs ML. Distinct
macrophage subpopulations characterize acute infection and chronic inflammatory lung disease. J Immunol (2012) 189(2):946–55. doi:10.4049/jimmunol.
1200660
19. Geissmann F, Jung S, Littman DR. Blood monocytes consist of two principal subsets with distinct migratory properties. Immunity (2003) 19(1):71–82.
doi:10.1016/S1074-7613(03)00174-2
20. Taylor PR, Martinez-Pomares L, Stacey M, Lin HH, Brown GD, Gordon S.
Macrophage receptors and immune recognition. Annu Rev Immunol (2005)
23:901–44. doi:10.1146/annurev.immunol.23.021704.115816
21. Nathan CF, Hibbs JB Jr. Role of nitric oxide synthesis in macrophage antimicrobial activity. Curr Opin Immunol (1991) 3(1):65–70. doi:10.1016/0952-7915(91)
90079-G
22. Fantone JC, Ward PA. Role of oxygen-derived free radicals and metabolites in leukocyte-dependent inflammatory reactions. Am J Pathol (1982)
107(3):395–418.
23. Hickman-Davis JM, O’Reilly P, Davis IC, Peti-Peterdi J, Davis G, Young KR, et al.
Killing of Klebsiella pneumoniae by human alveolar macrophages. Am J Physiol
Lung Cell Mol Physiol (2002) 282:L944–56. doi:10.1152/ajplung.00216.2001
24. Hashimoto S, Pittet JF, Hong K, Folkesson H, Bagby G, Kobzik L, et al. Depletion of alveolar macrophages decreases neutrophil chemotaxis to Pseudomonas
airspace infections. Am J Physiol (1996) 270(5 Pt 1):L819–28.
25. Maus UA, Koay MA, Delbeck T, Mack M, Ermert M, Ermert L, et al. Role of
resident alveolar macrophages in leukocyte traffic into the alveolar air space
of intact mice. Am J Physiol Lung Cell Mol Physiol (2002) 282(6):L1245–52.
doi:10.1152/ajplung.00453.2001
26. Dhaliwal K, Scholefield E, Ferenbach D, Gibbons M, Duffin R, Dorward DA,
et al. Monocytes control second-phase neutrophil emigration in established
lipopolysaccharide-induced murine lung injury. Am J Respir Crit Care Med
(2012) 186(6):514–24. doi:10.1164/rccm.201112-2132OC
27. Beck-Schimmer B, Schwendener R, Pasch T, Reyes L, Booy C, Schimmer RC.
Alveolar macrophages regulate neutrophil recruitment in endotoxin-induced
lung injury. Respir Res (2005) 6:61. doi:10.1186/1465-9921-6-61

Frontiers in Immunology | Molecular Innate Immunity

Alveolar macrophages in COPD

28. Knapp S, Leemans JC, Florquin S, Branger J, Maris NA, Pater J, et al. Alveolar macrophages have a protective antiinflammatory role during murine
pneumococcal pneumonia. Am J Respir Crit Care Med (2003) 167(2):171–9.
doi:10.1164/rccm.200207-698OC
29. Cox G, Crossley J, Xing Z. Macrophage engulfment of apoptotic neutrophils contributes to the resolution of acute pulmonary inflammation
in vivo. Am J Respir Cell Mol Biol (1995) 12(2):232–7. doi:10.1165/ajrcmb.12.2.
7865221
30. Fadok VA, Bratton DL, Konowal A, Freed PW, Westcott JY, Henson PM.
Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production through autocrine/paracrine mechanisms involving
TGF-beta, PGE2, and PAF. J Clin Invest (1998) 101(4):890–8. doi:10.1172/
JCI1112
31. Tate MD, Pickett DL, van Rooijen N, Brooks AG, Reading PC. Critical role
of airway macrophages in modulating disease severity during influenza virus
infection of mice. J Virol (2010) 84(15):7569–80. doi:10.1128/JVI.00291-10
32. Ghoneim HE, Thomas PG, McCullers JA. Depletion of alveolar macrophages
during influenza infection facilitates bacterial superinfections. J Immunol (2013)
191(3):1250–9. doi:10.4049/jimmunol.1300014
33. Sun K, Metzger DW. Inhibition of pulmonary antibacterial defense by
interferon-gamma during recovery from influenza infection. Nat Med (2008)
14(5):558–64. doi:10.1038/nm1765
34. Arredouani M, Yang Z, Ning Y, Qin G, Soininen R, Tryggvason K, et al. The
scavenger receptor MARCO is required for lung defense against pneumococcal pneumonia and inhaled particles. J Exp Med (2004) 200(2):267–72.
doi:10.1084/jem.20040731
35. Lopez AD, Murray CC. The global burden of disease, 1990-2020. Nat Med (1998)
4(11):1241–3. doi:10.1038/3218
36. Barnes PJ, Shapiro SD, Pauwels RA. Chronic obstructive pulmonary disease: molecular and cellular mechanisms. Eur Respir J (2003) 22(4):672–88.
doi:10.1183/09031936.03.00040703
37. Dennis RJ, Maldonado D, Norman S, Baena E, Martinez G. Woodsmoke exposure and risk for obstructive airways disease among women. Chest (1996)
109(1):115–9. doi:10.1378/chest.109.1.115
38. Pauwels RA, Buist AS, Calverley PM, Jenkins CR, Hurd SS, Committee GS. Global
strategy for the diagnosis, management, and prevention of chronic obstructive pulmonary disease. NHLBI/WHO global initiative for chronic obstructive
lung disease (GOLD) workshop summary. Am J Respir Crit Care Med (2001)
163(5):1256–76. doi:10.1164/ajrccm.163.5.2101039
39. Shapiro SD. The macrophage in chronic obstructive pulmonary disease. Am
J Respir Crit Care Med (1999) 160(5 Pt 2):S29–32. doi:10.1164/ajrccm.160.
supplement_1.9
40. Pesci A, Balbi B, Majori M, Cacciani G, Bertacco S, Alciato P, et al. Inflammatory cells and mediators in bronchial lavage of patients with chronic obstructive
pulmonary disease. Eur Respir J (1998) 12(2):380–6. doi:10.1183/09031936.98.
12020380
41. Keatings VM, Collins PD, Scott DM, Barnes PJ. Differences in interleukin-8
and tumor necrosis factor-alpha in induced sputum from patients with chronic
obstructive pulmonary disease or asthma. Am J Respir Crit Care Med (1996)
153(2):530–4. doi:10.1164/ajrccm.153.2.8564092
42. Retamales I, Elliott WM, Meshi B, Coxson HO, Pare PD, Sciurba FC, et al. Amplification of inflammation in emphysema and its association with latent adenoviral infection. Am J Respir Crit Care Med (2001) 164(3):469–73. doi:10.1164/
ajrccm.164.3.2007149
43. Di Stefano A, Capelli A, Lusuardi M, Balbo P, Vecchio C, Maestrelli P, et al. Severity of airflow limitation is associated with severity of airway inflammation in
smokers. Am J Respir Crit Care Med (1998) 158(4):1277–85. doi:10.1164/ajrccm.
158.4.9802078
44. Beckett EL, Stevens RL, Jarnicki AG, Kim RY, Hanish I, Hansbro NG, et al. A
new short-term mouse model of chronic obstructive pulmonary disease identifies a role for mast cell tryptase in pathogenesis. J Allergy Clin Immunol (2013)
131(3):752–62. doi:10.1016/j.jaci.2012.11.053
45. Finlay GA, O’Driscoll LR, Russell KJ, D’Arcy EM, Masterson JB, FitzGerald
MX, et al. Matrix metalloproteinase expression and production by alveolar
macrophages in emphysema. Am J Respir Crit Care Med (1997) 156(1):240–7.
doi:10.1164/ajrccm.156.1.9612018

September 2014 | Volume 5 | Article 435 | 192

Vlahos and Bozinovski

46. Ohnishi K, Takagi M, Kurokawa Y, Satomi S, Konttinen YT. Matrix
metalloproteinase-mediated extracellular matrix protein degradation in human
pulmonary emphysema. Lab Invets (1998) 78(9):1077–87.
47. Lim S, Roche N, Oliver BG, Mattos W, Barnes PJ, Chung KF. Balance
of matrix metalloprotease-9 and tissue inhibitor of metalloprotease-1 from
alveolar macrophages in cigarette smokers. Regulation by interleukin-10.
Am J Respir Crit Care Med (2000) 162(4 Pt 1):1355–60. doi:10.1164/ajrccm.
162.4.9910097
48. Russell RE, Culpitt SV, DeMatos C, Donnelly L, Smith M, Wiggins J, et al.
Release and activity of matrix metalloproteinase-9 and tissue inhibitor of
metalloproteinase-1 by alveolar macrophages from patients with chronic
obstructive pulmonary disease. Am J Respir Cell Mol Biol (2002) 26(5):602–9.
doi:10.1165/ajrcmb.26.5.4685
49. Monso E, Ruiz J, Rosell A, Manterola J, Fiz J, Morera J, et al. Bacterial infection in
chronic obstructive pulmonary disease. A study of stable and exacerbated outpatients using the protected specimen brush. Am J Respir Crit Care Med (1995)
152(4 Pt 1):1316–20. doi:10.1164/ajrccm.152.4.7551388
50. Pela R, Marchesani F, Agostinelli C, Staccioli D, Cecarini L, Bassotti C, et al. Airways microbial flora in COPD patients in stable clinical conditions and during
exacerbations: a bronchoscopic investigation. Monaldi Arch Chest Dis (1998)
53(3):262–7.
51. Papi A, Bellettato CM, Braccioni F, Romagnoli M, Casolari P, Caramori G,
et al. Infections and airway inflammation in chronic obstructive pulmonary disease severe exacerbations. Am J Respir Crit Care Med (2006) 173(10):1114–21.
doi:10.1164/rccm.200506-859OC
52. Hurst JR, Vestbo J, Anzueto A, Locantore N, Mullerova H, Tal-Singer R, et al.
Susceptibility to exacerbation in chronic obstructive pulmonary disease. N Engl
J Med (2010) 363(12):1128–38. doi:10.1056/NEJMoa0909883
53. Birrell MA, Wong S, Catley MC, Belvisi MG. Impact of tobacco-smoke on key
signaling pathways in the innate immune response in lung macrophages. J Cell
Physiol (2008) 214(1):27–37. doi:10.1002/jcp.21158
54. Muller T, Gebel S. The cellular stress response induced by aqueous extracts of
cigarette smoke is critically dependent on the intracellular glutathione concentration. Carcinogenesis (1998) 19(5):797–801. doi:10.1093/carcin/19.5.797
55. Rahman I, MacNee W. Oxidative stress and regulation of glutathione in lung
inflammation. Eur Respir J (2000) 16(3):534–54. doi:10.1034/j.1399-3003.2000.
016003534.x
56. Marti-Lliteras P, Regueiro V, Morey P, Hood DW, Saus C, Sauleda J, et al.
Nontypeable Haemophilus influenzae clearance by alveolar macrophages is
impaired by exposure to cigarette smoke. Infect Immun (2009) 77(10):4232–42.
doi:10.1128/IAI.00305-09
57. Hodge S, Hodge G, Ahern J, Jersmann H, Holmes M, Reynolds PN. Smoking alters alveolar macrophage recognition and phagocytic ability: implications
in chronic obstructive pulmonary disease. Am J Respir Cell Mol Biol (2007)
37(6):748–55. doi:10.1165/rcmb.2007-0025OC
58. Hodge S, Matthews G, Mukaro V, Ahern J, Shivam A, Hodge G, et al. Cigarette smoke-induced changes to alveolar macrophage phenotype and function
are improved by treatment with procysteine. Am J Respir Cell Mol Biol (2011)
44(5):673–81. doi:10.1165/rcmb.2009-0459OC
59. Richens TR, Linderman DJ, Horstmann SA, Lambert C, Xiao YQ, Keith RL, et al.
Cigarette smoke impairs clearance of apoptotic cells through oxidant-dependent
activation of RhoA. Am J Respir Crit Care Med (2009) 179(11):1011–21.
doi:10.1164/rccm.200807-1148OC
60. Minemetsu N, Blumental-Perry A, Shapiro SD. Cigarette smoke inhibits engulfment of apoptotic cells by macrophages through inhibition of actin rearrangement. Am J Respir Cell Mol Biol (2011) 44(4):474–82. doi:10.1165/rcmb.20090463OC
61. Damiano VV, Tsang A, Kucich U, Abrams WR, Rosenbloom J, Kimbel P, et al.
Immunolocalization of elastase in human emphysematous lungs. J Clin Invest
(1986) 78(2):482–93. doi:10.1172/JCI112600
62. Shao MX, Nadel JA. Neutrophil elastase induces MUC5AC mucin production in human airway epithelial cells via a cascade involving protein kinase C,
reactive oxygen species, and TNF-alpha-converting enzyme. J Immunol (2005)
175(6):4009–16. doi:10.4049/jimmunol.175.6.4009
63. Liu K, Gualano RC, Hibbs ML, Anderson GP, Bozinovski S. Epidermal growth
factor receptor signaling to Erk1/2 and STATs control the intensity of the
epithelial inflammatory responses to rhinovirus infection. J Biol Chem (2008)
283(15):9977–85. doi:10.1074/jbc.M710257200

www.frontiersin.org

Alveolar macrophages in COPD

64. Banan A, Zhang Y, Losurdo J, Keshavarzian A. Carbonylation and disassembly of the F-actin cytoskeleton in oxidant induced barrier dysfunction and its
prevention by epidermal growth factor and transforming growth factor alpha in
a human colonic cell line. Gut (2000) 46(6):830–7. doi:10.1136/gut.46.6.830
65. Smerjac SM, Bizzozero OA. Cytoskeletal protein carbonylation and degradation in experimental autoimmune encephalomyelitis. J Neurochem (2008)
105(3):763–72. doi:10.1111/j.1471-4159.2007.05178.x
66. Bozinovski S, Vlahos R, Zhang Y, Lah LC, Seow HJ, Mansell A, et al. Carbonylation caused by cigarette smoke extract is associated with defective macrophage
immunity. Am J Respir Cell Mol Biol (2011) 45(2):229–36. doi:10.1165/rcmb.
2010-0272OC
67. Kirkham PA, Spooner G, Rahman I, Rossi AG. Macrophage phagocytosis of
apoptotic neutrophils is compromised by matrix proteins modified by cigarette
smoke and lipid peroxidation products. Biochem Biophys Res Commun (2004)
318(1):32–7. doi:10.1016/j.bbrc.2004.04.003
68. Barnes PJ. Alveolar macrophages as orchestrators of COPD. COPD (2004)
1(1):59–70. doi:10.1081/COPD-120028701
69. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
70. Agusti AG, Villaverde JM, Togores B, Bosch M. Serial measurements of exhaled
nitric oxide during exacerbations of chronic obstructive pulmonary disease. Eur
Respir J (1999) 14(3):523–8. doi:10.1034/j.1399-3003.1999.14d34.x
71. Bhowmik A, Seemungal TA, Donaldson GC, Wedzicha JA. Effects of exacerbations and seasonality on exhaled nitric oxide in COPD. Eur Respir J (2005)
26(6):1009–15. doi:10.1183/09031936.05.00047305
72. Maziak W, Loukides S, Culpitt S, Sullivan P, Kharitonov SA, Barnes PJ. Exhaled
nitric oxide in chronic obstructive pulmonary disease. Am J Respir Crit Care
Med (1998) 157(3 Pt 1):998–1002. doi:10.1164/ajrccm.157.3.97-05009
73. Maarsingh H, Pera T, Meurs H. Arginase and pulmonary diseases. Naunyn
Schmiedebergs Arch Pharmacol (2008) 378(2):171–84. doi:10.1007/s00210-0080286-7
74. Gebel S, Gerstmayer B, Kuhl P, Borlak J, Meurrens K, Muller T. The kinetics of
transcriptomic changes induced by cigarette smoke in rat lungs reveals a specific
program of defense, inflammation, and circadian clock gene expression. Toxicol
Sci (2006) 93(2):422–31. doi:10.1093/toxsci/kfl071
75. Ichinose M, Sugiura H, Yamagata S, Koarai A, Shirato K. Increase in reactive
nitrogen species production in chronic obstructive pulmonary disease airways.
Am J Respir Crit Care Med (2000) 162(2 Pt 1):701–6. doi:10.1164/ajrccm.162.2.
9908132
76. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity.
J Innate Immun (2014) 6(6). doi:10.1159/000364945
77. Woodruff PG, Koth LL, Yang YH, Rodriguez MW, Favoreto S, Dolganov GM,
et al. A distinctive alveolar macrophage activation state induced by cigarette
smoking. Am J Respir Crit Care Med (2005) 172(11):1383–92. doi:10.1164/rccm.
200505-686OC
78. Shaykhiev R, Krause A, Salit J, Strulovici-Barel Y, Harvey BG, O’Connor TP,
et al. Smoking-dependent reprogramming of alveolar macrophage polarization: implication for pathogenesis of chronic obstructive pulmonary disease.
J Immunol (2009) 183(4):2867–83. doi:10.4049/jimmunol.0900473
79. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
80. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
81. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine
system in diverse forms of macrophage activation and polarization. Trends
Immunol (2004) 25(12):677–86. doi:10.1016/j.it.2004.09.015
82. Van Gorp H, Delputte PL, Nauwynck HJ. Scavenger receptor CD163, a Jackof-all-trades and potential target for cell-directed therapy. Mol Immunol (2010)
47(7–8):1650–60. doi:10.1016/j.molimm.2010.02.008
83. Kunz LI, Lapperre TS, Snoeck-Stroband JB, Budulac SE, Timens W, van Wijngaarden S, et al. Smoking status and anti-inflammatory macrophages in bronchoalveolar lavage and induced sputum in COPD. Respir Res (2011) 12:34.
doi:10.1186/1465-9921-12-34
84. Dolinay T, Choi AM, Ryter SW. Heme oxygenase-1/CO as protective mediators in cigarette smoke-induced lung cell injury and chronic obstructive pulmonary disease. Curr Pharm Biotechnol (2012) 13(6):769–76. doi:10.2174/
138920112800399338

September 2014 | Volume 5 | Article 435 | 193

Vlahos and Bozinovski

85. Williams L, Jarai G, Smith A, Finan P. IL-10 expression profiling in human
monocytes. J Leukoc Biol (2002) 72(4):800–9.
86. Takanashi S, Hasegawa Y, Kanehira Y, Yamamoto K, Fujimoto K, Satoh K,
et al. Interleukin-10 level in sputum is reduced in bronchial asthma, COPD
and in smokers. Eur Respir J (1999) 14(2):309–14. doi:10.1183/09031936.99.
14230999
87. Anthony D, McQualter JL, Bishara M, Lim EX, Yatmaz S, Seow HJ,
et al. SAA drives proinflammatory heterotypic macrophage differentiation in
the lung via CSF-1R-dependent signaling. FASEB J (2014). doi:10.1096/fj.14250332
88. Bozinovski S, Uddin M, Vlahos R, Thompson M, McQualter JL, Merritt AS, et al.
Serum amyloid A opposes lipoxin A(4) to mediate glucocorticoid refractory lung
inflammation in chronic obstructive pulmonary disease. Proc Natl Acad Sci U S
A (2012) 109(3):935–40. doi:10.1073/pnas.1109382109
89. Bozinovski S, Anthony D, Anderson GP, Irving LB, Levy BD, Vlahos R. Treating neutrophilic inflammation in COPD by targeting ALX/FPR2 resolution
pathways. Pharmacol Ther (2013) 140(3):280–9. doi:10.1016/j.pharmthera.
2013.07.007
90. El Kebir D, Jozsef L, Khreiss T, Pan W, Petasis NA, Serhan CN, et al. Aspirintriggered lipoxins override the apoptosis-delaying action of serum amyloid
A in human neutrophils: a novel mechanism for resolution of inflammation.
J Immunol (2007) 179(1):616–22. doi:10.4049/jimmunol.179.1.616

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Alveolar macrophages in COPD

91. Anthony D, Seow HJ, Uddin M, Thompson M, Dousha L, Vlahos R, et al. Serum
amyloid A promotes lung neutrophilia by increasing IL-17A levels in the mucosa
and gammadelta T cells. Am J Respir Crit Care Med (2013) 188(2):179–86.
doi:10.1164/rccm.201211-2139OC
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 22 July 2014; paper pending published: 18 August 2014; accepted: 26 August
2014; published online: 10 September 2014.
Citation: Vlahos R and Bozinovski S (2014) Role of alveolar macrophages
in chronic obstructive pulmonary disease. Front. Immunol. 5:435. doi:
10.3389/fimmu.2014.00435
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Vlahos and Bozinovski. This is an open-access article distributed
under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s)
or licensor are credited and that the original publication in this journal is cited, in
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permitted which does not comply with these terms.

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MINI REVIEW ARTICLE
published: 26 September 2014
doi: 10.3389/fimmu.2014.00470

Macrophage polarization in obesity and type 2 diabetes:
weighing down our understanding of macrophage
function?
Michael James Kraakman 1,2 , Andrew James Murphy 2,3 , Karin Jandeleit-Dahm 4,5 † and
Hélène L. Kammoun 1 *†
1
2
3
4
5

Cellular and Molecular Metabolism Laboratory, Baker IDI Heart and Diabetes Institute, Melbourne, VIC, Australia
Haematopoiesis and Leukocyte Biology Laboratory, Baker IDI Heart and Diabetes Institute, Melbourne, VIC, Australia
Department of Immunology, Monash University, Melbourne, VIC, Australia
Diabetic Complications Laboratory, Baker IDI Heart and Diabetes Institute, Melbourne, VIC, Australia
Department of Medicine, Central Clinical School, Monash University, Melbourne, VIC, Australia

Edited by:
Klaus Ley, La Jolla Institute for Allergy
and Immunology, USA
Reviewed by:
Paul Quax, Leiden University Medical
Center, Netherlands
Philippe Georgel, Strasbourg
University, France
*Correspondence:
Hélène L. Kammoun, Baker IDI Heart
and Diabetes Institute, P.O. Box 6492,
St Kilda Road Central, Melbourne
3004, VIC, Australia
e-mail: helene.kammoun@
bakeridi.edu.au
†

Co-senior authors.

Obesity and type 2 diabetes are now recognized as chronic pro-inflammatory diseases.
In the last decade, the role of the macrophage in particular has become increasingly
implicated in their pathogenesis. Abundant literature now establishes that monocytes
get recruited to peripheral tissues (i.e., pancreas, liver, and adipose tissue) to become
resident macrophages and contribute to local inflammation, development of insulin resistance, or even pancreatic dysfunction. Furthermore, an accumulation of evidence has
established an important role for macrophage polarization in the development of metabolic
diseases. The general view in obesity is that there is an imbalance in the ratio of M1/M2
macrophages, with M1 “pro-inflammatory” macrophages being enhanced compared with
M2 “anti-inflammatory” macrophages being down-regulated, leading to chronic inflammation and the propagation of metabolic dysfunction. However, there is emerging evidence
revealing a more complex scenario with the spectrum of macrophage states exceeding
well beyond the M1/M2 binary classification and confused further by human and animal
models exhibiting different macrophage profiles. In this review, we will discuss the recent
findings regarding macrophage polarization in obesity and type 2 diabetes.
Keywords: macrophage, inflammation, obesity, type 2 diabetes, M1/M2, polarization

Inflammation is a fundamental biological process whose role is
not only to enable host protection against pathogens, but also to
stimulate and modulate repair and healing when cellular damage
occurs. During an inflammatory event, once the initial insult is
contained, a primary objective is the restoration of tissue homeostasis. Failure to appropriately resolve an inflammatory stimulus
can result in persistent immune system activation, which can actually cause tissue damage and disease. Significant literature over the
last decades has established obesity to induce a state of chronic
low-grade systemic inflammation (1). Importantly, the inflammation accompanying obesity is distinctly different to that of acute
inflammation, as the inflammatory stimulus fails to be resolved.
This is of particular significance as chronic low-grade inflammation is implicated in the etiology of atherosclerosis, hypertension,
type 2 diabetes (T2D), and even certain cancers, which can all
be associated with obesity (2, 3). Considering the economic burden from the increasing prevalence of these chronic metabolic
diseases, it is not surprising that considerable scientific attention
has focused on how and why obesity promotes chronic low-grade
inflammation.
Immune cells are the primary effectors of most inflammatory
reactions and are categorized into the innate and acquired immune
systems. With respect to obesity, leukocytes from both immune

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systems have been implicated in the development of chronic
low-grade inflammation and metabolic dysfunction (1). However, adipose tissue macrophages have received the lion’s share of
attention of the immune cells involved. Macrophages can display
remarkable phenotypic heterogeneity with the ability to perform
vastly different roles depending on the biological situation (4).
Thus, a spectrum of many different macrophage populations has
been characterized by combinations of membrane markers and
gene expression profiles. This led to the establishment of a complex nomenclature, which over time has become simplified into
two main macrophage phenotypes, M1 pro-inflammatory and M2
anti-inflammatory macrophages. This classification is obviously a
simplistic view of the situation as it is now clear that macrophages
dynamically evolve from one phenotype to the other according to
their environment and can occupy various points of the spectrum
with mixed characteristics [for updated nomenclature see Ref. (5,
6)]. For clarity purposes, we will mostly mention macrophages as
either M1 or M2.

MACROPHAGE ACCUMULATION IN THE OBESE ADIPOSE
TISSUE
Phagocytosis is a main function of macrophages that allows
them to contribute to tissue homeostasis through surveillance,

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Kraakman et al.

maintenance, and repair. Though long-lived macrophage populations reside within almost all bodily tissues, in 2003, two
separate laboratories reported macrophage accumulation in the
white adipose tissue (WAT) of both obese patients and rodents
(7, 8). Importantly, this was completely in line with Hotamisligil’s landmark article 10 years earlier observing TNFα secretion
from the adipose tissue in obese rodents (9). Indeed, it appeared
these macrophages were the source of the elevated inflammatory
cytokines reported in obesity and their accumulation was associated with insulin resistance (7, 8). These landmark studies were
the first to link obesity and insulin resistance with adipose tissue
macrophage content and inflammation. Later it appeared that the
accumulation of macrophages is not limited to the WAT in obesity
with macrophages found to accumulate in many other organs critical for glucose homeostasis such as liver, pancreas, gut, and even the
brain (3). Regardless, the most significant immunological changes
occurring during obesity originate within the adipose tissue. Of
interest, despite the adipose tissue representing a small portion
of whole body glucose disposal, immune dysfunction within this
tissue is sufficient to impair systemic glucose metabolism (1).

MACROPHAGE POLARIZATION IN OBESITY AND T2D
Macrophages constitute an important fraction of non-adipocyte
cells within the WAT. Indeed, within a “normal” lean WAT, they
can account for almost 10% of total cell number. Profiling of
lean adipose tissue revealed these macrophages to possess an
M2 phenotype. These macrophages perform tissue surveillance
and remodeling functions and are associated with maintaining
WAT insulin sensitivity. Indeed, the manipulation of peroxisome proliferator activator receptor (PPAR) transcription factors
required for macrophage M2 polarization was associated with
metabolic dysfunction (10–12). The current theory supports that
weight gain induces local inflammation and chemokine production to promote recruitment of circulating pro-inflammatory
(Ly6Chi ) monocytes. Recruited monocytes differentiate into an
M1 macrophage phenotype and their accumulation leads to
an imbalance between M1 and M2 macrophages. Increased
cytokine production from M1 macrophages and/or reduced antiinflammatory signals from the M2 macrophages promote adipose
tissue dysfunction and impairs glucose tolerance.
Evidence for the detrimental role of M1 macrophages in promoting adipose tissue insulin resistance has been reported in
several studies. Lumeng and colleagues first reported that the
macrophages accumulating in obese WAT possessed an inflammatory CD11c+ M1 phenotype and gathered around necrotic
adipocytes in Crown like structures (CLS) (13, 14). Furthermore, M1 macrophage numbers were shown to correlate with
insulin resistance in high-fat fed rodents (15). Whether WAT
M1 macrophages could be targeted therapeutically has proven
more challenging. Clodronate is a toxic compound administered in liposomes that gets taken up by macrophages thereby
inducing their apoptosis. Clodronate liposome injections succeed in decreasing visceral WAT macrophage accumulation and
improve glucose metabolism and insulin sensitivity in HFD-fed
mice (16). However, this approach has limitations as clodronate
liposomes targeted all WAT macrophage phenotypes as well as

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Macrophage polarization in metabolic diseases

liver Kuppfer cells. One could envisage a deleterious effect from
depleting M2 macrophages on a long term basis. A more specific
removal of M1 macrophages was achieved in an elegant study
published by Olefsky’s group. They demonstrated that ablation
of M1 macrophages, achieved by targeting diphtheria-sensitive
CD11c+ cells with diphtheria toxin, was associated with improved
glucose tolerance (17). While CD11c expression allows discrimination between M1 and M2 macrophages, its expression is not
exclusive for M1 macrophages. Dendritic cells and neutrophils also
express CD11c and the elevation of these cells in obese WAT may
contribute to the increased CD11c+ cell population and insulin
resistance (18, 19). From these studies, it is clear that targeting
established M1 populations within obese WAT will prove complex.
For these reasons, studies investigating mechanisms leading to
macrophage accumulation may yield more promising therapeutic
targets.

POTENTIAL MECHANISMS FOR MACROPHAGE
ACCUMULATION IN OBESITY
White adipose tissue macrophage accumulation is thought to
occur through two main processes. First, increased chemokine
secretion from adipocytes and resident macrophages promotes
the recruitment of Ly6C+ blood monocytes to obese WAT. Most of
these monocytes subsequently differentiate into M1 macrophages
in response to inflammatory signals within the adipose tissue. Of the chemokines produced from obese WAT, monocyte
chemoattractant protein-1(MCP-1) and its receptor C–C motif
receptor-2 (CCR2) appear particularly important. For example,
mice lacking either CCR2 or MCP-1 have reduced ATMs, whereas
adipocyte specific over-expression of MCP-1 leads to enhanced
ATMs (20, 21). However, these knockout models did not normalize macrophage numbers suggesting additional mechanisms are
likely to be involved in obesity-induced macrophage accumulation (20). Indeed, obese WAT secretes many chemokines including
LTB4, MIP, MIF, and MCP-3 implicated in macrophage accumulation and glucose intolerance (22). Interestingly, in addition
to chemokine secretion, signals from obese WAT also influence
bone marrow progenitor cells to increase myelopoiesis. Indeed, we
have recently demonstrated that in obese mice, IL1β production
from CD11c+ ATMs promoted bone marrow myelopoiesis further
perpetuating adipose tissue inflammation (23). Second, resident
ATMs have a strong proliferation capacity in both human beings
and rodents. Indeed, Jenkins et al. showed that IL4 is a strong promoter of macrophage proliferation (24). Hence, in the lean state
where eosinophils secrete high levels of IL4 in the WAT, proliferation is considered the major mechanism to maintain resident
M2 macrophage populations (24–26). Interestingly, Amano et al.
revealed that MCP-1 could enhance macrophage proliferation in
the visceral WAT independent of its chemokine function (27). It
is clear that further studies are required to determine the contribution of macrophage proliferation and recruitment to adipose
tissue macrophage accumulation in obesity.

MACROPHAGE FATE
While the recruitment and source of macrophages present in obese
adipose tissue are well documented, the fate of these recruited

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macrophages remains less studied. Unlike resolving inflammation in which levels of inflammatory leukocytes subside following
the restoration of tissue homeostasis, adipose tissue inflammation
induced by excessive adiposity fails to resolve naturally. During
resolving inflammation, it is well appreciated that the initially
recruited polymorphonuclear neutrophils undergo swift apoptosis
prompting their phagocytosis by macrophages. Despite early evidence for macrophage emigration and drainage to lymph nodes as
a significant contributor to macrophage disappearance following
acute inflammation, recent work from the Randolph laboratory
has revealed that macrophage apoptosis is largely responsible
for their removal in acute inflammation (28). Whether these
mechanisms contribute to adipose tissue macrophage accumulation in non-resolving inflammation is not well studied. For
example, whether M1 macrophages accumulate within obese adipose tissue due to pro-survival/anti-apoptotic signals remains
unknown. However, it is also possible that macrophages are
actively retained within the adipose tissue in response to various
cues. Netrin-1, best described for its role in neural development,
has recently been implicated in macrophage retention within obese
adipose tissue (29). However, these findings are somewhat at
odds with the plethora of studies showing the importance of
monocyte/macrophage recruitment, repopulation (e.g., after bone
marrow transplantation), and proliferation to the macrophage
burden in the obese adipose tissue and perhaps plays only a
minor role.

A MORE COMPLEX PICTURE THAN M1 OR M2
Macrophages are able to modify their phenotype according
to their environment. However, whether macrophage phenotype switching occurs in obesity remains unresolved (30, 31).
Shaul et al. demonstrated that in HFD-fed animals’ classical M1
macrophage accumulation is observed after 8 weeks of diet, however, after 12 weeks of diet, these CD11c+ macrophages exhibited an increased expression of M2 associated transcripts (32).
While the M1 macrophages maintained their pro-inflammatory
phenotype, they also adopted some remodeling features in a context of increased adipogenesis reminiscent of M2. Conversely,
M2 macrophages are able to secrete pro-inflammatory mediators in specific conditions (33). These studies reveal extremely
dynamic macrophage populations and newer technologies such
as live imaging undoubtedly will enhance our understanding of
the changes occurring in macrophage polarization state upon
weight gain. Indeed, Haase et al. showed recently that most adipose tissue macrophages arising in situ stained positive for the
M2 markers CD206 and CD301 (34). Live imagery and tracking techniques allowed the revelation that newly formed M2
macrophages originates within the CLS before they migrate to the
interstitial space. Given that CLS form around dying adipocytes,
the presence of M2 macrophages could be viewed as a resolving
mechanism. Given that macrophages are able to alter their phenotype and that most studies assess adipose tissue macrophage
content at one single time, care must be exercised when interpreting data. For example, one cannot expect to predict a metabolic phenotype based upon observed adipose tissue macrophage
polarity.

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Macrophage polarization in metabolic diseases

It remains plausible that the inflammation associated with obesity initially constitutes a physiological rather than pathological
process within the adipose tissue. Indeed, in an elegant study,
Scherer and colleagues have demonstrated that adipocyte inflammation is essential for adipose tissue expansion and remodeling
(35). Using the “adipochaser mouse,” they tracked newly formed
adipocytes and distinguished them from older “blue” adipocytes
expressing β-galactosidase. They determined that acute inflammation promotes adipogenesis and improved adipose tissue function
and insulin sensitivity. Conversely, they demonstrated that abrogation of inflammation within the adipose tissue led to defective
adipogenesis followed by ectopic lipid accumulation and glucose
intolerance (35). These findings highlight a previously unappreciated role for inflammation per se in healthy adipose tissue
function.

CAN WE TARGET IMMUNITY TO TREAT T2D?
It is important to ensure that the findings made in rodent models are useful in human pathology. It is clear that macrophages
also accumulate in adipose tissue of obese humans (7) and have
been correlated with insulin resistance (36).Wentworth et al. also
showed that pro-inflammatory CD11c+ macrophages are positively associated with systemic insulin resistance in obese patients
(37). Furthermore, macrophage content is reduced in the adipose
tissue following gastric bypass surgery (38). Importantly, in these
patients, the macrophage status was switched toward a less proinflammatory profile. However, other groups have shown that the
accrued macrophages in human adipose tissue present M2 surface
markers associated with a more anti-inflammatory phenotype (39,
40). Nonetheless, Zeyda et al. showed that these M2 macrophages
possess a strong capacity to produce pro-inflammatory mediators (39). These discrepancies may be explained by different
experimental protocols. In most studies, WAT is obtained from
subcutaneous depots whereas fewer studies report data obtained
from omental WAT. Furthermore, the sex and degree of adiposity of the patients may also account for some of the differences
reported. The same reasons may partly explain the inconsistencies
observed between mice and human beings. In addition, important metabolic differences between rodents and human beings
contribute to the difficulty of translating mouse data into human
therapies.
In addition, there are further evidences that inflammation and
macrophages should be targeted with the greatest care in T2D.
Indeed, Chawla’s group recently reported that M2 macrophages
can secrete catecholamines in response to cold exposure, activating thermogenesis of brown adipose tissue and lipolysis of WAT
(41). They later found that M2 macrophages were directly involved
in the “beginning” of adipose tissue (42). In addition, a current
article proposed that M2 macrophages strongly promote β-cell
proliferation (43). The presented studies tend to suggest that the
positive effects of macrophages in obesity and T2D are carried
on by repair macrophages exhibiting an M2 phenotype. Hence,
there appears to be rationale targeting the enhancement of M2
populations as opposed to depleting M1 macrophages, which may
jeopardize the patient’s immune function. However, increased M2
macrophage function is also tightly linked to tumor proliferation

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Kraakman et al.

FIGURE 1 | Immune cells modulation in adipose tissue during obesity. In
the lean healthy adipose tissue, M2 like macrophages self-maintain through
proliferation. They ensure tissue remodeling and pathogen screening. In the
obese adipose tissue, overnutrition leads to larger adipocytes, which coupled
with various intrinsic and extrinsic cellular stress, and promotes the

so manipulating M2 populations must be considered carefully
as obese patients already present with an increased incidence of
cancer (2).

CONCLUSION
It is increasingly appreciated that immune cells play a crucial role
in the control of whole body metabolism (2). In light of this view,
it seems clear that the macrophage accumulation within the adipose tissue is associated with metabolic dysfunction observed in
obesity. However, the role of macrophages in the pathogenesis of
obesity and diabetes is still conflicting. In addition to the complexity of the immune response itself, different rodent models,
the use of different high-fat diets and of different intervention
time courses not to add the different environments of each animal facility, have often led to contradictory results raising more
confusion within the field. As it was nicely exposed in a recent
review by Murray et al., it is important for the coherence of results
in this domain that researchers try and decide which models to
focus on to allow the field to progress and result in more human
translational potential (5).
In this review, we have summarized the recent findings on
the role of macrophages in obesity and T2D. M1 macrophage

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Macrophage polarization in metabolic diseases

development of a pro-inflammatory environment. In this context, M1 like
macrophages start to accumulate due to some M2 macrophages switching
phenotype, bone marrow stimulation of myelopoiesis contributing to the
recruitment of pro-inflammatory monocytes, and possibly reduced egress of
macrophages.

accumulation within the adipose tissue remains tightly associated
with obesity. It is still unclear how much this accumulation contributes to the glucose intolerance and insulin resistance described
in obese rodent models. There is growing literature suggesting
their presence could be required for physiological adaptations
of the adipose tissue (35). Perhaps these findings reflect the
dynamic nature of macrophage polarity and the essential role of
macrophages in the biology of adipose tissue (Figure 1). However,
it is also important to take a “non-glucose/insulin resistant-centric
view” with respect to the role of adipose tissue macrophages
and to appreciate that they could also contribute significantly
to the risk of other associated diseases including cardiovascular disease and cancer. As a whole, these data advocate that
macrophages should be targeted with the greatest care in metabolic
diseases.

REFERENCES
1. Kammoun HL, Kraakman MJ, Febbraio MA. Adipose tissue inflammation in
glucose metabolism. Rev Endocr Metab Disord (2014) 15(1):31–44. doi:10.1007/
s11154-013-9274-4
2. Noto H, Goto A, Tsujimoto T, Osame K, Noda M. Latest insights into the
risk of cancer in diabetes. J Diabetes Investig (2013) 4(3):225–32. doi:10.1111/
jdi.12068

September 2014 | Volume 5 | Article 470 | 198

Kraakman et al.

3. Osborn O, Olefsky JM. The cellular and signaling networks linking the
immune system and metabolism in disease. Nat Med (2012) 18(3):363–74.
doi:10.1038/nm.2627
4. Okabe Y, Medzhitov R. Tissue-specific signals control reversible program
of localization and functional polarization of macrophages. Cell (2014)
157(4):832–44. doi:10.1016/j.cell.2014.04.016
5. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
6. Guilliams M, Ginhoux F, Jakubzick C, Naik SH, Onai N, Schraml BU, et al.
Dendritic cells, monocytes and macrophages: a unified nomenclature based on
ontogeny. Nat Rev Immunol (2014) 14(8):571–8. doi:10.1038/nri3712
7. Weisberg SP, McCann D, Desai M, Rosenbaum M, Leibel RL, Ferrante AW Jr.
Obesity is associated with macrophage accumulation in adipose tissue. J Clin
Invest (2003) 112(12):1796–808. doi:10.1172/JCI19246
8. Xu H, Barnes GT, Yang Q, Tan G, Yang D, Chou CJ, et al. Chronic inflammation
in fat plays a crucial role in the development of obesity-related insulin resistance.
J Clin Invest (2003) 112(12):1821–30. doi:10.1172/JCI19451
9. Hotamisligil GS, Shargill NS, Spiegelman BM. Adipose expression of tumor
necrosis factor-alpha: direct role in obesity-linked insulin resistance. Science
(1993) 259(5091):87–91. doi:10.1126/science.7678183
10. Desvergne B. PPARdelta/beta: the lobbyist switching macrophage allegiance
in favor of metabolism. Cell Metab (2008) 7(6):467–9. doi:10.1016/j.cmet.2008.
05.002
11. Odegaard JI, Ricardo-Gonzalez RR, Goforth MH, Morel CR, Subramanian
V, Mukundan L, et al. Macrophage-specific PPARgamma controls alternative
activation and improves insulin resistance. Nature (2007) 447(7148):1116–20.
doi:10.1038/nature05894
12. Hevener AL, Olefsky JM, Reichart D, Nguyen MT, Bandyopadyhay G, Leung
HY, et al. Macrophage PPAR gamma is required for normal skeletal muscle
and hepatic insulin sensitivity and full antidiabetic effects of thiazolidinediones.
J Clin Invest (2007) 117(6):1658–69. doi:10.1172/JCI31561
13. Lumeng CN, Bodzin JL, Saltiel AR. Obesity induces a phenotypic switch in
adipose tissue macrophage polarization. J Clin Invest (2007) 117(1):175–84.
doi:10.1172/JCI29881
14. Lumeng CN, DelProposto JB, Westcott DJ, Saltiel AR. Phenotypic switching of adipose tissue macrophages with obesity is generated by spatiotemporal differences in macrophage subtypes. Diabetes (2008) 57(12):3239–46.
doi:10.2337/db08-0872
15. Fujisaka S, Usui I, Bukhari A, Ikutani M, Oya T, Kanatani Y, et al. Regulatory
mechanisms for adipose tissue M1 and M2 macrophages in diet-induced obese
mice. Diabetes (2009) 58(11):2574–82. doi:10.2337/db08-1475
16. Feng B, Jiao P, Nie Y, Kim T, Jun D, van Rooijen N, et al. Clodronate liposomes
improve metabolic profile and reduce visceral adipose macrophage content in
diet-induced obese mice. PLoS One (2011) 6(9):e24358. doi:10.1371/journal.
pone.0024358
17. Patsouris D, Li PP, Thapar D, Chapman J, Olefsky JM, Neels JG. Ablation of
CD11c-positive cells normalizes insulin sensitivity in obese insulin resistant
animals. Cell Metab (2008) 8(4):301–9. doi:10.1016/j.cmet.2008.08.015
18. Stefanovic-Racic M, Yang X, Turner MS, Mantell BS, Stolz DB, Sumpter TL,
et al. Dendritic cells promote macrophage infiltration and comprise a substantial proportion of obesity-associated increases in CD11c+ cells in adipose tissue
and liver. Diabetes (2012) 61(9):2330–9. doi:10.2337/db11-1523
19. Talukdar S, Oh da Y, Bandyopadhyay G, Li D, Xu J, McNelis J, et al. Neutrophils
mediate insulin resistance in mice fed a high-fat diet through secreted elastase.
Nat Med (2012) 18(9):1407–12. doi:10.1038/nm.2885
20. Weisberg SP, Hunter D, Huber R, Lemieux J, Slaymaker S, Vaddi K, et al. CCR2
modulates inflammatory and metabolic effects of high-fat feeding. J Clin Invest
(2006) 116(1):115–24. doi:10.1172/JCI24335C1
21. Kamei N, Tobe K, Suzuki R, Ohsugi M, Watanabe T, Kubota N, et al.
Overexpression of monocyte chemoattractant protein-1 in adipose tissues
causes macrophage recruitment and insulin resistance. J Biol Chem (2006)
281(36):26602–14. doi:10.1074/jbc.M601284200
22. Surmi BK, Hasty AH. The role of chemokines in recruitment of immune cells
to the artery wall and adipose tissue. Vascul Pharmacol (2010) 52(1–2):27–36.
doi:10.1016/j.vph.2009.12.004

www.frontiersin.org

Macrophage polarization in metabolic diseases

23. Nagareddy PR, Kraakman M, Masters SL, Stirzaker RA, Gorman DJ,
Grant RW, et al. Adipose tissue macrophages promote myelopoiesis and monocytosis in obesity. Cell Metab (2014) 19(5):821–35. doi:10.1016/j.cmet.2014.03.
029
24. Jenkins SJ, Ruckerl D, Cook PC, Jones LH, Finkelman FD, van Rooijen
N, et al. Local macrophage proliferation, rather than recruitment from the
blood, is a signature of TH2 inflammation. Science (2011) 332(6035):1284–8.
doi:10.1126/science.1204351
25. Wu D, Molofsky AB, Liang HE, Ricardo-Gonzalez RR, Jouihan HA, Bando JK,
et al. Eosinophils sustain adipose alternatively activated macrophages associated
with glucose homeostasis. Science (2011) 332(6026):243–7. doi:10.1126/science.
1201475
26. Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, et al. Tissueresident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity (2013) 38(4):792–804.
doi:10.1016/j.immuni.2013.04.004
27. Amano SU, Cohen JL, Vangala P, Tencerova M, Nicoloro SM, Yawe JC, et al.
Local proliferation of macrophages contributes to obesity-associated adipose
tissue inflammation. Cell Metab (2014) 19(1):162–71. doi:10.1016/j.cmet.2013.
11.017
28. Randolph GJ. Mechanisms that regulate macrophage burden in atherosclerosis.
Circ Res (2014) 114(11):1757–71. doi:10.1161/CIRCRESAHA.114.301174
29. Ramkhelawon B, Hennessy EJ, Menager M, Ray TD, Sheedy FJ, Hutchison S, et al.
Netrin-1 promotes adipose tissue macrophage retention and insulin resistance
in obesity. Nat Med (2014) 20(4):377–84. doi:10.1038/nm.3467
30. Stout RD, Jiang C, Matta B, Tietzel I, Watkins SK, Suttles J. Macrophages sequentially change their functional phenotype in response to changes in microenvironmental influences. J Immunol (2005) 175(1):342–9. doi:10.4049/jimmunol.
175.1.342
31. Li P, Lu M, Nguyen MT, Bae EJ, Chapman J, Feng D, et al. Functional heterogeneity of CD11c-positive adipose tissue macrophages in diet-induced obese mice.
J Biol Chem (2010) 285(20):15333–45. doi:10.1074/jbc.M110.100263
32. Shaul ME, Bennett G, Strissel KJ, Greenberg AS, Obin MS. Dynamic, M2like remodeling phenotypes of CD11c+ adipose tissue macrophages during high-fat diet – induced obesity in mice. Diabetes (2010) 59(5):1171–81.
doi:10.2337/db09-1402
33. Gundra UM, Girgis NM, Ruckerl D, Jenkins S, Ward LN, Kurtz ZD, et al. Alternatively activated macrophages derived from monocytes and tissue macrophages
are phenotypically and functionally distinct. Blood (2014) 123(20):e110–22.
doi:10.1182/blood-2013-08-520619
34. Haase J, Weyer U, Immig K, Kloting N, Bluher M, Eilers J, et al. Local proliferation of macrophages in adipose tissue during obesity-induced inflammation.
Diabetologia (2014) 57(3):562–71. doi:10.1007/s00125-013-3139-y
35. Wernstedt Asterholm I, Tao C, Morley TS, Wang QA, Delgado-Lopez F,
Wang ZV, et al. Adipocyte inflammation is essential for healthy adipose tissue
expansion and remodeling. Cell Metab (2014) 20(1):103–18. doi:10.1016/j.cmet.
2014.05.005
36. Kloting N, Fasshauer M, Dietrich A, Kovacs P, Schon MR, Kern M, et al.
Insulin-sensitive obesity. Am J Physiol Endocrinol Metab (2010) 299(3):E506–15.
doi:10.1152/ajpendo.00586.2009
37. Wentworth JM, Naselli G, Brown WA, Doyle L, Phipson B, Smyth GK, et al.
Pro-inflammatory CD11c+CD206+ adipose tissue macrophages are associated with insulin resistance in human obesity. Diabetes (2010) 59(7):1648–56.
doi:10.2337/db09-0287
38. Tordjman J, Poitou C, Hugol D, Bouillot JL, Basdevant A, Bedossa P, et al. Association between omental adipose tissue macrophages and liver histopathology
in morbid obesity: influence of glycemic status. J Hepatol (2009) 51(2):354–62.
doi:10.1016/j.jhep.2009.02.031
39. Zeyda M, Farmer D, Todoric J, Aszmann O, Speiser M, Gyori G, et al. Human
adipose tissue macrophages are of an anti-inflammatory phenotype but capable
of excessive pro-inflammatory mediator production. Int J Obes (Lond) (2007)
31(9):1420–8. doi:10.1038/sj.ijo.0803632
40. Fjeldborg K, Pedersen SB, Moller HJ, Christiansen T, Bennetzen M, Richelsen
B. Human adipose tissue macrophages are enhanced but changed to an antiinflammatory profile in obesity. J Immunol Res (2014) 2014:309548. doi:10.
1155/2014/309548

September 2014 | Volume 5 | Article 470 | 199

Kraakman et al.

41. Nguyen KD, Qiu Y, Cui X, Goh YP, Mwangi J, David T, et al. Alternatively activated macrophages produce catecholamines to sustain adaptive thermogenesis.
Nature (2011) 480(7375):104–8. doi:10.1038/nature10653
42. Qiu Y, Nguyen KD, Odegaard JI, Cui X, Tian X, Locksley RM, et al. Eosinophils
and type 2 cytokine signaling in macrophages orchestrate development of functional beige fat. Cell (2014) 157(6):1292–308. doi:10.1016/j.cell.2014.03.066
43. Xiao X, Gaffar I, Guo P, Wiersch J, Fischbach S, Peirish L, et al. M2 macrophages
promote beta-cell proliferation by up-regulation of SMAD7. Proc Natl Acad Sci
U S A (2014) 111(13):E1211–20. doi:10.1073/pnas.1321347111

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.

Frontiers in Immunology | Molecular Innate Immunity

Macrophage polarization in metabolic diseases

Received: 21 August 2014; accepted: 12 September 2014; published online: 26 September
2014.
Citation: Kraakman MJ, Murphy AJ, Jandeleit-Dahm K and Kammoun HL
(2014) Macrophage polarization in obesity and type 2 diabetes: weighing
down our understanding of macrophage function? Front. Immunol. 5:470. doi:
10.3389/fimmu.2014.00470
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Kraakman, Murphy, Jandeleit-Dahm and Kammoun. This is an
open-access article distributed under the terms of the Creative Commons Attribution
License (CC BY). The use, distribution or reproduction in other forums is permitted,
provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use,
distribution or reproduction is permitted which does not comply with these terms.

September 2014 | Volume 5 | Article 470 | 200

OPINION ARTICLE
published: 18 November 2014
doi: 10.3389/fimmu.2014.00594

Are “resting” microglia more “M2”?
Jonathan D. Cherry 1 , John A. Olschowka 2 and M. Kerry O’Banion 2 *
1

Department of Pathology and Laboratory Medicine, University of Rochester School of Medicine and Dentistry, Rochester, NY, USA
Department of Neurobiology and Anatomy, University of Rochester School of Medicine and Dentistry, Rochester, NY, USA
*Correspondence: kerry_obanion@urmc.rochester.edu
2

Edited by:
Charles Dudley Mills, BioMedical Consultants, USA
Reviewed by:
Geanncarlo Lugo-Villarino, Centre National de la Recherche Scientifique (CNRS), France
Charles Dudley Mills, BioMedical Consultants, USA
Keywords: microglia, M2, alternative activation, neuroinflammation, resting, activation

Over the last decade, the concept and
nomenclature of microglial phenotype
polarization has been carried over from the
peripheral macrophage literature. However, it is not entirely correct to view these
two cell types as overlapping. Microglia,
although related to macrophages, have
several differences and their own unique
repertoire of features. In particular,
microglia arise from a distinct early yolk
sac progenitor population and therefore
have different developmental origins than
macrophages (1). Furthermore, once in the
central nervous system (CNS), microglia
are maintained through local self-renewal
(2). Under normal conditions, there is no
infiltration of peripheral macrophages (3).
Microglia also maintain expression profiles
distinct from peripheral macrophages (4).
This demonstrates that microglia are not
simply macrophages that have migrated
into the brain; rather, they are a distinct
cell type.
These ideas have led to a reassessment of microglia activity, leading many
researchers to shift their thinking on
glial biology in general. One of the
more outdated concepts carried over from
macrophages is the idea that microglia
in the healthy brain exist in a “resting” state. Through a variety of stimuli
these “resting” cells can rapidly be “activated,” yielding microglia that are cytotoxic (5). These “activated” microglia were
described in and were thought to play a
major driving role in many neurodegenerative diseases (6). Thus, microglia “activation” took on a largely negative connotation. Taking cues from ongoing work in
peripheral macrophages, this initial concept of microglia “activation” being generally harmful gave way to more specific
ideas suggesting there was not just one

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kind of “activation.” Under the influence of
either pro- or anti-inflammatory cytokines,
microglia could be polarized into an
inflammatory or anti-inflammatory phenotype, designated classically and alternatively activated microglia, respectively (7).
Others have given different designations
not solely based on inflammation but from
inhibiting vs. healing functions, and label
them M1 and M2, respectively (8). It is
now becoming clear that the responses of
microglia, like macrophages, are heterogeneous: these responses can include proand anti-inflammatory signatures within
single cells and small, nearby populations and are driven by the local environment that can supply M1 and M2 polarizing cues simultaneously (9). Therefore,
microglial responses are much richer than
the dichotomous nomenclature suggests. It
is common to represent microglia polarization as a spectrum with each respective phenotype occupying the extremes
of the scale (10). However, it is unclear
whether the diverse functional responses
observed are a product of many different
kinds of microglia subsets or simply varying ratios of M1 and M2 microglia. While
there is much work that details this spectrum of activation, and much remains to be
investigated, an interesting but overlooked
area is the phenotype of the “resting”
microglial cell.
Classically, those who study microglial
function in a healthy, normal brain are not
the same groups that study microglia in an
“activated” or disease setting. However, it
appears that the microglia in each of these
settings may be more similar than originally thought. The notion that microglia
are truly “resting” has long been cast aside.
The advent of in vivo techniques, in particular 2-photon microscopy, has revealed

the constant surveillance and activity of
microglia, even in the absence of traditional activation signals (11). Therefore, it
might be incorrect to view microglia in the
normal, healthy CNS as a distinct population that can become “activated.” Rather, it
seems likely that even at baseline, microglia
are already on the activation spectrum. So
the question arises, where on the spectrum
are they?
Inflammation in the brain is typically
associated with harmful outcomes. Even
acute, low level inflammation can impair
synaptic function, leading to cognitive
dysfunction and behavioral abnormalities
(12). Moreover, neuroinflammation has
been recognized as a pathological hallmark
in most chronic neurodegenerative diseases
(13). This demonstrates the sensitivity of
the brain to inflammation and the importance of protecting the CNS from insult.
This protective role has generally been
assigned to M2 microglia. Many investigators have observed protective effects of
M2 cells; such as elevated neuronal survival
and process extension after treatment with
M2 conditioned media (14), or as reported
in numerous papers that detail the beneficial effect of direct treatment with M2
inducing agents (15). However, all of these
take place during pathology and do not
consider the normal protective function of
microglia. Thus, given the critical role of
basal microglia in maintaining homeostasis, an attractive hypothesis is that under
normal conditions, microglia are skewed
toward a protective, anti-inflammatory
phenotype. In fact many of the normal
functions of microglia are reminiscent of
M2 cells, although they are not as prominent as fully polarized cells. Even without stimulation, microglia are vital sources
of important, neurosupportive cytokines

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Cherry et al.

such as IGF-1 and BDNF (16–18). Neuroprotective cytokine secretion is generally
considered an M2 microglial function, but
we now know it also occurs in “resting”
microglia, albeit at lower levels. Indeed,
in models that lack proper M2 inducing
signals like IL-4−/− and SCID mice, cognitive impairment is observed, which was
attributed to decreased production of these
necessary neurotrophins (19, 20).
Furthermore, microglia in a basal state
share another important function with
fully polarized M2 microglia, namely, rapid
and efficient debris clearance (21). This
process, which is one of the quintessential defining functions of M2 polarized
cells, seems to be a default function for
microglia. Although the concept of phagocytosis is not unique to alternatively activated microglia, the speed and quality at
which this occurs differs between the phenotypes. In contrast to M1 polarized cells,
where a slower and less acidic phagosome
is beneficial for downstream immune functions such as antigen-presenting abilities, a
rapid, more acidic phagosome aids M2 cells
in quick, efficient removal of debris (22).
This speed and efficiency can be observed
in the highly dynamic process called synaptic pruning, which is characterized as rapid
elimination of developing synapses. In the
past several years, microglia have been
shown to be crucial to this process (23).
This concept is thought to be carried over
into the adult but at a less dramatic level
(24). The role of microglia in normal
synaptic maintenance is not fully understood, so it is difficult to directly attribute
baseline phagocytic function to an M2
related mechanism. However, due to the
speed and efficiency at which this process
occurs, we can speculate that this type of
phagocytosis shares more similarities with
M2 than M1 polarized microglia.
In addition to the neuroprotective and
functional similarities resting microglia
share with traditionally polarized M2
microglia, skewing to an M2 state can
be seen in the receptor profile resting
microglia express. For example, unlike
macrophages, microglia express very low
levels of MHCII (25). Only when microglia
polarize to an inflammatory phenotype do
they upregulate MHCII expression (26).
Contrary to MHCII, DC-SIGN has been
observed on microglia in the normal brain.
This c-type lectin receptor, which has been

Are “resting” microglia more “M2”?

implicated in promoting immune homeostasis, also maintains the immunosuppressive environment in the healthy brain
(27). By limiting particular surface receptors while expressing others, microglia
are biased toward a particular phenotype,
namely anti-inflammatory. However, it
appears that this M2 biased phenotype may
change with age. For still unknown reasons,
there is a loss of signals that keep microglia
anti-inflammatory. During normal aging,
a reduction in IL-4Rα is observed as well
as a decreased sensitivity to other antiinflammatory cytokines (28). This is mirrored by an increase in sensitivity to proinflammatory cytokines, suggesting a switch
in the basal phenotype of microglia (29).
A similar switch, termed priming, has been
observed after microglia were exposed to
inflammatory cytokines. Following the initial inflammatory insult, microglia appear
to return to their basal state. However,
with a second inflammatory stimulus
these “primed” microglia produce significantly more inflammatory cytokines than
unprimed microglia (30), suggesting that
their basal state is altered toward a more
M1 phenotype. Age-associated and priming switches may be involved in increased
susceptibility to neurodegenerative disorders (31).
Keeping microglia skewed toward a
non-inflamed state is critical for normal
homeostasis and specific control mechanisms exist that actively prevent microglia
from adopting an inflammatory profile. In
particular, neurons express several receptors and ligands that signal to their counterparts specifically localized on microglia.
CX3CR1, CD200, CD47, TREM2, and several other receptors have been identified that participate in constant cross-talk
between microglia and neurons (32). Interestingly, in genetic knockout mice missing
CX3CR1, impaired cognition was observed
(33). Two explanations appear likely. First,
the loss of direct inhibition resulted in
increased inflammation, which in turn,
caused cognitive dysfunction (33). Secondly, the loss of CX3CR1 resulted in
impaired phagocytic ability (23) and subsequent loss of proper synaptic pruning
during a critical developmental period,
as discussed previously. Although different, both of these explanations share the
idea that divergence from proper normal baseline function (namely M2 skewed

Frontiers in Immunology | Molecular Innate Immunity

functions) results in CNS pathology. Furthermore, this suggests that neurons, under
normal conditions, are active in controlling microglial polarization. This environmental control on phenotype skewing
poses an interesting question, is basal state
polarization present in other tissue specific macrophages? The answer is most
likely yes. However, it seems that other tissue macrophage cells such as peritoneal,
lung, or splenic red pulp macrophages all
exhibit greater diversity in their normal
gene expression and most likely, basal function when compared to microglia, even
though they a share common yolk sac
progenitor (1, 34). Therefore, it is hard to
directly compare between the cell types.
This demonstrates the large role the environment plays in skewing cells to the needs
of a specific tissue. In the CNS, an antiinflammatory state is most beneficial so
the environment favors slight M2 skewing. It would be interesting to characterize
potential skewing in other organs and how
that relates to normal resident macrophage
function.
The idea that microglia exist as a skewed
population in normal, non-pathological
tissue highlights this phenotype as an
innate characteristic of microglia. The classical view of macrophage biology is that
the adaptive arm of the immune system,
primarily T cells, controls phenotypes via
release of cytokines such as IFN-γ or IL4 (35, 36). Unfortunately, it is difficult to
prove innate or adaptive control in peripheral tissue due to the normal presence of T
cells. However, the healthy brain is largely
devoid of T cells, which accentuates the idea
that M2 phenotype skewing is a default,
innate function of microglia (37). Interestingly, similar ideas have been proposed for
macrophages (38).
In conclusion, to properly understand
and discuss microglia, we have to do
away with terms such as “activation” and
“resting.” By using these outdated concepts, we fail to acknowledge the complex plastic nature of microglia. In addition to not correctly representing the
non-pathological, normal functions of
microglia, the term “activation” is vague
and provides no specific information about
the many possible microglial phenotypes.
Microglia are always “active,” so the true
distinguishing feature is where they exist on
the phenotype spectrum. Healthy, normal

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Cherry et al.

brain microglia do not sit precisely in the
center between inflammatory and antiinflammatory cells. Rather, these sentinel
microglia are slightly shifted toward an
anti-inflammatory phenotype, which is
beneficial to brain homeostasis. The field of
microglial biology does not need to battle
over nomenclature for baseline microglia.
No single term can adequately encompass
all microglial functions at all times. But it is
important to recognize that these cells are
a plastic population that can dynamically
shift between a spectrum of phenotypes
and should not be boxed into fixed, rigid
“activation” states.

AUTHORS CONTRIBUTION
Jonathan D. Cherry researched the literature and drafted the manuscript. M. Kerry
O’Banion and John A. Olschowka critically reviewed and edited the work. All
the authors read and approved the final
manuscript.

ACKNOWLEDGMENTS
The authors thank Dr. Ania Majewska for
her editorial assistance as well as scientific insight on the subject matter. This
work was supported by the NIH/NIA grant
R01AG030149.

REFERENCES
1. Schulz C, Gomez Perdiguero E, Chorro L, SzaboRogers H, Cagnard N, Kierdorf K, et al. A lineage of
myeloid cells independent of Myb and hematopoietic stem cells. Science (2012) 336:86–90. doi:10.
1126/science.1219179
2. Elmore MR, Najafi AR, Koike MA, Dagher
NN, Spangenberg EE, Rice RA, et al. Colonystimulating factor 1 receptor signaling is necessary for microglia viability, unmasking a microglia
progenitor cell in the adult brain. Neuron (2014)
82:380–97. doi:10.1016/j.neuron.2014.02.040
3. Ajami B, Bennett JL, Krieger C, Tetzlaff W, Rossi
FM. Local self-renewal can sustain CNS microglia
maintenance and function throughout adult life.
Nat Neurosci (2007) 10:1538–43. doi:10.1038/
nn2014
4. Butovsky O, Jedrychowski MP, Moore CS, Cialic
R, Lanser AJ, Gabriely G, et al. Identification of a
unique TGF-beta-dependent molecular and functional signature in microglia. Nat Neurosci (2014)
17:131–43. doi:10.1038/nn.3599
5. Banati RB, Gehrmann J, Schubert P, Kreutzberg
GW. Cytotoxicity of microglia. Glia (1993)
7:111–8. doi:10.1002/glia.440070117
6. Streit WJ, Mrak RE, Griffin WS. Microglia and
neuroinflammation: a pathological perspective.
J Neuroinflammation (2004) 1:14. doi:10.1186/
1742-2094-1-14
7. Gordon S. Alternative activation of macrophages.
Nat Rev Immunol (2003) 3:23–35. doi:10.1038/
nri978

www.frontiersin.org

Are “resting” microglia more “M2”?

8. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill
AM. M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.
4049/jimmunol.164.12.6166
9. Martinez FO, Gordon S. The M1 and M2 paradigm
of macrophage activation: time for reassessment.
Prime Rep (2014) 6:13. doi:10.12703/P6-13
10. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation. Nat Rev Immunol
(2008) 8:958–69. doi:10.1038/nri2448
11. Nimmerjahn A, Kirchhoff F, Helmchen F. Resting microglial cells are highly dynamic surveillants of brain parenchyma in vivo. Science (2005)
308:1314–8. doi:10.1126/science.1110647
12. Rao JS, Kellom M, Kim HW, Rapoport SI, Reese
EA. Neuroinflammation and synaptic loss. Neurochem Res (2012) 37:903–10. doi:10.1007/s11064012-0708-2
13. Amor S, Puentes F, Baker D,Van Der Valk P. Inflammation in neurodegenerative diseases. Immunology (2010) 129:154–69. doi:10.1111/j.1365-2567.
2009.03225.x
14. Kigerl KA, Gensel JC, Ankeny DP, Alexander
JK, Donnelly DJ, Popovich PG. Identification of
two distinct macrophage subsets with divergent
effects causing either neurotoxicity or regeneration in the injured mouse spinal cord. J Neurosci
(2009) 29:13435–44. doi:10.1523/JNEUROSCI.
3257-09.2009
15. Cherry JD, Olschowka JA, O’Banion MK. Neuroinflammation and M2 microglia: the good, the
bad, and the inflamed. J Neuroinflammation (2014)
11:98. doi:10.1186/1742-2094-11-98
16. Parkhurst CN, Yang G, Ninan I, Savas JN,
Yates JR III, Lafaille JJ, et al. Microglia promote
learning-dependent synapse formation through
brain-derived neurotrophic factor. Cell (2013)
155:1596–609. doi:10.1016/j.cell.2013.11.030
17. Suh HS, Zhao ML, Derico L, Choi N, Lee
SC. Insulin-like growth factor 1 and 2 (IGF1,
IGF2) expression in human microglia: differential regulation by inflammatory mediators. J Neuroinflammation (2013) 10:37. doi:10.1186/17422094-10-37
18. Ueno M, Fujita Y, Tanaka T, Nakamura Y, Kikuta
J, Ishii M, et al. Layer V cortical neurons require
microglial support for survival during postnatal development. Nat Neurosci (2013) 16:543–51.
doi:10.1038/nn.3358
19. Derecki NC, Cardani AN, Yang CH, Quinnies KM,
Crihfield A, Lynch KR, et al. Regulation of learning and memory by meningeal immunity: a key
role for IL-4. J Exp Med (2010) 207:1067–80.
doi:10.1084/jem.20091419
20. Derecki NC, Quinnies KM, Kipnis J. Alternatively
activated myeloid (M2) cells enhance cognitive
function in immune compromised mice. Brain
Behav Immun (2011) 25:379–85. doi:10.1016/j.bbi.
2010.11.009
21. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism and
functions.
Immunity
(2010)
32:593–604.
doi:10.1016/j.immuni.2010.05.007
22. Canton J. Phagosome maturation in polarized
macrophages. J Leukoc Biol (2014) 96:729–38.
doi:10.1189/jlb.1MR0114-021R
23. Paolicelli RC, Bolasco G, Pagani F, Maggi L,
Scianni M, Panzanelli P, et al. Synaptic pruning

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

by microglia is necessary for normal brain
development. Science (2011) 333:1456–8. doi:10.
1126/science.1202529
Morris GP, Clark IA, Zinn R, Vissel B. Microglia:
a new frontier for synaptic plasticity, learning and
memory, and neurodegenerative disease research.
Neurobiol Learn Mem (2013) 105:40–53. doi:10.
1016/j.nlm.2013.07.002
de Haas AH, Boddeke HW, Biber K. Regionspecific expression of immunoregulatory proteins
on microglia in the healthy CNS. Glia (2008)
56:888–94. doi:10.1002/glia.20663
Loane DJ, Deighan BF, Clarke RM, Griffin RJ, Lynch AM, Lynch MA. Interleukin-4
mediates the neuroprotective effects of rosiglitazone in the aged brain. Neurobiol Aging
(2009) 30:920–31. doi:10.1016/j.neurobiolaging.
2007.09.001
Garcia-Vallejo JJ, Ilarregui JM, Kalay H, Chamorro
S, Koning N, Unger WW, et al. CNS myelin induces
regulatory functions of DC-SIGN-expressing,
antigen-presenting cells via cognate interaction
with MOG. J Exp Med (2014) 211:1465–83. doi:
10.1084/jem.20122192
Fenn AM, Henry CJ, Huang Y, Dugan A, Godbout
JP. Lipopolysaccharide-induced interleukin (IL)-4
receptor-alpha expression and corresponding sensitivity to the M2 promoting effects of IL-4 are
impaired in microglia of aged mice. Brain Behav
Immun (2012) 26:766–77. doi:10.1016/j.bbi.2011.
10.003
Lee DC, Ruiz CR, Lebson L, Selenica ML, Rizer
J, Hunt JB Jr, et al. Aging enhances classical
activation but mitigates alternative activation
in the central nervous system. Neurobiol Aging
(2013) 34:1610–20. doi:10.1016/j.neurobiolaging.
2012.12.014
Teeling JL, Perry VH. Systemic infection and
inflammation in acute CNS injury and chronic
neurodegeneration: underlying mechanisms. Neuroscience (2009) 158:1062–73. doi:10.1016/j.
neuroscience.2008.07.031
Perry VH. The influence of systemic inflammation on inflammation in the brain: implications
for chronic neurodegenerative disease. Brain Behav
Immun (2004) 18:407–13. doi:10.1016/j.bbi.2004.
01.004
Kierdorf K, Prinz M. Factors regulating microglia
activation. Front Cell Neurosci (2013) 7:44. doi:10.
3389/fncel.2013.00044
Rogers JT, Morganti JM, Bachstetter AD, Hudson CE, Peters MM, Grimmig BA, et al. CX3CR1
deficiency leads to impairment of hippocampal cognitive function and synaptic plasticity. J Neurosci (2011) 31:16241–50. doi:10.1523/
JNEUROSCI.3667-11.2011
Gautier EL, Shay T, Miller J, Greter M, Jakubzick C, Ivanov S, et al. Gene-expression profiles and
transcriptional regulatory pathways that underlie the identity and diversity of mouse tissue
macrophages. Nat Immunol (2012) 13:1118–28.
doi:10.1038/ni.2419
Stein M, Keshav S, Harris N, Gordon S. Interleukin 4 potently enhances murine macrophage
mannose receptor activity: a marker of alternative immunologic macrophage activation.
J Exp Med (1992) 176:287–92. doi:10.1084/jem.
176.1.287

November 2014 | Volume 5 | Article 594 | 203

Cherry et al.

36. Dalton DK, Pitts-Meek S, Keshav S, Figari IS,
Bradley A, Stewart TA. Multiple defects of immune
cell function in mice with disrupted interferongamma genes. Science (1993) 259:1739–42. doi:10.
1126/science.8456300
37. Ransohoff RM, Brown MA. Innate immunity in
the central nervous system. J Clin Invest (2012)
122:1164–71. doi:10.1172/JCI58644
38. Mills CD, Ley K. M1 and M2 macrophages: the
chicken and the egg of immunity. J Innate Immunity (2014) 6:716–26. doi:10.1159/000364945

Are “resting” microglia more “M2”?

Conflict of Interest Statement: The authors declare
that the research was conducted in the absence of any
commercial or financial relationships that could be
construed as a potential conflict of interest.

Received: 06 August 2014; accepted: 05 November 2014;
published online: 18 November 2014.
Citation: Cherry JD, Olschowka JA and O’Banion
MK (2014) Are “resting” microglia more “M2”? Front.
Immunol. 5:594. doi: 10.3389/fimmu.2014.00594

Frontiers in Immunology | Molecular Innate Immunity

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology.
Copyright © 2014 Cherry, Olschowka and O’Banion.
This is an open-access article distributed under the terms
of the Creative Commons Attribution License (CC BY).
The use, distribution or reproduction in other forums is
permitted, provided the original author(s) or licensor are
credited and that the original publication in this journal
is cited, in accordance with accepted academic practice.
No use, distribution or reproduction is permitted which
does not comply with these terms.

November 2014 | Volume 5 | Article 594 | 204

REVIEW ARTICLE
published: 24 November 2014
doi: 10.3389/fimmu.2014.00606

M1/M2 macrophage polarity in normal and complicated
pregnancy
Mary B. Brown, Maria von Chamier , Ayman B. Allam and Leticia Reyes*
D. H. Barron Reproductive and Perinatal Biology Research Program, Department of Infectious Disease and Pathology, College of Veterinary Medicine, University of
Florida, Gainesville, FL, USA

Edited by:
Laurel L. Lenz, University of Colorado
School of Medicine, USA
Reviewed by:
Andrew Tasman Hutchinson,
University of Technology Sydney,
Australia
Devendra K. Agrawal, Creighton
University School of Medicine, USA
*Correspondence:
Leticia Reyes, D. H. Barron
Reproductive and Perinatal Biology
Research Program, Department of
Infectious Disease and Pathology,
College of Veterinary Medicine,
University of Florida, PO Box 110880,
Gainesville, FL 32611-0880, USA
e-mail: lreyes@ufl.edu

Tissue macrophages play an important role in all stages of pregnancy, including uterine stromal remodeling (decidualization) before embryo implantation, parturition, and post-partum
uterine involution. The activation state and function of utero-placental macrophages are
largely dependent on the local tissue microenvironment. Thus, macrophages are involved
in a variety of activities such as regulation of immune cell activities, placental cell invasion,
angiogenesis, and tissue remodeling. Disruption of the uterine microenvironment, particularly during the early stages of pregnancy (decidualization, implantation, and placentation)
can have profound effects on macrophage activity and subsequently impact pregnancy
outcome. In this review, we will provide an overview of the temporal and spatial regulation
of utero-placental macrophage activation during normal pregnancy in human beings and
rodents with a focus on more recent findings. We will also discuss the role of M1/M2
dysregulation within the intrauterine environment during adverse pregnancy outcomes.
Keywords: macrophages, M1/M2, normal pregnancy, complicated pregnancy, uterine atherosis, spiral artery,
chorioamnionitis, spontaneous abortion

INTRODUCTION
Macrophages within the maternal–fetal compartment have been
a research focus for over 30 years. Both maternal and fetal derived
macrophages play an important role in all stages of pregnancy.
These cells support a variety of processes essential for successful
pregnancy such as remodeling of the uterine connective tissues and
blood vessels, regulation of trophoblast (fetal cell) implantation,
immune-tolerance toward fetal antigens, immunomodulation of
neighboring leukocytes, and initiation of parturition (1–5). All of
these functions are manifestations of macrophage polarity or state
of activation, which at the most basic level has been referred to
as M1 and M2. M1 macrophages display the capacity to present
antigen, produce IL-12, IL-23, reactive oxygen species (6, 7), and
skew T cell responses toward a TH1 or cell mediated immune
response (8). In contrast, M2 macrophages participate in tissue
remodeling, have immunosuppressive qualities, and promote TH2
or antibody mediated immune responses (6). In essence, M1/M2
activities arise from arginine metabolism via two enzymatic pathways (iNOS and arginase) that down regulate each other (7, 9).
The factors that influence which pathway dominates are based on
the surrounding signals that the macrophage receives as well as the
available arginine pool (7, 10). Thus, macrophage function, which
is a manifestation of the cell’s polarization or activation state, is
ultimately decided by the surrounding milieu.
The maternal–fetal interface is a unique environment in that
it comprises three distinct compartments: the placenta (fetal
origin), and maternal origin (endometrium or decidua, and
myometrium), which are infiltrated with fetal cells. Furthermore,
these compartments undergo dramatic changes in architecture
and leukocyte composition as gestation progresses. These changes
are necessary for placental development and tolerance of foreign

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fetal antigens. Macrophages are present in all compartments
and during all stages of pregnancy (implantation, placentation,
fetal growth, and parturition) (11). Successful pregnancy requires
that the macrophage activation state remain regulated throughout pregnancy. Indeed, inappropriate macrophage polarization
within the maternal–fetal compartments is associated with spontaneous abortion or miscarriage (12), inadequate remodeling of
the uterine vessels during placentation (12–14), and intrauterine
parasitic infections (15). In this review, we provide an overview
of M1/M2 dynamics relevant to the maternal–fetal interface,
the temporal and spatial changes in macrophage M1/M2 within
the maternal and fetal compartments during normal pregnancy,
and imbalanced M1/M2 dynamics associated with complicated
pregnancies.

M1 AND THE MANY SHADES OF M2
Macrophages display divergent phenotypes that were originally
described as M1or M2 polarity (7). M1 refers to the classically
activated macrophage whereby the cell displays the capacity to
present antigen, produce IL-12, IL-23, and reactive oxygen species
(6, 7). M1 polarized macrophages are more effective at antimicrobial killing and skewing T cell responses toward a TH1 or cell
mediated immune response (8). External stimuli known to promote M1 polarization include toll-like receptor (TLR4) agonists
such lipopolysaccharide (LPS) from E. coli, IFN-γ, TNF-α, and
GM-CSF (7, 16). In contrast, M2 polarized cells (alternatively activated cells) display abundant expression of mannose and scavenger
receptors and produce IL-10 and TGF-β (6). M2 macrophages participate in tissue remodeling, have immunosuppressive qualities,
and promote TH2 or antibody mediated immune responses (6).
Stimuli that induce M2 polarization include IL-4, IL-10, IL-13,

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M1/M2 in pregnancy

IL-33, TGF-β, and G-CSF (7, 16–19). Because of the broad range of
activities carried out by M2 macrophages, Mantovani et al. (6) later
proposed that these cells be further divided into M2a (induced by
IL-4 and IL-13), M2b (macrophages exposed to immune complexes or toll-like receptor agonists), and M2c (induced by IL-10
and glucocorticoid hormones). These subcategories were based
on both the type of agonists responsible for triggering their differentiation and the distinct functional profile induced by these
agonists. For example, M2a cells display the alternatively activated
phenotype typically attributed to M2 cells. M2b confers a Type
II phenotype, because it promotes TH2 responses and produces
both pro- and anti-inflammatory cytokines (TNF-α, IL-1, IL-6,
IL-10high , and IL-12low ) (6). M2c confers an M2 macrophage that
produces IL-10, TGF-β, and extracellular matrix components (6).
At the most fundamental level, M1/M2 polarity is driven by
arginine metabolism and available arginine within the extracellular milieu. In essence, M1 is a product of iNOS pathway, which
drives the conversion of arginine to citrulline and NO, whereas
M2 is a product of the arginase pathway in which arginine is
hydrolyzed into ornithine and urea (7, 20). iNOS and arginase
pathways are antagonistic, not only do they compete for arginine

but also arginase directly inhibits NO synthase 2 (NOS2) (10).
Although M2 is defined by the arginase pathway (7, 9), in Mantovani et al.’s designation, M2b develops independent of arginase
activity, while M2a and M2c still require arginase activity (6).
A distinctive feature of the iNOS pathway is that its activation
requires extracellular arginine, even if an adequate level of intracellular arginine is already present (21). Without sufficient arginine
iNOS is blocked at the translational level and NO is not produced
even in the presence of IFN-γ or LPS. Under these conditions, the
macrophage does not acquire an M1 phenotype, but still produces
TNF-α in response to IFN-γ or LPS (21). This condition bears
a resemblance to M2b macrophages (6). This dependency upon
extracellular arginine reserves may be particularly important during pregnancy since placental tissues are rich in arginase I and
II (22), and therefore, placental extracellular arginine levels may
actually be too low for induction of M1 macrophages.
Several studies have attempted to characterize the functional state of macrophages at the maternal–fetal interface during different stages of a healthy pregnancy (Table 1). These
include transcriptome analysis (23, 24), methylation profiling (25), characterization of cell surface markers (26–28), and

Table 1 | Summary of recent studies that characterized M1/M2 status of macrophages within the maternal–fetal interface during healthy
pregnancy.
First

Year

author

Gestation

Methods

Results

Reference

at sampling
(weeks)

Laskarin

Repnik

2005

2008

6–8

Immunohistochemistry,

Confirmed that decidual and placental Mφ express mannose receptor

Term

costaining for CD14 decidual

in vivo

and placental macrophages

CD14+ decidual Mφ surround endometrial glands in early pregnancy

(Mφ) and mannose receptor

50% Of CD14+ Mφ in term decidua and villi (Hofbauer) express

(M2 biomarker)

mannose receptor

9–18

Flow cytometry of decidual

Decidual Mφ from early/mid gestation have higher CD80, CD86, and

19–23

Mφ

HLA-DR expression than term Mφ

Term
Gustafsson

Houser

2008

2011

7–11

6–12

Kim

2011

2012

7–12

Term

(26)

Mφ in decidua basalis express CD105, DC-SIGN, and mannose receptor
RNA Microarray analysis of

The majority of genes upregulated in decidual Mφ were functionally

CD14+ Mφ isolated from

related to immunomodulation and tissue remodeling consistent with an

decidua and peripheral blood

activated M2 phenotype

Flow cytometry and RNA

Identified 2 decidual Mφ subsets: CD11cHI and CD11cLO

microarray of

Svensson

(29)

CD14+

Mφ

(24)

CD11cLO decidual Mφ express genes associated with extracellular

subsets isolated from

matrix formation, muscle regulation, and tissue growth

decidua basalis and

CD11cHI decidual Mφ express genes associated with lipid metabolism

peripheral blood

and inflammation

Phenotypic characterization

Identified 2 decidual Mφ subsets: ICAM-3HI and ICAM-3LO

of decidual Mφ by flow

ICAM-3 expression on decidual Mφ positively correlated with CD11c

cytometry

expression as reported by Houser and colleagues

RNA microarray on in vitro

M-CSF/IL-10 in vitro differentiated Mφ and decidual Mφ showed similar

differentiated Mφ cultures

cytokine secretion patterns (↑IL-10, IL-6, TNF, and CCL4)

Methylome analysis of

Both decidual Mφ and Hofbauer cells exhibited hypermethylation of

decidual Mφ, Hofbauer

genes encoding classical Mφ activation

cells, neonatal, and maternal

Both decidual Mφ and Hofbauer cells exhibited hypomethylation of

blood monocytes (Mo)

genes encoding alternative Mφ macrophage activation

Frontiers in Immunology | Molecular Innate Immunity

(23)

(27)

(25)

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Brown et al.

immunohistochemistry (29). When viewed in the context of
M1 and M2, the results from these studies suggest that M2
macrophages or M2 subgroups are the predominant phenotype
in the decidua with a smaller subset of macrophages bearing some
characteristics of M1 or M2b (24). One drawback is that most of
these studies analyzed macrophages out of their in situ context so
the local factors responsible for triggering the differentiation of
these macrophage subsets are still unknown.
The role of phosphatidylserine receptor Tim-3 in macrophages
from the maternal–fetal interface has gained recent attention (30).
Tim-3 is recognized as a regulator of pro- and anti-inflammatory
innate immune responses (31), and Tim-3 expression is enhanced
in M2 immunoregulatory macrophages (32, 33). Therefore, Tim-3
may play an important role in maintaining macrophage mediated immune tolerance at the maternal–fetal interface (31).
Indeed, Tim-3 facilitates phagocytic activity of apoptotic bodies
at the maternal–fetal interface, and blockade of Tim-3 on uterine
macrophages led to increased production of IFN-γ and TNF-α by
inflammatory granulocytes with subsequent rejection of the fetus
(30). To date, it is unknown if loss of Tim-3 or dysregulation of
Tim-3 occurs in adverse pregnancy outcome.
Hofbauer cells are macrophages that reside within the mesenchymal stroma of the chorionic villi (34), which are thought
to originate from fetal hematopoietic cells (35). In human placenta, Hofbauer cells have been detected by 4 weeks gestation, and
their numbers increase during the first trimester and then decline
by term (36). Based on their hypermethylation patterns, Hofbauer
cells display a commitment to M2 in healthy pregnancy (25). Other
phenotypic features that suggest these cells are pro-M2 include
constitutive expression of DC-sign (CD209), high levels of CD163,
CD68, CD45, hyaluronan receptor LYVE-1, and HLA antigens A,
B, and C, IL-10, and TGF-β (37–39). Hofbauer cells may have a role
in placental angiogenesis, remodeling of the extracellular matrix,
and modulation of inflammation, which are M2 characteristics
(39–41). Although not a particular feature of macrophage function, it has been suggested that Hofbauer cells participate in the
regulation of stromal fluid balance, ion exchange, and transfer of
serum proteins to the vascular system (36).

ANATOMY OF THE MATERNAL–FETAL INTERFACE IN
RELATION TO MACROPHAGES
The various compartments of the maternal–fetal interface (placenta, decidua, and myometrium) are anatomically distinguished
by the composition of the connective tissue and the leukocyte populations present in each compartment. For example, macrophages
residing in the myometrium may be receiving signals from T-cells,
dendritic cells, and myocytes, whereas macrophages within the
decidua receive signals from decidualized stromal cells, uterine NK
cells, T-cells, and trophoblasts. In addition, decidual macrophages
respond to immunoregulatory compounds such as HLA-G, TGFβ, and vasoactive intestinal peptide (VIP) that are produced by
adjacent trophoblasts (40–42).
The mammalian uterus has two distinct tissue layers: the
endometrium and myometrium. Macrophages are found interspersed in both layers of the uterus. The luminal surface of
the endometrium consists of simple cuboidal epithelium that is
supported by underlying connective tissue. The endometrium

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M1/M2 in pregnancy

contains a complex vascular supply and a diverse population of
leukocytes that changes with stage of the menstrual cycle and stage
of pregnancy (43). The myometrium is composed of interlacing
bundles of smooth muscle. In human beings, the myometrium
exhibits minimal change during pregnancy (44). In rodents, however, the myometrium below the site of placental attachment
expands and develops into the metrial triangle (44). Prior to
embryo implantation, the endometrium begins to undergo structural changes referred to as decidualization, which is initiated by
increased progesterone levels (44, 45). The process of decidualization in human beings stabilizes by 12 weeks gestation (46). The
physical changes that occur during decidualization include proliferation of luminal epithelium, development of secretory glandules with large apical protrusions (pinopodes) and microvilli,
and transformation of fibroblast-like endometrial stromal cells
into larger rounded cells (47, 48). In addition, endometrial cells
produce a variety of compounds that attract leukocytes to the
decidualized tissue such as prolactin, colony stimulating factor
1 (CSF-1), macrophage inhibitory factor (MIF), IL-15, insulin
growth factor binding protein-1 (IGFBP-1), and cyclooxygenase-2
(COX-2), and cell adhesion molecules (ICAM-1, VACM-1, LFA3, H-CAM) (41, 45, 48, 49). Both CSF-1and MIF influence the
recruitment of macrophages into the decidua (11, 43, 50, 51).
Following embryo implantation, the decidua differentiates into
two distinct regions: the decidua basalis, which refers to the portion of the uterus attached to the placenta, and decidua parietalis,
which refers to the rest of the endometrium lining the main cavity
of the pregnant uterus. In human beings, macrophages within the
decidua basalis coexist with trophoblasts, CD56bright /CD16low NK
cells, and T-cells, 25% of which are CD4+ /CD25+ /FOXP3+ regulatory T-cells (Tregs) (11, 44). Within the decidua basalis, CD14+
macrophages and NK cells are often observed in aggregates in
association with spiral arteries (11, 52). Decidual macrophages are
also observed in close association with trophoblasts (40). Factors
released by neighboring trophoblasts may induce decidual trophoblasts to display cell surface expression of CD14 and CD16,
exhibit increased phagocytic capacity, and secrete more IL-1β,
IL-10, and IP-10 (53).
The microenvironment within the myometrium slightly differs from the decidual basalis in that macrophages and T-cells
are the predominant leukocyte populations, and there are few
CD56bright /CD16low NK cells (54). In rats and mice, macrophages
are more predominant in the myometrium than in the decidua,
which has been attributed to higher CSF-1 concentrations within
this compartment (55). Trophoblasts are also present within the
inner third of the myometrium in both human beings and rats, but
not in mice (14, 56). Macrophages recruited into the myometrium
are important for the normal induction of labor and post-partum
uterine remodeling (5, 57).
Placentation in human beings, non-human primates, and
rodents is described as hemochorial because there are no barriers
between trophoblast cells and maternal blood (58). Hemochorial placentation requires extensive tissue remodeling in which
the endometrium that becomes the decidua basalis is obliterated
and the uterine arteries are transformed from high resistance,
low-flow arteries to low resistance, and high-flow vessels (14). During remodeling of spiral arteries, the endothelium and smooth

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Brown et al.

muscle cells are replaced by invading extravillous trophoblasts
and fibrinoid deposits (59). This complex process depends on
a coordinated crosstalk between decidual stromal cells, uterine
NK cells, and CD14+ /CD68+ macrophages (59–62). Under optimal circumstances, the transient invasion of uterine NK cells
and CD14+ /CD68+ macrophages into the spiral arteries pave the
way for the second stage of remodeling that involves invasion of
trophoblasts into the vessels (52, 59).

M1 OR M2 IN NORMAL PREGNANCY, IT IS IN THE TIMING
It has been recently proposed that pregnancy is actually an
active and highly regulated immunologic process (63). Successful implantation requires a transient inflammatory phase that is
initiated by cytokines and prostaglandins within seminal fluid
(64). During the peri-implantation period, the polarization pattern of decidual macrophages is skewed toward M1 (65). However,
as trophoblasts establish attachment into the endometrial lining
and invade the uterine stroma, decidual macrophages begin to
transition to a mixed M1/M2 profile (65). This mixed population may represent M1/M2 or possibly a blend of M2 subtypes
(pro-inflammatory M2b mixed with anti-inflammatory M2a and
M2c). This mixed polarization pattern continues through the first
trimester and the early phase of the second trimester of pregnancy
when the uterine vasculature is undergoing extensive remodeling
in order to establish an adequate placental–fetal blood supply (63).
After placental development is complete, the decidua shifts toward
a pro-M2 environment, which prevents rejection of the fetus and
allows fetal growth until parturition. Parturition, which is another
inflammatory event, is preceded by an influx of macrophages into
the myometrium and decidua (3–5). This inflammatory process
promotes the contraction of the uterus, expulsion of the baby,
ejection of the placenta, and uterine involution (57).
A2V, which is an isoform of the alpha subunit of vacuolar
ATPase (V-ATPase), is a key immune regulator important for
implantation (66). At least in normal murine pregnancy, A2V is
expressed in sperm, embryo, and endometrium (65), and loss of
a2v results in reproductive failure (67). In the reproductive tract,
a2V promotes a transient pro-inflammatory effect followed by a
balanced immune response that facilitates embryo implantation
without rejection. During the pre-implantation period, cleavage
of a2V in semen fluid releases a soluble N-terminus portion of the
protein (a2NTD). Seminal a2NTD has pro-inflammatory effects
such as upregulation of leukemia inhibitory factor (LIF), IL-1β,
TNF-α, and MCP-1 gene expression in the uterus, creating a transient pro-M1 effect. The developing embryo continues to produce
a2V, which continues to stimulate uterine MCP-1. This in turn
attracts new macrophages into the endometrium leading to a
more balanced M1/M2 ratio and protection against rejection of the
fetus. In the BALB/c mouse, these studies clearly demonstrate that
a2V mediated induction of MCP-1 is important for proper regulation of M1/M2 during early pregnancy (65, 67). Additional factors
present within the decidualized tissue that influence macrophage
polarization of invading macrophages also contribute to tolerance.
As pregnancy progresses, the M1/M2 ratio decreases (23, 27)
and the population of decidual macrophages become more heterogeneous. This, in part, is caused by the uneven spatial distribution of neighboring leukocytes and trophoblasts within the

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M1/M2 in pregnancy

decidua, which provide different cues to resident macrophages.
Events that account for these changes include the contribution of immunoregulatory molecules from trophoblasts such
as HLA-G (68, 69), TGF-β (40), and VIP (41). These compounds have direct effects on macrophages or on Tregs, which
express pro-M2 factors such as IDO and IL-10 (1). In contrast, CD56bright /CD16low NK cells, which are intimately associated with decidual macrophages invading the spiral arteries,
produce pro-M1 IFN-γ (70) and TNF-α (11). Houser et al. identified two distinct macrophage subsets in first trimester decidua
based on the degree of CD11c expression (24). Regardless of
the degree of Cd11c expression, these cells uniformly expressed
CD68 and CD14. CD11cHI macrophages expressed less phagocytic receptors such as CD209 (DC-SIGN) and CD206 (mannose receptor) compared to CD11cLO macrophages. CD11cHI and
CD11cLO macrophages also had different transcriptional profiles.
For example, CD11cHI macrophages express genes involved in
invasion, mobility, inflammatory processes including lipid metabolism, and anti-apoptotic effects, whereas CD11cLO macrophages
express genes that regulate growth and development, as well
extracellular communication including networking. CD11cLO
macrophages may be the subpopulation of cells that actively suppress CD56bright /CD16low NK cell mediated killing of invading
trophoblasts (2). On the other hand, the transcriptional profile of
CD11cHI macrophages suggests that these cells participate in the
remodeling of the uterine arteries that occurs during this stage of
pregnancy.
During the growth phase of pregnancy, the predominant
immunological feature is the induction of an anti-inflammatory
state. It is presumed that the immunosuppressive qualities
observed in decidual macrophages studied from first trimester
pregnancies continue to be present as pregnancy advances (71).
This notion is supported by a several studies that evaluated
macrophages isolated from term decidua. First, the methylation
pattern of these cells show silencing of genes that encode M1
markers such as TLR9, IL1B, IL12RB2, CD48, and FGR, whereas
genes associated with M2 polarity such as CCL13, CCL14, A2M,
HNMT, and IL10 were hypomethylated (25). Second, decidual
macrophages express M2 polarity characteristics such as reduced
expression of co-stimulatory molecule CD86 with spontaneous
production of high levels of IL-10 and IDO (72). Third, decidual macrophages express C-type lectin receptors such as CD206
throughout pregnancy (73), which is more in line with an M2
phenotype (71).
Normal parturition is an inflammatory process characterized
by an enhanced expression of IL-1β, IL-6, and IL-8 with an accumulation of leukocytes in the cervix, fetal membranes, decidua,
and myometrium (74). Macrophages are one of the predominant
cell types in the decidua and myometrium during normal parturition. CD68+ macrophages begin to invade the decidua a few days
before parturition (3, 4), which coincides with a global increase
in the production of chemokines such as CCL2 (MCP-1), CCL4
(MIP-1β), CCL5 (Rantes), CXCL8 (IL-8), and CXCL10 (IP-10)
within the decidua (75). Immediately before parturition, a wave of
macrophages invades the myometrium (4), resulting in an increase
in pro and anti-inflammatory cytokines such as IL-1β, IL-6, IL-12,
and IL-10. To date, the polarity profile of macrophages invading

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Brown et al.

the maternal–fetal interface before or during labor has not been
determined.
Although the density of Hofbauer cells within the villous mesenchyme fluctuates with gestational age (36), the limited characterization of these cells suggests that they display a commitment
to M2 related functions. DNA methylation profiling of Hofbauer
cells obtained from term placentas show that pro-M1 genes such as
TLR9, IL1B, IL12RB2, CD48, and FGR are silenced (25). In contrast, pro-M2 genes such as CCL2, CCL13, CCL14, CD209, and
A2M are hypomethylated, and thus, available for expression (25).
Hofbauer cells constitutively express anti-inflammatory cytokines
such as IL-10 and TGF-β1 (38), which may in part be related
to the immune suppressive effects of neighboring mesenchymal
stem cells that have the capacity to shift macrophages from an
M1 phenotype to M2 (76). It is also likely that arginase I and
II activity from surrounding villous trophoblasts (22) reduces
extracellular arginine pools within the villous mesenchyme that
would be required for translation of iNOS and differentiation
into M1.

M1/M2 IMBALANCES DURING EARLY PREGNANCY
It is noteworthy that some obstetric complications associated with
decidual M1/M2 imbalances occur during the early inflammatory phase of pregnancy. Under these circumstances, it is likely
that the surrounding environment supports M1 polarization over
M2. Some examples include spontaneous abortions and disorders
involving inadequate remodeling of the uterine arteries.
Spontaneous abortions that occur within 12 weeks gestation are
associated with an increased influx of macrophages in the decidual stroma (12, 77). In one study, decidual macrophages from
spontaneous abortions had increased Fas-L expression, which
coincided with an increased rate of trophoblast apoptosis (12).
It was proposed that the increased FasL-expressing population of
macrophages reflected M1 activation (12); however, this is yet to be
proven. Jaiswal et al. recently demonstrated that a shift toward M1
polarization at the maternal–fetal interface enhances abortion in
CBA × DBA/2 mouse matings treated with LPS (67). The underlying mechanism involved a significant decrease in placental a2V
coupled with a decrease in uterine MCP-1 expression.
Inadequate remodeling of the uterine spiral arteries is associated with a spectrum of obstetric complications such as preeclampsia, intrauterine growth restriction, pre-term birth, preterm premature rupture of membranes (PPROM), late sporadic
miscarriage, and premature separation of the placenta from the
uterus (78). During optimal spiral artery remodeling, there is a
transient influx of NK cells and activated CD68+ macrophages
into the vessel wall. This leukocytic influx, which is described
as stage 1 remodeling, is thought to initiate the disruption of
the organized smooth muscle layer and endothelium needed
for subsequent trophoblast invasion into the vessel (59). ProM2 stromal macrophages in the outer periphery of the vessel
assist in controlling inflammation by phagocytizing potentially
pro-inflammatory cellular debris and secreting anti-inflammatory
TGF-β1 (40). Excessive M1 or M2b activity during the early phase
of remodeling can prevent the resolution of this inflammatory
phase, thus, disrupting trophoblast invasion into the arterial wall
and preventing complete remodeling of the vessel (52, 59). Indeed,

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M1/M2 in pregnancy

reduced trophoblast invasion into utero-placental spiral arteries is associated with an excess of activated macrophages in and
around these arteries (79, 80). Macrophages activated by exogenous TNF-α, tryptophan depletion (which reduces production
of IDO), GM-CSF, or M-CSF promote apoptosis of extravillous
trophoblasts in vitro (80, 81).

M1/M2 IMBALANCES DURING LATE PREGNANCY
Decidual inflammation is also presumed to produce M1 or M2b
excess in the pathogenesis of uterine atherosis. Uterine atherosis
is a late pregnancy lesion of the decidua that is characterized by
the accumulation of lipid filled CD68+ foamy macrophages in
the subendothelial layer of uterine spiral arteries (14, 82). This
lesion is also associated with increased local TNF-α (83). In rare
instances, acute atherosis has also been observed in the myometrial
segments of the spiral arteries (78, 82). During early pregnancy,
acute atherosis can also be seen in cases of defective spiral artery
remodeling (78). In this instance, the lesions are present in the
lower section of the arteries where they feed into the intervillous
space (82).
Staff and Redman proposed that acute atherosis is an end result
of various inflammatory pathways triggered by immunologic,
genetic, and/or hemodynamic influences (sequela to impaired spiral arterial remodeling and perturbed laminar blood flow) that
may be working singly or in combination (82). This hypothesis is based on the observation that acute atherosis is found in
a wide range of pregnancy complications such as pre-eclampsia,
fetal growth restriction, and certain autoimmune diseases (systemic lupus erythematosus and antiphospholipid syndrome). It
has also been suggested that infectious organisms implicated in
promoting cardiovascular disease such as Chlamydia pneumoniae may play a role in initiating or activating uterine atherosis
(84). Recent work in our laboratory suggests that the periodontal pathogen, Porphyromonas gingivalis, which is implicated in
promoting cardiovascular disease (85), may indeed have such an
effect.

M1/M2 IMBALANCES ASSOCIATED WITH INTRAUTERINE
INFECTION
Porphyromonas gingivalis is a common periodontal pathogen of
human beings that is also implicated in low-birth weight, fetal
growth restriction, pre-eclampsia, and spontaneous pre-term birth
(86–93). In prototypical M2 Balb/c mice, intrauterine infection
with P. gingivalis induces pro-M1 or M2b inflammatory responses
such as increased TNF-α and IFN-γ with suppression of IL10, and fetal growth restriction (94, 95). Using a rat model of
intrauterine infection, we observed that the local presence of P.
gingivalis within the uterus produced lesions suggestive of an
M1 > M2 or M2b > M2a, M2c imbalance. Namely, infected dams
exhibited acute arteritis within the endometrium and metrial triangle characterized by perivascular necrosis, hyaline degeneration
with varying degrees of thrombosis (96, 97). Moreover, the spiral arteries had increased densities of CD68+ macrophages, with
increased stromal TNF-α and a concomitant decrease in extravillous trophoblast invasion into the placental bed. Since this was
an acute study (4 days duration), we did not observe lipid filled
macrophages surrounding affected spiral arteries. Nevertheless,

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these results provide a compelling argument that certain bacterial
infections may promote M1/M2 imbalances that in the placental
bed have a negative impact on spiral artery remodeling.
To date, there is no evidence that pro-M1 activity is actually
involved in the pathogenesis of chorioamnionitis and pre-term
delivery. Rather, acute chorioamnionitis may actually promote M2
polarization. Decidual macrophages in gestation day 16 FVB/NJ
mice sustain an M2 polarity phenotype even after LPS treatment (98). C57BL/6 mice, which are resistant to chorioamnionitis
during intrauterine infection with Ureaplasma parvum (99), do
not develop pro-M1 responses, instead C57BL/6 mice exhibit a
pro-M2 profile within the placenta and decidua (100). This is
particularly intriguing since C57BL/6 mice are known for their
prototypical M1 immune responses (8). Human chorioamniotic and umbilical cord macrophages express more nuclear IL-33
(also known as full length IL-33) during acute chorioamnionitis (101). Nuclear IL-33 acts as a transcriptional regulator that
has been shown to suppress production of LPS-stimulated proinflammatory cytokines in vitro (102). Moreover, extracellular or
mature IL-33 released from activated or dying cells promotes
M2 polarization of macrophages at the site of inflammation
(103). At best, pro-M2 activation during chorioamnionitis may
minimize detrimental inflammation associated with chorioamnionitis, or it may enhance infection by suppressing antimicrobial
responses.
Manipulation of macrophage polarity at the maternal–fetal
interface by infectious agents can impact pregnancy outcomes. The
obligate intracellular parasite Toxoplasma gondii can invade the
placenta and fetus, resulting in spontaneous abortion, stillbirth,
fetal neurological, and ocular damage (104). A virulent strain of
T. gondii, TgCtwh3, actively induces M2 polarization in infected
macrophages (15). TgCtwh3 rapidly invades the placenta producing a heavy parasite burden with minimal inflammation but a
high rate of trophoblast apoptosis (15). On the other hand, less
virulent strain TgCtwh6 triggers pro-M1 responses in infected
macrophages (15). TgCtwh6 can still invade the placenta, but
infection is contained and associated with a profound lymphocytic inflammatory response with trophoblast apoptosis that is
less extensive than TgCtwh3 (15).

HOFBAUER CELLS IN COMPLICATED PREGNANCY
Placental Hofbauer cells are responsive to paracrine signals from
surrounding cells within the chorionic villus (105). Alterations in
Hofbauer cell densities within the villi have been reported during
chorioamnionitis (106, 107), but there is no evidence that these
cells shift to M1 polarity phenotype during chorioamnionitis.
Instead, HIV infected Hofbauer cells or cells from chorioamnionitis cases continue to display M2 characteristics such as production
of IL-10, TGF-β, expression of DC-SIGN, CD163, and mannose
receptor/CD206 (38, 39). In another study, Hofbauer cells from
chorioamnionitis patients and healthy controls had similar gene
expression profiles, which were not considered to be committed to
either M1 or M2 (108).
Interestingly, it was reported that hyperglycemia can shift Hofbauer cells toward M1 activation (109). Hofbauer cells isolated
from diabetic women displayed characteristics that could be attributed to either M1or M2b phenotype (decreased CD163, CD209,

Frontiers in Immunology | Molecular Innate Immunity

M1/M2 in pregnancy

IL-10 with increased CD68, CCR7, and IL-1β) (109). Further,
when rat Hofbauer cells were cultured in high-glucose conditions
in vitro, these cells expressed increased levels of NOS2 gene expression and NO, clear markers of M1 activation (109). But a caveat
of that experiment was that Hofbauer cells were cultured in
RPMI media, which contains supraphysiologic levels of arginine
(7). High-arginine concentrations in RPMI are not representative
of the tissue microenvironment, especially during inflammation
(7). Further, high-arginine levels will promote iNOS activation in
macrophages in vitro (7).
Folate receptor β (FR-β) is preferentially expressed on M2
polarized macrophages, and is considered a biomarker for
immunoregulatory M2 macrophages (110). Decreased expression of FR-β and CD163, but not CD68, has been observed in
Hofbauer cells isolated from women with severe pre-term preeclampsia (111) suggesting that in this syndrome, Hofbauer cells
may have shifted toward M1 polarity. This would be consistent with Aziza et al.’s study (112) that reported increased iNOS
with concurrent decreased eNOS in placentas from women with
pre-eclampsia.
Villitis of unknown etiology (VUE) is an inflammatory lesion
of the chorionic villi that is associated with intrauterine fetal
growth restriction and perinatal morbidity and mortality (105).
VUE is characterized by influxes of Hofbauer cells, maternal
T-cells (more CD8+ than CD4+ ), and increased expression of
chemokines (CXCL9, CXCL10, CXCL11, CXCL13, CCL4, CCL5,
CXCR3, CCR5) within the placental villi (113, 114). It has been
suggested that the increased expression of chemokines and their
receptors within the villous mesenchyme of VUE patients provides the pro-migratory signals that promote increased migration
of Hofbauer cells and maternal T-cells into the villous (115).
Hofbauer cells do migrate in response to fibroblast secreted
MCP-1 in vitro (106). Whether or not changes in villous mesenchyme promote M1 polarization of Hofbauer cells remains to
be determined.

CONCLUSION
Macrophages serve important roles in the development of hemochorial placentation, maintenance of pregnancy, and initiation of
parturition. As cells that are highly responsive to altering their
polarization pattern in vivo, the mixed M polarity phenotypes
present in the decidua reflect the heterogeneous nature of the
maternal–fetal interface. Both spatial and temporal regulation of
M1 and M2 polarization is required for successful pregnancy.
Similarly, ill-timed or ill-placed macrophage polarization at the
maternal–fetal interface is associated with pregnancy complications and poor outcomes. Despite recent insights into M1/M2
dynamics at the maternal–fetal interface, there are still critical
knowledge gaps concerning the in situ context of macrophage
polarity, the mechanisms that promote dysregulation, and its
impact on healthy pregnancy and obstetric disease.

ACKNOWLEDGMENTS
This project was funded in part by R15 HD81439-01, National
Institute of Child Health and Human Development. The funders had no role in the decision to publish or preparation of the
manuscript.

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M1/M2 in pregnancy

REFERENCES
1. Nagamatsu T, Schust DJ. The immunomodulatory roles of macrophages at
the maternal-fetal interface. Reprod Sci (2010) 17(3):209–18. doi:10.1177/
1933719109349962
2. Co EC, Gormley M, Kapidzic M, Rosen DB, Scott MA, Stolp HA, et al. Maternal
decidual macrophages inhibit NK cell killing of invasive cytotrophoblasts during human pregnancy. Biol Reprod (2013) 88(6):155. doi:10.1095/biolreprod.
112.099465
3. Hamilton S, Oomomian Y, Stephen G, Shynlova O, Tower CL, Garrod A, et al.
Macrophages infiltrate the human and rat decidua during term and preterm
labor: evidence that decidual inflammation precedes labor. Biol Reprod (2012)
86(2):39. doi:10.1095/biolreprod.111.095505
4. Shynlova O, Nedd-Roderique T, Li Y, Dorogin A, Nguyen T, Lye SJ. Infiltration of myeloid cells into decidua is a critical early event in the labour cascade
and post-partum uterine remodelling. J Cell Mol Med (2013) 17(2):311–24.
doi:10.1111/jcmm.12012
5. Shynlova O, Nedd-Roderique T, Li Y, Dorogin A, Lye SJ. Myometrial immune
cells contribute to term parturition, preterm labour and post-partum involution in mice. J Cell Mol Med (2013) 17(1):90–102. doi:10.1111/j.1582-4934.
2012.01650.x
6. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine
system in diverse forms of macrophage activation and polarization. Trends
Immunol (2004) 25(12):677–86. doi:10.1016/j.it.2004.09.015
7. Mills CD. Macrophage arginine metabolism to ornithine/urea or nitric
oxide/citrulline: a life or death issue. Crit Rev Immunol (2001) 21(5):399–425.
doi:10.1615/CritRevImmunol.v21.i5.10
8. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
9. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
10. Comalada M, Yeramian A, Modolell M, Lloberas J, Celada A. Arginine and
macrophage activation. Methods Mol Biol (2012) 844:223–35. doi:10.1007/9781-61779-527-5_16
11. Erlebacher A. Immunology of the maternal-fetal interface. Annu Rev Immunol
(2013) 31:387–411. doi:10.1146/annurev-immunol-032712-100003
12. Guenther S, Vrekoussis T, Heublein S, Bayer B, Anz D, Knabl J, et al. Decidual macrophages are significantly increased in spontaneous miscarriages and
over-express FasL: a potential role for macrophages in trophoblast apoptosis.
Int J Mol Sci (2012) 13(7):9069–80. doi:10.3390/ijms13079069
13. Wang WJ, Hao CF, Lin QD. Dysregulation of macrophage activation by decidual regulatory T cells in unexplained recurrent miscarriage patients. J Reprod
Immunol (2011) 92(1–2):97–102. doi:10.1016/j.jri.2011.08.004
14. Pijnenborg R, Vercruysse L, Hanssens M. The uterine spiral arteries in
human pregnancy: facts and controversies. Placenta (2006) 27(9–10):939–58.
doi:10.1016/j.placenta.2005.12.006
15. Liu T, Zhang Q, Liu L, Xu X, Chen H, Wang H, et al. Trophoblast apoptosis through polarization of macrophages induced by Chinese Toxoplasma
gondii isolates with different virulence in pregnant mice. Parasitol Res (2013)
112(8):3019–27. doi:10.1007/s00436-013-3475-3
16. Martinez FO, Gordon S. The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000Prime Rep (2014) 6:13. doi:10.12703/P6-13
17. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
18. Kurowska-Stolarska M, Stolarski B, Kewin P, Murphy G, Corrigan CJ, Ying S,
et al. IL-33 amplifies the polarization of alternatively activated macrophages
that contribute to airway inflammation. J Immunol (2009) 183(10):6469–77.
doi:10.4049/jimmunol.0901575
19. Nelson MP, Christmann BS, Werner JL, Metz AE, Trevor JL, Lowell CA, et al.
IL-33 and M2a alveolar macrophages promote lung defense against the atypical fungal pathogen Pneumocystis murina. J Immunol (2011) 186(4):2372–81.
doi:10.4049/jimmunol.1002558
20. Pesce JT, Ramalingam TR, Mentink-Kane MM, Wilson MS, El Kasmi KC, Smith
AM, et al. Arginase-1-expressing macrophages suppress Th2 cytokine-driven
inflammation and fibrosis. PLoS Pathog (2009) 5(4):e1000371. doi:10.1371/
journal.ppat.1000371
21. El-Gayar S, Thuring-Nahler H, Pfeilschifter J, Rollinghoff M, Bogdan C. Translational control of inducible nitric oxide synthase by IL-13 and arginine

www.frontiersin.org

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

availability in inflammatory macrophages. J Immunol (2003) 171(9):4561–8.
doi:10.4049/jimmunol.171.9.4561
Ishikawa T, Harada T, Koi H, Kubota T, Azuma H, Aso T. Identification of
arginase in human placental villi. Placenta (2007) 28(2–3):133–8. doi:10.1016/
j.placenta.2006.03.015
Gustafsson C, Mjosberg J, Matussek A, Geffers R, Matthiesen L, Berg G,
et al. Gene expression profiling of human decidual macrophages: evidence
for immunosuppressive phenotype. PLoS One (2008) 3(4):e2078. doi:10.1371/
journal.pone.0002078
Houser BL, Tilburgs T, Hill J, Nicotra ML, Strominger JL. Two unique
human decidual macrophage populations. J Immunol (2011) 186(4):2633–42.
doi:10.4049/jimmunol.1003153
Kim SY, Romero R, Tarca AL, Bhatti G, Kim CJ, Lee J, et al. Methylome of
fetal and maternal monocytes and macrophages at the feto-maternal interface. Am J Reprod Immunol (2012) 68(1):8–27. doi:10.1111/j.1600-0897.2012.
01108.x
Repnik U, Tilburgs T, Roelen DL, van der Mast BJ, Kanhai HH, Scherjon S, et al. Comparison of macrophage phenotype between decidua basalis
and decidua parietalis by flow cytometry. Placenta (2008) 29(5):405–12.
doi:10.1016/j.placenta.2008.02.004
Svensson J, Jenmalm MC, Matussek A, Geffers R, Berg G, Ernerudh J.
Macrophages at the fetal-maternal interface express markers of alternative activation and are induced by M-CSF and IL-10. J Immunol (2011)
187(7):3671–82. doi:10.4049/jimmunol.1100130
Laskarin G, Medancic SS, Redzovic A, Duric D, Rukavina D. Specific decidual
CD14(+) cells hamper cognate NK cell proliferation and cytolytic mediator
expression after mucin 1 treatment in vitro. J Reprod Immunol (2012) 95(1–
2):36–45. doi:10.1016/j.jri.2012.06.002
Laskarin G, Cupurdija K, Tokmadzic VS, Dorcic D, Dupor J, Juretic K,
et al. The presence of functional mannose receptor on macrophages at the
maternal-fetal interface. Hum Reprod (2005) 20(4):1057–66. doi:10.1093/
humrep/deh740
Chabtini L, Mfarrej B, Mounayar M, Zhu B, Batal I, Dakle PJ, et al. TIM-3 regulates innate immune cells to induce fetomaternal tolerance. J Immunol (2013)
190(1):88–96. doi:10.4049/jimmunol.1202176
Han G, Chen G, Shen B, Li Y. Tim-3: an activation marker and activation limiter of innate immune cells. Front Immunol (2013) 4:449. doi:10.3389/fimmu.
2013.00449
Zhang ZY, Schluesener HJ, Zhang Z. Distinct expression of Tim-3 during different stages of rat experimental autoimmune neuritis. Brain Res Bull (2011)
86(3–4):229–34. doi:10.1016/j.brainresbull.2011.07.005
Frisancho-Kiss S, Nyland JF, Davis SE, Barrett MA, Gatewood SJ, Njoku
DB, et al. Cutting edge: T cell Ig mucin-3 reduces inflammatory heart disease by increasing CTLA-4 during innate immunity. J Immunol (2006)
176(11):6411–5. doi:10.4049/jimmunol.176.11.6411
Takahashi K, Naito M, Katabuchi H, Higashi K. Development, differentiation, and maturation of macrophages in the chorionic villi of mouse placenta
with special reference to the origin of Hofbauer cells. J Leukoc Biol (1991)
50(1):57–68.
Karakaya YA, Ozer E. The role of Hofbauer cells on the pathogenesis of early
pregnancy loss. Placenta (2013) 34(12):1211–5. doi:10.1016/j.placenta.2013.
10.010
Ingman K, Cookson VJ, Jones CJ, Aplin JD. Characterisation of Hofbauer cells
in first and second trimester placenta: incidence, phenotype, survival in vitro
and motility. Placenta (2010) 31(6):535–44. doi:10.1016/j.placenta.2010.03.
003
Bockle BC, Solder E, Kind S, Romani N, Sepp NT. DC-sign+ CD163+
macrophages expressing hyaluronan receptor LYVE-1 are located within
chorion villi of the placenta. Placenta (2008) 29(2):187–92. doi:10.1016/j.
placenta.2007.11.003
Johnson EL, Chakraborty R. Placental Hofbauer cells limit HIV-1 replication
and potentially offset mother to child transmission (MTCT) by induction of
immunoregulatory cytokines. Retrovirology (2012) 9:101. doi:10.1186/17424690-9-101
Joerink M, Rindsjo E, van Riel B, Alm J, Papadogiannakis N. Placental
macrophage (Hofbauer cell) polarization is independent of maternal allergensensitization and presence of chorioamnionitis. Placenta (2011) 32(5):380–5.
doi:10.1016/j.placenta.2011.02.003

November 2014 | Volume 5 | Article 606 | 211

Brown et al.

40. Abrahams VM, Kim YM, Straszewski SL, Romero R, Mor G. Macrophages
and apoptotic cell clearance during pregnancy. Am J Reprod Immunol (2004)
51(4):275–82. doi:10.1111/j.1600-0897.2004.00156.x
41. Grasso E, Paparini D, Aguero M, Mor G, Perez Leiros C, Ramhorst R. VIP
contribution to the decidualization program: regulatory T cell recruitment.
J Endocrinol (2014) 221(1):121–31. doi:10.1530/JOE-13-0565
42. Hunt JS, Petroff MG, Burnett TG. Uterine leukocytes: key players in pregnancy.
Semin Cell Dev Biol (2000) 11(2):127–37. doi:10.1006/scdb.2000.0158
43. Thiruchelvam U, Dransfield I, Saunders PT, Critchley HO. The importance
of the macrophage within the human endometrium. J Leukoc Biol (2013)
93(2):217–25. doi:10.1189/jlb.0712327
44. Hunt JS, Pollard JW. Macrophages in the uterus and placenta. Curr Top Microbiol Immunol (1992) 181:39–63.
45. Kammerer U, von Wolff M, Markert UR. Immunology of human
endometrium. Immunobiology (2004) 209(7):569–74. doi:10.1016/j.imbio.
2004.04.009
46. Svensson-Arvelund J, Ernerudh J, Buse E, Cline JM, Haeger JD, Dixon
D, et al. The placenta in toxicology. Part II: systemic and local immune
adaptations in pregnancy. Toxicol Pathol (2014) 42(2):327–38. doi:10.1177/
0192623313482205
47. Paria BC, Reese J, Das SK, Dey SK. Deciphering the cross-talk of implantation: advances and challenges. Science (2002) 296(5576):2185–8. doi:10.1126/
science.1071601
48. Dunn CL, Kelly RW, Critchley HO. Decidualization of the human endometrial stromal cell: an enigmatic transformation. Reprod Biomed Online (2003)
7(2):151–61. doi:10.1016/S1472-6483(10)61745-2
49. Strakova Z, Srisuparp S, Fazleabas AT. IL-1beta during in vitro decidualization in primate. J Reprod Immunol (2002) 55(1–2):35–47. doi:10.1016/S01650378(01)00141-3
50. Bulmer JN, Pace D, Ritson A. Immunoregulatory cells in human decidua:
morphology, immunohistochemistry and function. Reprod Nutr Dev (1988)
28(6B):1599–613. doi:10.1051/rnd:19881006
51. Arcuri F, Buchwalder L, Toti P, Cintorino M, Tosi P, Lockwood CJ, et al. Differential regulation of colony stimulating factor 1 and macrophage migration inhibitory factor expression by inflammatory cytokines in term human
decidua: implications for macrophage trafficking at the fetal-maternal interface. Biol Reprod (2007) 76(3):433–9. doi:10.1095/biolreprod.106.054189
52. Harris LK. IFPA Gabor than Award Lecture: transformation of the spiral arteries in human pregnancy: key events in the remodelling timeline. Placenta
(2011) 32(Suppl 2):S154–8. doi:10.1016/j.placenta.2010.11.018
53. Aldo PB, Racicot K, Craviero V, Guller S, Romero R, Mor G. Trophoblast induces
monocyte differentiation into CD14+/CD16+ macrophages. Am J Reprod
Immunol (2014) 72(3):270–84. doi:10.1111/aji.12288
54. Ivanisevic M, Segerer S, Rieger L, Kapp M, Dietl J, Kammerer U, et al. Antigenpresenting cells in pregnant and non-pregnant human myometrium. Am J
Reprod Immunol (2010) 64(3):188–96. doi:10.1111/j.1600-0897.2010.00858.x
55. Tagliani E, Shi C, Nancy P, Tay CS, Pamer EG, Erlebacher A. Coordinate regulation of tissue macrophage and dendritic cell population dynamics by CSF-1.
J Exp Med (2011) 208(9):1901–16. doi:10.1084/jem.20110866
56. Rosario GX,Ain R, Konno T, Soares MJ. Intrauterine fate of invasive trophoblast
cells. Placenta (2009) 30(5):457–63. doi:10.1016/j.placenta.2009.02.008
57. Shynlova O, Tsui P, Dorogin A, Lye SJ. Monocyte chemoattractant protein-1
(CCL-2) integrates mechanical and endocrine signals that mediate term and
preterm labor. J Immunol (2008) 181(2):1470–9. doi:10.4049/jimmunol.181.
2.1470
58. Benirschke K. Pathology of the Human Placenta. New York, NY: Springer (2012).
59. Harris LK. Review: trophoblast-vascular cell interactions in early pregnancy:
how to remodel a vessel. Placenta (2010) 31(Suppl):S93–8. doi:10.1016/j.
placenta.2009.12.012
60. Smith SD, Dunk CE, Aplin JD, Harris LK, Jones RL. Evidence for immune cell
involvement in decidual spiral arteriole remodeling in early human pregnancy.
Am J Pathol (2009) 174(5):1959–71. doi:10.2353/ajpath.2009.080995
61. Bulmer JN, Burton GJ, Collins S, Cotechini T, Crocker IP, Croy BA, et al.
IFPA meeting 2011 workshop report II: angiogenic signaling and regulation
of fetal endothelial function; placental and fetal circulation and growth; spiral
artery remodeling. Placenta (2012) 33(Suppl):S9–14. doi:10.1016/j.placenta.
2011.11.014

Frontiers in Immunology | Molecular Innate Immunity

M1/M2 in pregnancy

62. Hazan AD, Smith SD, Jones RL, Whittle W, Lye SJ, Dunk CE. Vascular-leukocyte
interactions: mechanisms of human decidual spiral artery remodeling in vitro.
Am J Pathol (2010) 177(2):1017–30. doi:10.2353/ajpath.2010.091105
63. Mor G, Cardenas I, Abrahams V, Guller S. Inflammation and pregnancy: the
role of the immune system at the implantation site. Ann N Y Acad Sci (2011)
1221:80–7. doi:10.1111/j.1749-6632.2010.05938.x
64. Robertson SA. Seminal fluid signaling in the female reproductive tract:
lessons from rodents and pigs. J Anim Sci (2007) 85(13 Suppl):E36–44.
doi:10.2527/jas.2006-578
65. Jaiswal MK, Mallers TM, Larsen B, Kwak-Kim J, Chaouat G, Gilman-Sachs A,
et al. V-ATPase upregulation during early pregnancy: a possible link to establishment of an inflammatory response during preimplantation period of pregnancy. Reproduction (2012) 143(5):713–25. doi:10.1530/REP-12-0036
66. Ntrivalas E, Levine R, Kwong C, Gilman-Sachs A, Beaman K. The a2 isoform of vacuolar ATPase is a modulator of implantation and feto-maternal
immune tolerance in early pregnancy. J Reprod Immunol (2010) 85(1):106–11.
doi:10.1016/j.jri.2009.10.010
67. Jaiswal MK, Gilman-Sachs A, Chaouat G, Beaman KD. Placental ATPase expression is a link between multiple causes of spontaneous abortion in mice. Biol
Reprod (2011) 85(3):626–34. doi:10.1095/biolreprod.111.092494
68. Lombardelli L, Aguerre-Girr M, Logiodice F, Kullolli O, Casart Y, Polgar B, et al.
HLA-G5 induces IL-4 secretion critical for successful pregnancy through differential expression of ILT2 receptor on decidual CD4(+) T cells and macrophages.
J Immunol (2013) 191(7):3651–62. doi:10.4049/jimmunol.1300567
69. Apps R, Gardner L, Sharkey AM, Holmes N, Moffett A. A homodimeric complex of HLA-G on normal trophoblast cells modulates antigen-presenting cells
via LILRB1. Eur J Immunol (2007) 37(7):1924–37. doi:10.1002/eji.200737089
70. Fu B, Li X, Sun R, Tong X, Ling B, Tian Z, et al. Natural killer cells promote immune tolerance by regulating inflammatory TH17 cells at the human
maternal-fetal interface. Proc Natl Acad Sci U S A (2013) 110(3):E231–40.
doi:10.1073/pnas.1206322110
71. Nagamatsu T, Schust DJ. The contribution of macrophages to normal and
pathological pregnancies. Am J Reprod Immunol (2010) 63(6):460–71. doi:10.
1111/j.1600-0897.2010.00813.x
72. Heikkinen J, Mottonen M, Komi J, Alanen A, Lassila O. Phenotypic characterization of human decidual macrophages. Clin Exp Immunol (2003)
131(3):498–505. doi:10.1046/j.1365-2249.2003.02092.x
73. Mues B, Langer D, Zwadlo G, Sorg C. Phenotypic characterization of
macrophages in human term placenta. Immunology (1989) 67(3):303–7.
74. Osman I, Young A, Ledingham MA, Thomson AJ, Jordan F, Greer IA, et al.
Leukocyte density and pro-inflammatory cytokine expression in human fetal
membranes, decidua, cervix and myometrium before and during labour at
term. Mol Hum Reprod (2003) 9(1):41–5. doi:10.1093/molehr/gag001
75. Hamilton SA, Tower CL, Jones RL. Identification of chemokines associated
with the recruitment of decidual leukocytes in human labour: potential novel
targets for preterm labour. PLoS One (2013) 8(2):e56946. doi:10.1371/journal.
pone.0056946
76. Abumaree MH, Al Jumah MA, Kalionis B, Jawdat D, Al Khaldi A, Abomaray FM,
et al. Human placental mesenchymal stem cells (pMSCs) play a role as immune
suppressive cells by shifting macrophage differentiation from inflammatory
M1 to anti-inflammatory M2 macrophages. Stem Cell Rev (2013) 9(5):620–41.
doi:10.1007/s12015-013-9455-2
77. Lambropoulou M, Tamiolakis D, Venizelos J, Liberis V, Galazios G, Tsikouras P, et al. Imbalance of mononuclear cell infiltrates in the placental tissue from foetuses after spontaneous abortion versus therapeutic termination
from 8th to 12th weeks of gestational age. Clin Exp Med (2006) 6(4):171–6.
doi:10.1007/s10238-006-0111-x
78. Khong Y, Brosens I. Defective deep placentation. Best Pract Res Clin Obstet
Gynaecol (2011) 25(3):301–11. doi:10.1016/j.bpobgyn.2010.10.012
79. Reister F, Frank HG, Heyl W, Kosanke G, Huppertz B, Schroder W, et al.
The distribution of macrophages in spiral arteries of the placental bed in
pre-eclampsia differs from that in healthy patients. Placenta (1999) 20(2–
3):229–33. doi:10.1053/plac.1998.0373
80. Reister F, Frank HG, Kingdom JC, Heyl W, Kaufmann P, Rath W, et al.
Macrophage-induced apoptosis limits endovascular trophoblast invasion in
the uterine wall of preeclamptic women. Lab Invest (2001) 81(8):1143–52.
doi:10.1038/labinvest.3780326

November 2014 | Volume 5 | Article 606 | 212

Brown et al.

81. Wu ZM, Yang H, Li M, Yeh CC, Schatz F, Lockwood CJ, et al. Pro-inflammatory
cytokine-stimulated first trimester decidual cells enhance macrophage-induced
apoptosis of extravillous trophoblasts. Placenta (2012) 33(3):188–94. doi:10.
1016/j.placenta.2011.12.007
82. Staff AC, Redman CW. IFPA award in placentology lecture: preeclampsia, the
decidual battleground and future maternal cardiovascular disease. Placenta
(2014) 35(Suppl):S26–31. doi:10.1016/j.placenta.2013.12.003
83. Pijnenborg R, McLaughlin PJ, Vercruysse L, Hanssens M, Johnson PM, Keith
JC Jr, et al. Immunolocalization of tumour necrosis factor-alpha (TNF-alpha)
in the placental bed of normotensive and hypertensive human pregnancies.
Placenta (1998) 19(4):231–9. doi:10.1016/S0143-4004(98)90054-6
84. Staff AC, Dechend R, Pijnenborg R. Learning from the placenta: acute atherosis and vascular remodeling in preeclampsia-novel aspects for atherosclerosis and future cardiovascular health. Hypertension (2010) 56(6):1026–34.
doi:10.1161/HYPERTENSIONAHA.110.157743
85. Reyes L, Herrera D, Kozarov E, Rolda S, Progulske-Fox A. Periodontal bacterial
invasion and infection: contribution to atherosclerotic pathology. J Periodontol
(2013) 84(4 Suppl):S30–50. doi:10.1111/jcpe.12079
86. Chaparro A, Blanlot C, Ramirez V, Sanz A, Quintero A, Inostroza C, et al. Porphyromonas gingivalis, Treponema denticola and toll-like receptor 2 are associated with hypertensive disorders in placental tissue: a case-control study.
J Periodontal Res (2013) 48(6):802–9. doi:10.1111/jre.12074
87. Chaparro A, Sanz A, Quintero A, Inostroza C, Ramirez V, Carrion F, et al.
Increased inflammatory biomarkers in early pregnancy is associated with the
development of pre-eclampsia in patients with periodontitis: a case control
study. J Periodontal Res (2013) 48(3):302–7. doi:10.1111/jre.12008
88. Barak S, Oettinger-Barak O, Machtei EE, Sprecher H, Ohel G. Evidence of
periopathogenic microorganisms in placentas of women with preeclampsia.
J Periodontol (2007) 78(4):670–6. doi:10.1902/jop.2007.060362
89. Swati P, Thomas B, Vahab SA, Kapaettu S, Kushtagi P. Simultaneous detection of periodontal pathogens in subgingival plaque and placenta of women
with hypertension in pregnancy. Arch Gynecol Obstet (2012) 285(3):613–9.
doi:10.1007/s00404-011-2012-9
90. Moura da Silva G, Coutinho SB, Piscoya MD, Ximenes RA, Jamelli SR. Periodontitis as a risk factor for preeclampsia. J Periodontol (2012) 83(11):1388–96.
doi:10.1902/jop.2012.110256
91. Leon R, Silva N, Ovalle A, Chaparro A, Ahumada A, Gajardo M, et al. Detection
of Porphyromonas gingivalis in the amniotic fluid in pregnant women with a
diagnosis of threatened premature labor. J Periodontol (2007) 78(7):1249–55.
doi:10.1902/jop.2007.060368
92. Dasanayake AP, Boyd D, Madianos PN, Offenbacher S, Hills E. The association between Porphyromonas gingivalis-specific maternal serum IgG and low
birth weight. J Periodontol (2001) 72(11):1491–7. doi:10.1902/jop.2001.72.11.
1491
93. Sasahara J, Kikuchi A, Takakuwa K, Sugita N, Abiko Y, Yoshie H, et al.
Antibody responses to Porphyromonas gingivalis outer membrane protein
in the first trimester. Aust N Z J Obstet Gynaecol (2009) 49(2):137–41.
doi:10.1111/j.1479-828X.2009.00958.x
94. Lin D, Smith MA, Elter J, Champagne C, Downey CL, Beck J, et al. Porphyromonas gingivalis infection in pregnant mice is associated with placental dissemination, an increase in the placental Th1/Th2 cytokine ratio, and fetal
growth restriction. Infect Immun (2003) 71(9):5163–8. doi:10.1128/IAI.71.9.
5163-5168.2003
95. Lin D, Smith MA, Champagne C, Elter J, Beck J, Offenbacher S. Porphyromonas gingivalis infection during pregnancy increases maternal tumor
necrosis factor alpha, suppresses maternal interleukin-10, and enhances fetal
growth restriction and resorption in mice. Infect Immun (2003) 71(9):5156–62.
doi:10.1128/IAI.71.9.5156-5162.2003
96. Phillips P, Brown MB, Progulske-Fox A, Reyes L. Porphyromonas gingivalis
strain specific effects on the placental bed. Am J Reprod Immunol (2014)
71(Suppl 1):75–6. doi:10.1111/aji.12255
97. Belanger M, Reyes L, von Deneen K, Reinhard MK, Progulske-Fox A, Brown
MB. Colonization of maternal and fetal tissues by Porphyromonas gingivalis
is strain-dependent in a rodent animal model. Am J Obstet Gynecol (2008)
199(1):86.e1–7. doi:10.1016/j.ajog.2007.11.067
98. Gomez-Lopez N, Mial T, Robertson SA. Infection-induced preterm delivery
does not involve a shift in macrophage polarization from the M2/M1phenotype

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M1/M2 in pregnancy

but implicates a maternal pro-inflammatory state. Am J Reprod Immunol
(2014) 71(Suppl 1):68. doi:10.1111/aji.12255
99. von Chamier M, Allam A, Brown MB, Reinhard MK, Reyes L. Host genetic
background impacts disease outcome during intrauterine infection with Ureaplasma parvum. PLoS One (2012) 7(8):e44047. doi:10.1371/journal.pone.
0044047
100. Allam AB, von Chamier M, Brown MB, Reyes L. Immune profiling of BALB/C
and C57BL/6 mice reveals a correlation between Ureaplasma parvum-induced
fetal inflammatory response syndrome-like pathology and increased placental
expression of TLR2 and CD14. Am J Reprod Immunol (2014) 71(3):241–51.
doi:10.1111/aji.12192
101. Topping V, Romero R, Than NG, Tarca AL, Xu Z, Kim SY, et al. Interleukin33 in the human placenta. J Matern Fetal Neonatal Med (2013) 26(4):327–38.
doi:10.3109/14767058.2012.735724
102. Sharma S, Kulk N, Nold MF, Graf R, Kim SH, Reinhardt D, et al. The IL-1
family member 7b translocates to the nucleus and down-regulates proinflammatory cytokines. J Immunol (2008) 180(8):5477–82. doi:10.4049/jimmunol.
180.8.5477
103. Li D, Guabiraba R, Besnard AG, Komai-Koma M, Jabir MS, Zhang L, et al. IL-33
promotes ST2-dependent lung fibrosis by the induction of alternatively activated macrophages and innate lymphoid cells in mice. J Allergy Clin Immunol
(2014). doi:10.1016/j.jaci.2014.05.011
104. Chaudhry SA, Gad N, Koren G. Toxoplasmosis and pregnancy. Can Fam Physician (2014) 60(4):334–6.
105. Tang Z, Abrahams VM, Mor G, Guller S. Placental Hofbauer cells and complications of pregnancy. Ann N Y Acad Sci (2011) 1221:103–8. doi:10.1111/j.
1749-6632.2010.05932.x
106. Toti P, Arcuri F, Tang Z, Schatz F, Zambrano E, Mor G, et al. Focal
increases of fetal macrophages in placentas from pregnancies with histological
chorioamnionitis: potential role of fibroblast monocyte chemotactic protein1. Am J Reprod Immunol (2011) 65(5):470–9. doi:10.1111/j.1600-0897.2010.
00927.x
107. Vinnars MT, Rindsjo E, Ghazi S, Sundberg A, Papadogiannakis N. The number
of CD68(+) (Hofbauer) cells is decreased in placentas with chorioamnionitis
and with advancing gestational age. Pediatr Dev Pathol (2010) 13(4):300–4.
doi:10.2350/09-03-0632-OA.1
108. Ben Amara A, Gorvel L, Baulan K, Derain-Court J, Buffat C,Verollet C, et al. Placental macrophages are impaired in chorioamnionitis, an infectious pathology
of the placenta. J Immunol (2013) 191(11):5501–14. doi:10.4049/jimmunol.
1300988
109. Sisino G, Bouckenooghe T, Aurientis S, Fontaine P, Storme L, Vambergue A.
Diabetes during pregnancy influences Hofbauer cells, a subtype of placental
macrophages, to acquire a pro-inflammatory phenotype. Biochim Biophys Acta
(2013) 1832(12):1959–68. doi:10.1016/j.bbadis.2013.07.009
110. Samaniego R, Palacios BS, Domiguez-Soto A, Vidal C, Salas A, Matsuyama
T, et al. Macrophage uptake and accumulation of folates are polarizationdependent in vitro and in vivo and are regulated by activin A. J Leukoc Biol
(2014) 95(5):797–808. doi:10.1189/jlb.0613345
111. Tang Z, Buhimschi IA, Buhimschi CS, Tadesse S, Norwitz E, NivenFairchild T, et al. Decreased levels of folate receptor-beta and reduced numbers of fetal macrophages (Hofbauer cells) in placentas from pregnancies with severe pre-eclampsia. Am J Reprod Immunol (2013) 70(2):104–15.
doi:10.1111/aji.12112
112. Ariza AC, Bobadilla N, Diaz L, Avila E, Larrea F, Halhali A. Placental gene
expression of calcitonin gene-related peptide and nitric oxide synthases
in preeclampsia: effects of magnesium sulfate. Magnes Res (2009) 22(1):
44–9. doi:10.1684/mrh.2009.0154
113. Kim JS, Romero R, Kim MR, Kim YM, Friel L, Espinoza J, et al.
Involvement of Hofbauer cells and maternal T cells in villitis of unknown
aetiology. Histopathology (2008) 52(4):457–64. doi:10.1111/j.1365-2559.2008.
02964.x
114. Kim MJ, Romero R, Kim CJ, Tarca AL, Chhauy S, LaJeunesse C, et al.
Villitis of unknown etiology is associated with a distinct pattern of chemokine
up-regulation in the feto-maternal and placental compartments: implications for conjoint maternal allograft rejection and maternal anti-fetal graftversus-host disease. J Immunol (2009) 182(6):3919–27. doi:10.4049/jimmunol.
0803834

November 2014 | Volume 5 | Article 606 | 213

Brown et al.

115. Tamblyn JA, Lissauer DM, Powell R, Cox P, Kilby MD. The immunological
basis of villitis of unknown etiology – review. Placenta (2013) 34(10):846–55.
doi:10.1016/j.placenta.2013.07.002
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 14 August 2014; accepted: 11 November 2014; published online: 24 November
2014.

Frontiers in Immunology | Molecular Innate Immunity

M1/M2 in pregnancy

Citation: Brown MB, von Chamier M, Allam AB and Reyes L (2014) M1/M2
macrophage polarity in normal and complicated pregnancy. Front. Immunol. 5:606.
doi: 10.3389/fimmu.2014.00606
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Brown, von Chamier, Allam and Reyes. This is an open-access
article distributed under the terms of the Creative Commons Attribution License (CC
BY). The use, distribution or reproduction in other forums is permitted, provided the
original author(s) or licensor are credited and that the original publication in this
journal is cited, in accordance with accepted academic practice. No use, distribution or
reproduction is permitted which does not comply with these terms.

November 2014 | Volume 5 | Article 606 | 214

REVIEW ARTICLE
published: 22 January 2015
doi: 10.3389/fimmu.2014.00683

Ontogeny and polarization of macrophages in
inflammation: blood monocytes versus tissue
macrophages
Adwitia Dey 1,2 † , Joselyn Allen 1,3 † and Pamela A. Hankey-Giblin 1,2,3 *
1
2
3

Department of Veterinary and Biomedical Sciences, The Pennsylvania State University, University Park, PA, USA
Graduate Program in Physiology, The Pennsylvania State University, University Park, PA, USA
Graduate Program in Immunology and Infectious Disease, The Pennsylvania State University, University Park, PA, USA

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Wolfgang J. Streit, University of
Florida, USA
Masaaki Murakami, Hokkaido
University, Japan
*Correspondence:
Pamela A. Hankey-Giblin, 115 Henning
Building, University Park, PA 16802,
USA
e-mail: phc7@psu.edu
†

Co-first authors

The explosion of new information in recent years on the origin of macrophages in the
steady-state and in the context of inflammation has opened up numerous new avenues
of investigation and possibilities for therapeutic intervention. In contrast to the classical model of macrophage development, it is clear that tissue-resident macrophages can
develop from yolk sac-derived erythro-myeloid progenitors, fetal liver progenitors, and
bone marrow-derived monocytes. Under both homeostatic conditions and in response to
pathophysiological insult, the contribution of these distinct sources of macrophages varies
significantly between tissues. Furthermore, while all of these populations of macrophages
appear to be capable of adopting the polarized M1/M2 phenotypes, their respective contribution to inflammation, resolution of inflammation, and tissue repair remains poorly
understood and is likely to be tissue- and disease-dependent. A better understanding of
the ontology and polarization capacity of macrophages in homeostasis and disease will
be essential for the development of novel therapies that target the inherent plasticity of
macrophages in the treatment of acute and chronic inflammatory disease.
Keywords: tissue-resident macrophages, M1M2, microglia, adipose tissue macrophages, Kupffer cells, obesity,
neurodegenerative disease, hepatic steatosis

Macrophages are a heterogeneous population of immune cells
that have a range of roles in both the induction and resolution of inflammation. Tissue-resident macrophages promote tissue homeostasis and exhibit unique transcriptional profiles and
characteristics depending on the tissue in which they reside (1).
For instance, alveolar macrophages regulate pulmonary surfactant turnover while osteoclasts promote bone resorption, and
red-pulp macrophages (RPMs) in the spleen promote red blood
cell clearance and regulate iron recycling. In addition, multiple
macrophage subtypes occur within a given tissue, and they can
perform distinct functions depending on their anatomical location
(2). For example, in the bone there are at least two types of tissue
macrophages. TRAP+ F4/80− osteoclasts promote bone resorption while F4/80+ CD169+ TRAP− tissue-resident macrophages
promote red blood cell development by providing a niche that
promotes erythropoiesis and engulfing the extruded nuclei of
red blood cell (RBC) progenitors at late stages of development (Figure 1A). Depletion of CD169+ macrophages results in
impaired recovery of mice from hemolytic anemia (3). Recent
studies suggest these two macrophage populations in the bone
are replenished by distinct subsets of monocytes in response to
stress. Mac3+ F4/80− monocytes regulate osteoclast activity but
are not thought to be precursors of osteoclasts under homeostatic conditions (Figure 1A). However, these cells can differentiate
into osteoclasts under inflammatory conditions. Alternatively, in
the presence of increased levels of extracellular heme resulting

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from conditions of stress such as hemolytic anemia, a subpopulation of monocytes in the bone marrow and spleen develop into
F4/80+ CD11bhi progenitors termed pre-RPMs and subsequently
into RPMs and F4/80+VCAM+ bone marrow macrophages that
resemble RPMs to re-establish iron homeostasis (Figure 1A) (4).
In the developing embryo, hematopoiesis begins in the yolk
sac where primitive erythrocytes and macrophages develop in
the absence of hematopoietic stem cells (HSCs). Subsequently,
HSCs arise in the aorto-gonado-mesonephric (AGM) region
and eventually migrate to the fetal liver where development of
all hematopoietic lineages from HSCs occurs. In the neonate,
hematopoiesis moves from the fetal liver to the bone marrow
where it persists throughout adulthood. The traditional view of
macrophage origin contended that tissue-resident macrophages
develop from bone marrow-derived monocytes. It was subsequently believed that tissue-resident macrophages derive from
Ly6C− “resident” monocytes that traffic to tissues under homeostatic conditions. However, studies utilizing Runx1CreER and
Csf1CreER mice, which allow for tamoxifen-inducible activation
of Cre during the early stages of yolk sac hematopoiesis crossed
with Rosa26 mice to label yolk sac-derived (YS) cells (5) have
indicated that some tissue macrophages arise from the yolk sac
and seed tissues in the embryo where they repopulate throughout adulthood (6, 7). Studies utilizing Flt3Cre mice to mark
HSC-derived macrophages confirm that YS tissue macrophages
(Flt3Cre− ) are distinct from HSC-derived macrophages (Flt3Cre+ )

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Dey et al.

FIGURE 1 | Differentiation of resident macrophages in various
physiological states. (A) Two tissue macrophage populations reside in
bone, F4/80− TRAP+ osteoclasts and F4/80+ CD169+ TRAP−
tissue-resident macrophages (MΦs). Osteoclasts are developmentally
dependent on Mac3+ F4/80− monocytes but only during inflammation.
Alternatively, tissue-resident macrophages in bone can be maintained by a
heme-induced subset of monocytes (CD11bhi Ly6C+ SPI-C− ). (B) Cardiac
tissue-resident macrophage populations are primarily yolk sac-derived
(YS) however a minor subset of the population is derived from fetal liver
and HSC-derived progenitors. In homeostasis, primary yolk sac-derived

thus challenging the traditional view of macrophage ontogeny.
Kupffer cells, microglia, and cardiac tissue macrophages are primarily YS and are maintained throughout adulthood. These cells
are independent of the transcription factor Myb, which is required
for the development of HSCs, suggesting these populations of
tissue-resident macrophages develop and persist independent of
contribution from HSCs (8).
While Langerhans cells are first seeded by yolk sac macrophages,
these cells are largely replaced by fetal liver-derived monocytes at later stages of embryonic development as demonstrated
by adoptive transfer of fetal liver monocytes to host embryos
(9). In addition, studies utilizing the CX3 C chemokine receptor
1 (CX3 CR1) promoter, which marks fetal-derived resident

Frontiers in Immunology | Molecular Innate Immunity

Macrophage ontogeny and polarization

cardiac resident macrophages are maintained through self-renewal but in
response to cardiac insult, Ly6Chi monocytes contribute to all four
macrophage populations. (C) Selective transcriptional control plays an
important role in the development of different types of macrophage
populations. While red-pulp macrophages are dependent on SPI-C activity
for development and maintenance, marginal zone (MZ) macrophage
differentiation is mediated by LXRα. Alternatively, Gata6 is mandatory for
the differentiation and proliferation of peritoneal macrophages while
GM-CSF-dependent induction of PPARγ regulate alveolar macrophage
development.

macrophage populations to drive GFP, Cre or a tomoxifeninducible Cre together with adoptive transfer studies have demonstrated that peritoneal macrophages and alveolar macrophages are
also largely fetal liver-derived. While cardiac tissue macrophages
are also largely YS, two minor populations also display contribution from fetal liver and bone marrow progenitors (10–12). Other
tissue-resident macrophages including those from spleen, pancreas, and kidney also exhibit mixed contribution from fetal and
adult HSC-derived precursors (13), while intestinal macrophages
appear to be entirely derived from circulating monocytes (14, 15).
While these studies make a distinction between YS and fetal liverderived macrophages, a recent report describes an erythro-myeloid
progenitor (EMP) population that develops in the yolk sac and

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later migrates to the liver. These progenitors are distinct from
HSC-derived hematopoiesis, suggesting that YS and fetal liverderived tissue-resident macrophages have a common origin in the
yolk sac (16).
Unlike previous assumptions that macrophages are terminally
differentiated cells, YS macrophages appear to have stem celllike properties in that they can proliferate and self-renew (9).
In the lung, tissue-resident macrophages are maintained locally
in the steady state. Moreover, following irradiation and bone
marrow transplantation, residual lung-resident macrophages can
re-establish normal homeostasis when the development of donorderived macrophages is compromised (13). These cells expand in
response to colony-stimulating factors, M-CSF and GM-CSF, in
an interleukin 4 (IL-4)-independent manner. It is therefore likely
that, following irradiation, the expansion of resident macrophages
is delayed giving the monocyte-derived macrophages a developmental advantage. Alternatively, in response to helminth infection, Th2-dependent production of IL-4 promotes the proliferation of lung-resident macrophages (17) and this proliferation is independent of colony-stimulating factor-1 (CSF-1)
(18). However, while Langerhans cells are embryonic-derived
cells that exhibit hallmarks of self-maintenance, these cells are
eventually replaced by circulating precursors following nonmyeloablative transplantation (19). Similarly, YS cardiac resident
macrophages are maintained through local proliferation, while
monocyte-derived cells can replace all populations of resident cardiac macrophages following depletion or in response to damage
(Figure 1B) (10, 20).
The development and maintenance of tissue-resident
macrophages is also under tissue-selective transcriptional control.
The transcription factor Gata6 is responsible for the transcriptome profile of resident peritoneal macrophages as well as for
their proliferation under homeostatic conditions and in response
to inflammation (Figure 1C) (21) and this programing is regulated reversibly by retinoic acid-induced Gata6 expression (22).
Reflecting the fetal origin of peritoneal macrophages, retinoic
acid induces expression of Gata6 in fetal-derived macrophages
but not bone marrow-derived macrophages (BMDMs) due to
epigenetic silencing of the Gata6 locus in BMDMs (22). Alternatively, the development of alveolar macrophages from fetal
monocytes shortly after birth is regulated by the GM-CSFdependent induction of peroxisome proliferator-activated receptor γ (PPARγ), resulting in a transcriptosome profile unique
to alveolar macrophages (Figure 1C) (23). The development
of splenic RPMs in response to excess heme requires the transcription factor SPI-C (Figure 1C) (4), however, marginal zone
(MZ) macrophages develop normally in SPI-C knockout animals. MZ macrophages play a critical role in the recognition
and uptake of blood-borne antigens. While the differentiation
of RPMs requires SPI-C, the differentiation of both populations of MZ macrophages is under the regulatory control of the
nuclear receptor, liver x receptor α (LXRα) (Figure 1C) (24), and
LXRα-deficient mice exhibit abnormal responses to blood-borne
antigens.
Unlike previous assumptions that Ly6C− monocytes are precursors of tissue-resident macrophages, it has recently been
shown that these monocytes do not infiltrate tissues but rather

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Macrophage ontogeny and polarization

have a “patrolling” function in promoting endothelial integrity.
Alternatively, Ly6C+ “inflammatory” monocytes are actively
recruited to inflamed tissues by the chemokine, chemokine ligand 2 (CCL2) where they promote inflammation and pathogen
clearance. Analogous populations of monocytes in humans have
been described based on gene expression analysis and are classified as CD14hi CD16− , which are the primary subset in healthy
individuals and resemble Ly6C+ murine monocytes while the
minor CD14lo CD16+ population of human monocytes shares
gene expression profiles with murine Ly6C− monocytes (25). Furthermore, the CD14lo CD16+ population of human monocytes,
like Ly6C− monocytes, also play a role in patrolling the endothelium (26). Ly6C− monocytes derive in the circulation from the
Ly6C+ population and exhibit an extended half-life compared
with Ly6C+ monocytes (9). Intriguingly, recent studies indicate
that a subset of Ly6C+ monocytes can traffic to tissues under
homeostatic conditions where they fail to differentiate into tissueresident macrophages, rather they acquire antigen, which they
carry to the draining lymph nodes (27).
Under non-inflammatory conditions, tissue-resident macrophages largely exhibit an M2 phenotype that promotes tissue
homeostasis and repair. However, upon infection, M1 macrophage
activation is induced by the engagement of pattern recognition receptors (PPRs) by pathogen-associated molecular patterns
(PAMPs) such as lipopolysaccharide (LPS), in cooperation with
interferon γ (IFNγ) (28). These cells produce cytotoxic oxygen
and nitrogen radicals and pro-inflammatory cytokines resulting
in increased microbicidal activity. M2 macrophage activation is
induced by the Th2 chemokines IL-4 and interleukin-13 (IL-13)
(29, 30). Alternatively, during the resolution of inflammation, the
balance of macrophage activation toward an M2 phenotype occurs
in order to promote clearance of debris, inhibit the production
of inflammatory mediators, and restore tissue homeostasis. M2
macrophages produce anti-inflammatory cytokines and express
endocytic receptors, and these cells promote the clearance of apoptotic cells, proliferation, and wound healing (28). However, the
relative contribution and level of plasticity of tissue-resident versus monocyte-derived macrophage populations is only recently
beginning to be elucidated.
The M1/M2 macrophage subsets are most commonly distinguished based on the catabolism of l-arginine. While classically activated macrophages express increased levels of inducible
nitric oxide synthase (iNOS), which converts l-arginine to lcitrulline and nitric oxide, alternatively activated macrophages
express arginase I (ArgI), which catabolizes l-arginine to lornithine, a precursor of polyamines and proline. While Th1
and Th2 cytokines promote M1 and M2 macrophage activation,
respectively, these phenotypes are observed in T cell deficient mice
suggesting that T helper cells are not required to drive macrophage
polarization in vivo (29, 30). Conversely, M1 and M2 macrophages
are capable of promoting Th1 and Th2 differentiation, respectively,
suggesting these polarized macrophage phenotypes play an important role in driving the immune response to environmental insult
(29). Recent attempts have been made to re-classify macrophage
subpopulations in response to a range of stimuli and increasingly complex combination of markers (31). However, in vivo,
macrophages are exposed simultaneously to a diverse array of

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signals, so it remains to be determined to what extent distinct
subpopulations exist in vivo.
The origin and plasticity of macrophages in chronic inflammation remains poorly understood and is likely variable depending
on the tissue and the underlying cause of inflammation. Tissueresident macrophages as well as monocyte-derived macrophages
appear to have the capability of adopting both M1 and M2 phenotypes, however, the relative contribution of these subsets of
macrophages to the progression and resolution of chronic inflammation remains enigmatic. Here, we will review current progress in
understanding the complex relationships between tissue-resident
macrophages and infiltrating monocyte-derived macrophages in
homeostasis and chronic inflammation and their contributions to
M1/M2 polarized phenotypes in homeostasis and disease with a
focus on chronic inflammation in the brain, adipose tissue, and
liver.

MICROGLIA: RESIDENT CNS MACROPHAGES
Microglia, parenchymal tissue-resident macrophages in the central
nervous system, plays a central role in mediating tissue homeostasis in health and disease (32). Fate mapping studies conducted
independently by Ginhoux et al. and Prinz et al. demonstrated the
embryonic origin of microglia from c-kit+ EMP prior to the formation of blood–brain barrier and vascularization of the embryo
(7, 33, 34). The colony-stimulating-factor 1 receptor (CSF1R) and
its ligand, CSF-1, play a central role in macrophage development
and maintenance. However, while CSF1R is essential for the development and maintenance of microglia, mice deficient for CSF-1
exhibit only a partial reduction in microglial cells. Recent studies have identified an additional ligand for CSF1R, interleukin-34
(IL-34), which plays an essential role in the development and
maintenance of microglia. While mice deficient for IL-34 exhibit
significantly decreased numbers of microglia and Langerhans cells,
the development of other tissue macrophage populations remain
largely intact (35, 36). After birth, the microglia undergo massive
expansion in response to CSF-1 and IL-34 in order to populate
the developing nervous system. Cortical, optical, and spinal cord
microgliogenesis is sustained by the transcription factors Pu.1 and
interferon regulatory factor-8 (IRF-8), and is independent of Myb,
which is required for the development of hematopoietic stem cells
(34, 37). Microglia are sustained and self-renew within host tissues
throughout adulthood independent of progenitors from the bone
marrow (38).
In a healthy brain, microglia are dispersed relatively uniformly
throughout the parenchymal tissue. Microglia are characterized
by morphological features that reflect their functional capacity.
In a healthy brain, microglia are in a quiescent state or have a
“down-regulated” phenotype exhibited by a ramified shape, similar to Langerhans cells that have short fine processes and thus
increased surface area for tissue surveillance (39). This downregulated phenotype is characterized by an attenuated innate
immune function correlated with decreased expression of CD45,
MHCII, and Fc receptors (39–41). Initially, it was believed that
steady-state microglia are static, however, recent studies have characterized the quiescent phenotype as a more dynamic state in
which microglia are performing housekeeping functions by constitutively surveying the parenchymal tissue (42). This dynamic

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state is characterized by increased expression of oxidative genes
and favors homeostatic tissue remodeling and steady-state wound
healing (43).
Recent studies by Butofsky et al. (44) demonstrate that resident microglia exhibit a distinct expression profile that is not
observed in microglial cell lines and is distinct from M1 or M2
polarized microglia, but rather includes genes associated with
nervous system development. The induction of this unique profile is highly dependent on transforming growth factor β (TGFβ) – in vitro in human microglial cultures, and these cells are
absent in TGF-β knockout mice. The down-regulated phenotype
of microglia is critical for normal neuronal growth, and intimate interactions between neurons and microglia are important
for optimal synaptic growth and maintenance (Figure 2) (45).
The interaction between neurons and microglia is fostered by
the chemokine CX3 C chemokine ligand 1 (CX3 CL1; also known
as fractalkine) and CD200 membrane proteins expressed on
healthy neurons that interact with their respective transmembrane
protein receptors on microglia, CX3 CR1, and CD200R, respectively (32, 46–48). These microglial receptors carry immunoreceptor tyrosine-based inhibitory motifs (ITIMs) such that, upon
ligand–receptor interaction, the ITIM based receptors suppress
downstream immune signaling through the recruitment of Src
homology 2 domain-containing phosphatase 1 (SHP-1) (32, 48–
50). These cell–cell mediated interactions act synergistically with
constitutively released neurotropic elements in order to create an environment fostering the down-regulated phenotype of
microglia.
Whether or not bone marrow-derived cells can contribute to
the microglial population remains under debate (reviewed in Prinz
et al.). Bone marrow transplantation of irradiated recipient mice
demonstrated that macrophages in the perivascular space and
choroid plexus can be repopulated by donor cells (51, 52). While
microglia are largely radio-resistant cells, upon irradiation and
bone marrow transplantation, bone marrow-derived progenitor
cells that are distinct from monocytes have been shown to seed
the brain and differentiate into microglial-like macrophage cells
(45). However, it is not clear whether BMDMs maintain the range
of functions exhibited by YS microglia. In addition, irradiation
results in the breakdown of the blood–brain barrier, thus whether
the seeding of the brain with peripheral bone marrow-derived
cells can occur under steady-state conditions in the presence of an
intact blood–brain barrier is not known.
Like macrophages in the periphery, in vitro studies demonstrate that M1 and M2 activation can be induced in microglia by
LPS and IL-4, respectively (40, 53). However, whether this phenotypic switching of resident microglia occurs in vivo remains
unclear. Experimental models of Alzheimer’s disease (AD), aging,
and multiple sclerosis (MS) (54, 55) have demonstrated microglial
heterogeneity analogous to systemic tissue-resident populations.
These studies suggest that early in disease progression, microglial
cells develop an altered inducible “activated” state that is functionally different from steady-state microglia. This activated state is
then further subdivided into a classical M1 and alternative M2
state (32, 40). Morphologically, activated microglia exhibit an
amoeboid shape in contrast to the quiescent ramified shape of
steady-state microglia (42). While the quiescent state of microglia

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FIGURE 2 | Microglial–neuronal interactions in health and disease.
Healthy neurons expressing chemokine fractalkine (CX3 CL1) and CD200
membrane proteins intimately interact with their respective transmembrane
protein receptors on microglia, CX3 CR1, and CD200R to sustain a
down-regulated microglial phenotype. Microglial receptors have

is maintained by neuronal–glial interactions, in the course of
inflammation, neuronal, and glial interactions are disrupted due
to degeneration of neurons (43, 56). Disruption of CX3 CL1–
CX3 CR1 interactions releases microglia from tonic inhibition (48),
resulting in microglial activation as a consequence (Figure 2).
Additionally, a unique “primed” microglial state, which exhibits
a functional state somewhat distinct from the activated states,
has been described (57). These microglia are thought to function as more of an adaptive response (57). This primed state
refers to a mode of preconditioning underlying chronic inflammation whereby microglial cells are activated continuously by being
persistently exposed to a pro-inflammatory milieu (32, 57). Importantly this priming state enables the conditioned microglia to
mount a heightened response to secondary or future immunological insult in the course of disease progression. Primed microglia
exhibit a more robust inflammatory response as compared to
those that were not subjected to prior stimulatory challenges
(32, 57). Additionally, the heightened inflammatory response to a
physiological challenge is believed to promote a switch from a more

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Macrophage ontogeny and polarization

immunoreceptor tyrosine-based inhibitory motifs (ITIMs), which upon
ligand–receptor activation suppresses downstream immune signaling through
the recruitment of phosphatases including SHP-1. Chronic inflammation
disrupts this intimate neuronal–glial interaction, thus releasing the microglial
cells from a down-regulated inhibited state to an activated phenotype.

anti-inflammatory to a pro-inflammatory reactive phenotype
resulting in cellular stress and exacerbated inflammation (57).
Investigations in spinal cord injury (SCI), AD, and the animal
model of MS, also known as experimental autoimmune encephalitis (EAE) demonstrate an increase in both M1 and M2 phenotypes,
possibly acting synergistically in an attempt to mitigate inflammation (40). While both M1 and M2 transcripts increase in a
majority of chronic CNS disease models, the M1 phenotype offsets the M2 phenotype in the early stages of chronic inflammation,
disrupting normal neuronal/glial cross-talk, and promoting a proinflammatory milieu implicated in the progression of disease (39,
40, 53). With progressive degeneration, the M1 phenotype predominates, as M2 microglia exhibit decreased responsiveness to
anti-inflammatory cues (53). Alternatively, the classically activated
M1 phenotype is predominately induced by acute pathological
conditions such as stroke, traumatic brain injury, or experimentally induced systemic inflammation by LPS (42). The M1
state in chronic and acute manifestations is characterized by the
up-regulation of MHCII, CD86, and Fc receptors and increased

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production of pro-inflammatory cytokines such as interleukin6 (IL-6), interleukin-1 (IL-1), and tumor necrosis factor alpha
(TNFα) (58). Furthermore aging models hint at a possibility of
phenotypic switch in microglial activation from an M2 to M1
state over time, coincident with decreasing neurogenesis and a
growing inflammatory niche (32). However, it still remains unclear
whether these populations of macrophages are due to phenotypic
switching of resident microglia, or the activation states of infiltrating monocyte-derived macrophages. Interestingly, research in
microglial senescence sheds light into the notion that microglial
degradation precedes neurodegeneration and perhaps neurodegeneration is a secondary factor to aging induced dysregulation of
microglial populations (59, 60).
Several studies of chronic inflammation in the CNS have
demonstrated a degree of immune infiltration into the CNS, which
is presumed to promote tissue remodeling as an attempt to maintain homeostasis (61, 62). The observed monocytic infiltration is
attributed to the breakdown and weakening of the blood–brain
barrier, which then facilitates the entrance of immune cells (63,
64). However, while monocyte infiltration has traditionally been
considered a late event in disease development recent studies by
Yamasaki et al. using an EAE model, demonstrated that infiltration
of Ly6C+ monocytes occurs at the onset of disease and that these
infiltrating cells are responsible for initiating nodal demyelination
(65). It was further demonstrated that, while infiltrating monocytes are responsible for initiating nodal demyelination, resident
microglia are relatively quiescent upon disease onset (65–67). The
infiltrating monocytes promote disease progression though they
do not appear to contribute to the resident microglial population
(68). Similarly in a model of AD immune cell infiltration and
localization of microglial populations in areas of amyloid β (Aβ)
plaque deposits has been observed (69, 70).
Recently, researchers have been targeting microglial genes, such
as TGF-β activated kinase (TAK1) to elucidate how tissue-resident
macrophages are behaving during inflammation. While infiltrating monocytes are highly phagocytic and inflammatory, resident
microglia are relatively quiescent upon disease onset. However, a
recent study by Goldmann et al. demonstrated an essential role
of microglia in the onset and progression of disease in an EAE
model (71). By targeting TAK1 in microglia, but not peripheral monocytes/macrophages, they found that the lack of TAK1
in microglia inhibited the progression of inflammation in the
CNS, highlighting the contribution of both resident microglia
and bone marrow-derived monocytes in the onset and progression of EAE. However, in a model of EAE, infiltrating monocytes
are not maintained and do not appear to contribute to the resident microglial population (68). Furthermore, recent studies using
intravital correlated microscopy in zebrafish show that infiltrating phagocytes undergo cell death and are engulfed by resident
microglia during resolution of inflammation (72). Popovich et al.
investigated the potential for targeting CX3 CR1 in a model of SCI
as the ligand–receptor interactions between CX3 CR1 and CX3 CL1
are fundamental for optimal microglial function (47). Furthermore, Donelly et al. presented two important findings, targeted
CX3 CR1 deficiency/blocking in SCI results in enhanced recovery
and decreased monocytic infiltration to injury site (48).

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ADIPOSE TISSUE MACROPHAGES
While it has long been clear that undernutrition can impair
immunocompetence, more recently, the progression of obesity
has also been attributed to a shift in immune function from an
M2 to M1 type responses (73). Work by both Spiegelman (74, 75)
and Lumeng et al. demonstrate critical interactions between adipose tissue metabolism and the immune system, and the active
role of adipose tissue macrophage (ATM) polarization in the progression of obesity. Lean individuals in a non-inflammatory state
maintain a relatively low percentage (~10–15%) of resident ATMs
(76). Healthy adipose tissue consists of uniformly distributed alternatively activated M2 macrophages, expressing Arg1 and the cell
surface antigens CD206 and CD301 (Figure 3) (77–79). The M2
polarized state of the ATM population in healthy adipose tissue
is maintained by eosinophils (80), which secrete IL-4 (Figure 3)
(81, 82). Polarized M2 ATMs secrete interleukin-10 (IL-10), which
regulates glucose homeostasis within adipose tissue and systemic
tissues including muscle (Figure 3) (83). In turn adipose resident invariant natural killer T cells (iNKT), expressing nuclear
factor interleukin 3 regulated (E4BP4), induces the M2 phenotype through secretion of IL-10 (84). Healthy adipocytes, in turn,
secrete anti-inflammatory adipokines such as adiponectin, a hormone that acts synergistically with IL-4 to exert anti-inflammatory
effects through the activation of AMP kinase (AMPK) and signal
transducer and activator of transcription 6 (STAT6), respectively
(78, 85, 86). The alternative M2 state is further sustained by
the transcriptional regulators peroxisome proliferator-activated
receptor-δ (PPARδ), PPARγ, and Krüppel-like factor 4 (KLF4) (82,
87–89).
The intimate interaction between adipocytes and ATMs may
reflect a common origin of these two cell types. Studies have
shown not only cross-talk between ATMs and adipocytes, but a
great deal of plasticity between these two lineages compartments
(90, 91). Furthermore, while still controversial (92) several studies have suggested that some adipocytes and adipose progenitors
may be hematopoietic stem cell-derived (93, 94). In addition,
Coussin et al. have shown that hematopoietic stem cells that can
reconstitute lethally irradiated recipient mice exist within adipose
tissue (95), and more recent studies have supported the concept
that adipose tissue is an extramedullary source of hematopoietic
stem/progenitor cells (96, 97). Yet other studies have demonstrated the phagocytic capacity of pre-adipocytes and adipocytes
suggesting they can adopt macrophage-like functions (85, 98).
Interestingly, a recent study by Eto et al. has identified a novel
subpopulation of ATMs present in perivascular regions of adipose tissue (99). This population of ATMs is characterized by the
expression of CD206 and the stem cell marker CD34. The investigators further demonstrated that these cells can differentiate into
the adipogenic lineage in vitro, suggesting the possibility that this
subpopulation of ATMs might be a source of adipocytes in vivo.
Whether these cells are hematopoietic-derived or whether they
are precursors of the more abundant CD206+ CD34− ATMs is
unclear.
In the late 2000s, Lumeng et al. identified adipose tissue inflammation as an important early event in the development of obesity
related complications (82). At the onset of environmental and

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FIGURE 3 | Homeostatic regulation of ATM microenvironment.
Healthy adipose tissue contains a relatively low and uniformly dispersed
population of alternatively activated M2 macrophages, expressing cell
surface antigens CD206 and CD301. The M2 polarized state is maintained

metabolic perturbation, such as chronic exposure to a high-fat
diet (HFD), adipose tissue undergoes vast biochemical and morphological remodeling in part facilitated by infiltrating monocytes
that then differentiate into ATMs. An excess of nutrients stimulates adipose tissue hyperplasia and hypertrophy as individual
adipocytes enlarge with excess triglycerides (TAGs) and simultaneously undergo heightened levels of cellular stress imparted
by endoplasmic reticular (ER) stress, hypoxia, release of excess
free fatty acids (FFAs), increased reactive oxygen species (ROS)
production, and adipocyte necrosis (77, 82, 100). The “stressed
adipocytes” create an environmental milieu that promotes a more
pro-inflammatory state exemplified with an increase in local IL-6,
TNFα, and IL-1 production (101). Early adipocyte death is facilitated by ATMs in an effort to maintain homeostasis, followed by
the development of a new range of pre-adipocytes (77, 102). Further studies demonstrated that adipocyte necrosis is not only an
early event in diet-induced obesity (DIO), but it is a pivotal event,
which results in the release of chemokines that attract circulating monocytes to the site of inflammation (101, 103). It is well
established that throughout the course of adipose tissue inflammation, the resident macrophage population increases from ~10
to 50–60% associated with a parallel increase in F4/80+ CD11c+
inflammatory macrophages (78, 82). Interestingly, Kosteli et al.

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Macrophage ontogeny and polarization

by eosinophil and adipocyte derived adipokine secretions, IL-4, and
adiponectin, respectively. M2 ATMs maintain a homeostatic adipose
milieu with IL-10 secretions, which in turn regulate glucose homeostasis
within systemic tissues.

demonstrated that weight loss also leads to a rapid recruitment
and accumulation of ATM’s in white adipose tissue (WAT) (104).
ATM accumulation occurs with lipolysis-induced release of excessive FFAs and subsequent clearance of lipolysis byproducts by WAT
macrophages. Contrary to DIO ATM accumulation, weight loss
induced ATM recruitment does not exacerbate the existing inflammatory state and ATM populations diminish with the declining
rate of lipolysis (105).
At the onset of a physiological imbalance in the adipose
niche, such as weight gain, the uniformly dispersed small population of resident macrophages exhibits an alternatively activated
CD206+ CD301+ Arg1+ M2 phenotype (79, 82, 87). The ensuing pro-inflammatory milieu created by stressed adipocytes and
necrosis attracts circulating monocytes to the site of inflammation as early as 10 weeks from the onset of HFD introduction. Circulating Ly6C+ /chemokine receptor-2 (CCR2)+ monocytes are recruited and differentiate into the classically activated
CD11c+ CD11b+ F4/80+ M1 ATMs expressing high levels of iNOS
and TNFα (78, 106). Cinti et al. demonstrated that the differentiated M1 ATMs are exclusively localized in epididymal white adipose tissue (eWAT) and not inguinal white adipose tissue (iWAT),
more specifically within “crown-like structures (CLS)” composed
of necrotic adipocytes (107). The authors incorporated a HSL−/−

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(hormone sensitive lipase KO) model to show that adipocyte
hypertrophy itself triggers a pro-inflammatory microenvironment
that facilitates infiltration of monocytes (107). The M1 monocytederived macrophages are recruited and stabilized by CCR2 and
its ligand CCL2 in addition to other factors such as osteopontin
and fatty acids (FAs) (78, 106). CCL2 loss of function as well
as CCR2 gain of function or overexpression studies have collectively illustrated the fundamental role of CCR2 and CCL2 in
recruitment and differentiation of macrophages with an M1 phenotype (78, 85). However, Amano et al. recently demonstrated
that local proliferation of ATMs contributes significantly to tissue
ATM accumulation. They further demonstrated that this in situ
proliferation is driven by monocyte chemotactic protein (MCP-1)
(108). These studies highlight the contribution of both local proliferation and infiltrating monocytes to the accumulation of ATMs
in an obesity induced inflammatory state (108, 109).
While the observed ATM increase associated with obesity is
predominantly attributed to macrophages exhibiting an M1 phenotype, Shaul et al. recently suggested that there exists an M1/M2
hybrid ATM population and that the increase in ATMs with obesity
exhibit more of this mixed phenotype. The study noted an increase
in both M1 and M2 gene transcripts in a mouse DIO model
of chronic inflammation suggesting that perhaps both populations may influence the progression of obesity-associated chronic
inflammation (103). However, despite transcriptional increases in
both M1 and M2 genes, further studies by the authors demonstrated an overall decrease in the M2 populations in the DIO model
as exhibited by flow cytometry analysis of eWAT cells expressing macrophage galactose type-C lectin-1 (MGL1, also known as
CD301). It is not clear whether this intermediate population of
ATMs represents a truly separate population of cells or whether
these ATMs represent a transitional population, resulting from the
inherent plasticity of ATM phenotypes in vivo.
In healthy adipose tissue, tissue remodeling is accompanied
by angiogenesis to maintain oxygen supply and critical nutrients
to promote adipose tissue homeostasis. However, under pathological conditions, the existing adipocytes enlarge, angiogenesis
is limited, and tissue hypoxia ensues, resulting in the recruitment of inflammatory cells. Angiogenesis is induced by vascular
endothelial factor-A (VEGF-A) stimulation of vascular endothelial
growth factor receptor-2 (VEGFR-2) (110, 111). When VEGFR-2 is
blocked early in the onset of obesity, the development of metabolic
disorders is elevated suggesting that expanding vascularization can
promote diet-induced disease (112). However, Elias et al. demonstrated that overexpression of VEGF-A in adipose tissue in a DIO
model promotes the maintenance of a larger M2 population and
attenuates adipose tissue inflammation (113), coinciding with an
increase in local blood flow and decreased presence of necrotic
CLS. The authors further conclude that the correlative increase in
VEGF and M2 macrophage populations is likely due to recruitment of M2 macrophages to sites of inflammation in adipose
tissue rather than promoting a phenotypic switch between M1 and
M2 macrophage populations. Recent studies also demonstrate an
increase in brown adipose tissue thermogenesis as a response to
cold, particularly in the presence of VEGF overexpression (113,
114). Interestingly, pivotal studies in by the Chawla lab demonstrated that the absence of alternatively activated M2 macrophages

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results in impaired metabolic adaptation to cold, while administration of IL-4 promotes adaptation in a macrophage-dependent
manner (115), suggesting that the increased thermogenesis in
response to cold in the presence of VEGF could also be mediated,
at least in part, by M2 macrophages.
Therapeutic strategies in obesity and obesity-associated disease
may lie in understanding the molecular underpinnings of tissueresident macrophage heterogeneity. For instance, loss of KLF4
function has been associated with a loss of wound healing capacities under inflammatory conditions suggesting an important role
for KLF4 in maintaining an M2 phenotype in vivo (87). In addition
to KLF4, differential expression of PPARδ and PPARγ in current
experimental models, including HFD-induced obesity, illustrate
a critical role for activated PPARs in directing monocytic infiltration and polarization of macrophages toward the alternative
state in adipose tissue and liver (116, 117). Genetic deletion of
PPARδ and/or PPARγ in macrophages directs the differentiation of
monocytes toward a pro-inflammatory M1 phenotype (88, 118).
Conversely, agonist mediated PPAR-activation stimulates monocytic differentiation into an alternately activated M2 phenotype
(119). These agonists include synthetic anti-diabetic agents such
as Thiazolidinediones (TZD) or natural agents such as abscisic
acid (ABA) (116, 119, 120). Recent studies by Satoh et al. have
demonstrated an essential role for the pseudokinase, Trib1, in
the development of M2-like macrophage populations. Mice lacking Trib1 in the hematopoietic cell compartment exhibit a severe
reduction in M2-like macrophages in a number of tissues under
homeostatic conditions (121). In addition, these mice also exhibit
a significant reduction in eosinophils, suggesting that the defect
in M2 polarization could be, in part, secondary to diminished
levels of eosinophil-derived IL-4 under homeostatic conditions.
Importantly, these mice develop hypertriglyceridemia and insulin
resistance when maintained on a HFD, associated with increased
expression of pro-inflammatory cytokines (121, 122).

KUPFFER CELLS (LIVER-RESIDENT MACROPHAGES)
One of the largest resident populations of macrophages exists in
the liver. These YS macrophages, termed Kupffer cells, are positioned in the liver sinusoids where they efficiently clear microbes
and apoptotic cells from the portal circulation (Figure 4) (123).
While the proliferative capacity of hepatic macrophages remains
controversial, recent studies have suggested that Kupffer cells are
established prenatally and are maintained into adulthood independent of replenishment from blood monocytes in a manner
dependent on M-CSF and GM-CSF (9). Kupffer cells also play
essential roles in tissue homeostasis, tissue remodeling, and regulation of metabolic function in the liver (Figure 4). Due to
their constant exposure to blood-borne food antigens and bacterial products of commensal intestinal flora, these cells are
also required to prevent the onset of inflammation in response
to these non-pathogenic stimuli (124). In contrast, monocytederived macrophages in the liver play dual roles in promoting
both tissue inflammation and repair. These infiltrating cells are
initially CD11b+ F4/80+ and eventually differentiate into more
mature CD11blo F4/80hi macrophages.
Following acute hepatic injury induced by acetaminophen,
there is a large influx of monocyte-derived macrophages induced

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FIGURE 4 | The dual role of M1/M2 Kupffer cells in NAFLD. Kupffer
cells play a pivotal role in host defense where they routinely clear
microbes and apoptotic bodies from portal circulation. During
diet-induced liver injury, tissue-resident macrophages exhibiting a
classically activated M1 phenotype predominate and secrete cytokines

by CCL2, accompanied by a significant decrease in the resident
population of Kupffer cells (125–127). This influx is blocked in
CCR2-deficient mice and these mice exhibit a delay in tissue recovery indicating that the infiltrating population of macrophages
plays an important role in promoting liver repair following an
acute insult (126, 127). However, when both infiltrating hepatic
macrophages and resident Kupffer cells are depleted, tissue recovery is dramatically impaired (126, 128). These mice exhibited
impaired clearance of necrotic cells and a defect in the restoration
of the liver microvasculature (126, 128). During the resolution
phase of acute inflammation, the infiltrating monocyte-derived
cells differentiate into the more mature reparative macrophages
and the resident Kupffer cell population is restored. Genomic
analysis of these co-existing populations of cells demonstrated
that, while the recovered Kupffer cell population was similar to
resident Kupffer cells before injury, these cells were distinct from
the resolution phase monocyte-derived liver macrophage population, suggesting that these populations have distinct functions
in liver recovery from acute inflammation and that the infiltrating monocyte-derived macrophages do not replace the resident
Kupffer cells (126).
Non-alcoholic fatty liver disease is the most common liver disease in the western world, the prevalence of which has increased in

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Macrophage ontogeny and polarization

that can alter hepatocyte lipid metabolism and induce MCP-1
dependent recruitment of monocytes into the liver. In turn, these
infiltrating cells facilitate the development and progression of NAFLD.
Restrained induction of M1 Kupffer cell apoptosis further perpetuates
liver inflammation.

parallel to the increased incidence of obesity (129, 130). This clinicopathological condition is characterized by lipid accumulation
in hepatocytes and ranges from the non-progressive form termed
steatosis to non-alcoholic steatohepatitis (NASH), the progressive
form that is prone to the development of cirrhosis, liver cancer,
and liver failure (130, 131). In response to obesity, Kupffer cells are
engaged by cytokines, adipokines, and FAs secreted from adipose
tissue promoting a polarized M1 phenotype (132). In addition,
the release of FFAs into the liver by hypertrophic adipocytes leads
to hepatocyte ER stress and ROS production (132). Hepatocyte
death triggered by these responses results in the release of danger
associated molecular patterns (DAMPs) that further promote liver
macrophage polarization (123, 132).
In hepatic steatosis, pro-inflammatory cytokines produced by
M1 activated macrophages induce hepatic cholesterologenesis and
increase TAG production, resulting in discordant regulation of
lipid metabolism and homeostasis (Figure 4) (133–136). Sterol
regulatory element-binding proteins (SREBPs) directly activate
genes that mediate the synthesis and uptake of cholesterol and
FAs (137). Elevation of SREBPs often underlies the pathogenesis
of insulin resistance and type 2 diabetes (138, 139). In addition to
their initial propagation of fatty liver disease, activated M1 Kupffer cells trigger the more severe form of non-alcoholic fatty liver

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disease (NAFLD), NASH. A recent study demonstrated that TNFα
produced by M1 hepatic macrophages increased intrahepatic
expression of MCP-1 (Figure 4), a major chemokine responsible
for CCR2-dependent recruitment of Ly6Chi monocytes (135). The
vast infiltration of Ly6Chi monocytes further perturbs liver homeostasis amplifying hepatic inflammatory responses (Figure 4).
The recruited monocyte-derived macrophage population also
plays a key role in promoting fibrogenesis. In mice deficient for
CCR2 or depleted of Kupffer cells, inflammatory cell infiltration,
steatohepatitis, and fibrosis are ameliorated (135).
Progression of NAFLD to NASH occurs in a subset of patients
with fatty liver disease, and this progression is largely linked to
the activation of toll like receptor 4 (TLR4) by both exogenous
and endogenous ligands. Bacteria and bacterial products, such as
LPS, which translocate from the gut can activate TLR4 signaling to
induce NASH (140, 141). Furthermore, endogenous high mobility group box-1 (HMGB-1) and FFAs have also been shown to be
endogenous ligands for TLR4 (142, 143). A number of recent studies have also demonstrated a crucial link between the composition
of gut microbiota, obesity, and the development of NASH (144).
CD14 promotes the activation of TLR4 by promoting the recruitment of LPS to the TLR4 signaling complex (145). Interestingly,
Imajo et al. demonstrated that the increased levels of leptin associated with obesity promotes the enhanced expression of CD14
in Kupffer cells, resulting in increased sensitivity of the Kupffer
cells to low doses of LPS (146). Translocated bacterial DNA has
also been shown to promote the development of NASH through
engagement of TLR9 in Kupffer cells, which stimulates the production of interleukin-1β (IL-1β) (147). IL-1β in turn, promotes
hepatocyte lipid accumulation and death, as well inducing hepatic
stellate cells (HpSCs) to promote fibrosis.
Liver fibrosis is a common feature of chronic liver disease. It
is characterized by accumulation of extracellular matrix (ECM)
resulting from both increased synthesis and decreased degradation
of ECM proteins (148). During progressive liver injury, HpSCderived myofibroblast-like cells predominate and initiate collagen
deposition (148). Extensive evidence demonstrates a complex
interplay between Kupffer cells and HpSCs during hepatic fibrogenesis. In general, alternatively activated M2 macrophages are
involved in tissue remodeling through the production of transforming growth factor beta-1 (TGFβ-1), a potent inducer of the
HpSC fibrotic phenotype (149, 150). However, while arginase
produced by M2 cells during a Th2 response was thought to
play a role in progression of fibrosis, studies by Pesce et al.,
in which arginase is depleted in macrophages, highlight a role
for arginase-expressing macrophages in the resolution of fibrosis (151). Alternatively, in hepatic fibrosis, Ly6Chi inflammatory
monocyte-derived macrophages express higher levels of TGFβ1 and other pro-inflammatory cytokines that activate HpSCs
(152). TGFβ-1 not only promotes the fibrogenic activity of HpSCs
but also enhances expression of tissue inhibitors of metalloproteinases (TIMPs) by HpSCs that block the degradation of ECM.
These inflammatory monocyte-derived macrophages also exhibit
decreased expression of matrix metalloproteinases, thus directly
promoting fibrosis.
Interestingly, a recent report demonstrated a role for a novel
population of monocyte-derived macrophages in the resolution

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of fibrosis. These cells are derived from Ly6Chi monocytes and
exhibit a CD11bhi F4/80int Ly6Clo phenotype. These cells mediate MMP-dependent degradation of the ECM and the expression profile of this population of resolving monocyte-derived
macrophages demonstrates that these cells are distinct from the
typical M1/M2 populations of macrophages (152). More recently,
it has been reported that alternatively activated M2 Kupffer cells
can promote caspase-3-dependent apoptosis of classically activated M1 Kupffer cells, thus providing a protective mechanism
against NAFLD (Figure 4) (153). The M2 mediated apoptosis of
M1 Kupffer cells was shown to be arginase dependent by way of
IL-10 (153). Interestingly, alternatively activated M2 Kupffer cells
limit hepatocyte apoptosis and steatosis in alcohol induced liver
injury through the release of IL-6, a mechanism that may expand
to NAFLD (154).

SUMMARY
Based on recent studies, a unifying theme is beginning to emerge
between the contribution of tissue-resident macrophages and
infiltrating monocyte-derived macrophages in chronic inflammation (Figure 5). (1) Both microglia and Kupffer cells are of
embryonic origin. Microglia are YS, while contribution of yolk
sac and fetal liver has been described for Kupffer cells. However,
because it now appears that the yolk sac progenitors seed the liver
where they continue to contribute to tissue-resident macrophage
populations independent of hematopoietic stem cells, it is likely
that these cells share a common progenitor and are ontologically
indistinguishable. (2) In the brain and liver, resident macrophages
and infiltrating monocyte-derived macrophages play distinct roles
in the progression of inflammation. In the brain, microglia are
required for the initiation of disease, while infiltrating monocytederived macrophages promote demyelination. In the liver, both
resident and infiltrating populations contribute to both inflammation and repair, Kupffer cells are largely responsible for the
development of fibrosis. (3) In the brain and liver, infiltrating
monocytes do not contribute to the resident macrophage pool
rather, during recovery, resident cells are replenished through
local proliferation. (4) In both the brain and liver, there is
evidence that the resident population of macrophages plays a
role in promoting apoptosis and/or clearance of the infiltrating
monocyte-derived macrophage population during the recovery
phase.
However, the relative contribution of resident versus infiltrating populations of macrophages in adipose tissue remains
obscure. While ATMs are generally considered to be hematopoietic stem cells-derived, this assumption is based on studies using
myeloablation. Further fate mapping studies will be needed to
definitively identify the origin of ATMs. In addition, while it
is clear that both infiltration of monocytes and proliferation
of resident macrophage contribute to the massive expansion of
ATMs in obesity, the respective roles of resident versus infiltrating
macrophages in the propagation and/or resolution of inflammation is unclear. Finally, it unknown whether infiltrating monocytederived macrophages in adipose tissue ultimately contribute to the
resident population.
In all cases, the M1/M2 polarization of macrophages in inflammation and disease resolution has been described. However, these

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Dey et al.

Macrophage ontogeny and polarization

Microglia

Kupffer cells

Yolk sac

ATMs
Bloodd

Hepatocytes

Microglia
HpSC
MO ?

Yolk sac

Steady-State

Blood
Kupffer
cells
MO

ATMs

Brain
MO

Blood

Adipocytes

Steatosis

Inflamma#on
Demyelina#on

Fibrosis
Insulin
Resistance
FIGURE 5 | Resident macrophage populations in steady state and
inflammation. Resident microglia and Kupffer cell populations, both yolk
sac-derived, are maintained through self-renewal, unlike macrophages
resident to the adipose tissue, which are thought to originate from circulating

are largely based on studies that do not distinguish between resident macrophage and infiltrating macrophage populations. Given
the emergence of technologies that help to distinguish resident
cells from infiltrating cells, it will be essential to re-visit the M1/M2
paradigm in the context of macrophage ontogeny to determine the
extent of plasticity of individual populations of macrophages.
The recognition that the M1/M2 polarization of macrophages
plays a central role in the progression of chronic inflammation in a
wide range of diseases and that therapeutic approaches to chronic
inflammatory disease could involve the regulation of macrophage
polarization renders these outstanding questions highly significant. Taking into consideration not only the polarization state
of macrophages, but also their ontogeny and phenotypic plasticity, will be central in the development of therapeutic approaches
for the treatment of chronic inflammatory disease aimed at
redirecting macrophage polarization and function.

REFERENCES
1. Gautier EL, Shay T, Miller J, Greter M, Jakubzick C, Ivanov S, et al. Geneexpression profiles and transcriptional regulatory pathways that underlie the
identity and diversity of mouse tissue macrophages. Nat Immunol (2012)
13:1118–28. doi:10.1038/ni.2419

www.frontiersin.org

blood monocytes (MO). Although the progression of tissue specific
pathophysiological insults are dependent on both resident and infiltrating
macrophages, monocyte-derived macrophages do not replenish resident
microglia and Kupffer cell populations.

2. Davies LC, Jenkins SJ, Allen JE, Taylor PR. Tissue-resident macrophages. Nat
Immunol (2013) 14:986–95. doi:10.1038/ni.2705
3. Chow A, Huggins M,Ahmed J, Hashimoto D, Kunisaki Y, Pinho S, et al. CD169+
macrophages provide a niche promoting erythropoiesis under homeostasis,
myeloablation and in JAK2V617F-induced polycythemia vera. Nat Med (2014)
19:429–36. doi:10.1038/nm.3057
4. Haldar M, Kohyama M, So AY-L, Kc W, Wu X, Briseño CG, et al. Hememediated SPI-C induction promotes monocyte differentiation into ironrecycling macrophages. Cell (2014) 156:1223–34. doi:10.1016/j.cell.2014.01.
069
5. Epelman S, Lavine KJ, Randolph GJ. Origin and functions of tissue
macrophages. Immunity (2014) 41:21–35. doi:10.1016/j.immuni.2014.06.013
6. Wynn TA, Chawla A, Pollard JW. Macrophage biology in development, homeostasis and disease. Nature (2013) 496:445–55. doi:10.1038/nature12034
7. Ginhoux F, Greter M, Leboeuf M, Nandi S, See P, Gokhan S, et al. Fate mapping analysis reveals that adult microglia derive from primitive macrophages.
Science (2010) 330:841–5. doi:10.1126/science.1194637
8. Schulz C, Gomez Perdiguero E, Chorro L, Szabo-Rogers H, Cagnard N, Kierdorf K, et al. A lineage of myeloid cells independent of Myb and hematopoietic
stem cells. Science (2012) 336:86–90. doi:10.1126/science.1219179
9. Yona S, Kim KW, Wolf Y, Mildner A, Varol D, Breker M, et al. Fate mapping
reveals origins and dynamics of monocytes and tissue macrophages under
homeostasis. Immunity (2014) 38:79–91. doi:10.1016/j.immuni.2012.12.001
10. Epelman S, Lavine KJ, Beaudin AE, Sojka DK, Carrero JA, Calderon B, et al.
Embryonic and adult-derived resident cardiac macrophages are maintained

January 2015 | Volume 5 | Article 683 | 225

Dey et al.

through distinct mechanisms at steady state and during inflammation. Immunity (2014) 40:91–104. doi:10.1016/j.immuni.2013.11.019
11. Heidt T, Courties G, Dutta P, Sager HB, Sebas M, Iwamoto Y, et al. Differential contribution of monocytes to heart macrophages in steady-state
and after myocardial infarction. Circ Res (2014) 115:284–95. doi:10.1161/
CIRCRESAHA.115.303567
12. Pinto AR, Godwin JW, Rosenthal NA. Macrophages in cardiac homeostasis, injury responses and progenitor cell mobilisation. Stem Cell Res (2014)
13:705–14. doi:10.1016/j.scr.2014.06.004
13. Hashimoto D, Chow A, Noizat C, Teo P, Beth M, Leboeuf M, et al. Tissue resident macrophages self-maintain locally throughout adult life with
minimal contribution from circulating monocytes. Immunity (2014) 38:1–25.
doi:10.1016/j.immuni.2013.04.004
14. Bogunovic M, Ginhoux F, Helft J, Shang L, Greter M, Liu K, et al. Origin of
the lamina propria dendritic cell network Milena. Immunity (2010) 31:513–25.
doi:10.1016/j.immuni.2009.08.010
15. Varol C,Vallon-Eberhard A, Elinav E,Aychek T, Shapira Y, Luche H, et al. Intestinal lamina propria dendritic cell subsets have different origin and functions.
Immunity (2009) 31:502–12. doi:10.1016/j.immuni.2009.06.025
16. Perdiguero EG, Klapproth K, Schulz C, Busch K, Azzoni E, Crozet L, et al.
Tissue-resident macrophages originate from yolk-sac-derived erythro-myeloid
progenitors. Nature (2014). doi:10.1038/nature13989
17. Jenkins SJ, Ruckerl D, Cook PC, Jones LH, Finkelman FD, van Rooijen
N, et al. Local macrophage proliferation, rather than recruitment from the
blood, is a signature of TH2 inflammation. Science (2011) 332:1284–8.
doi:10.1126/science.1204351
18. Jenkins SJ, Ruckerl D, Thomas GD, Hewitson JP, Duncan S, Brombacher F,
et al. IL-4 directly signals tissue-resident macrophages to proliferate beyond
homeostatic levels controlled by CSF-1. J Exp Med (2013) 210:2477–91.
doi:10.1084/jem.20121999
19. Mielcarek M, Kirkorian AY, Hackman RC, Price J, Storer BE, Wood BL,
et al. Langerhans cell homeostasis and turnover after nonmyeloablative and
myeloablative allogeneic hematopoietic cell transplantation. Transplantation
(2014) 98:563–8. doi:10.1097/TP.0000000000000097
20. Cohen HB, Mosser DM. Cardiac macrophages: how to mend a broken heart.
Immunity (2014) 40:3–5. doi:10.1016/j.immuni.2013.12.005
21. Rosas M, Davies LC, Giles PJ, Liao C-T, Kharfan B, Stone TC, et al. The transcription factor Gata6 links tissue macrophage phenotype and proliferative
renewal. Science (2014) 344:645–8. doi:10.1126/science.1251414
22. Okabe Y, Medzhitov R. Tissue-specific signals control reversible program
of localization and functional polarization of macrophages. Cell (2014)
157:832–44. doi:10.1016/j.cell.2014.04.016
23. Schneider C, Nobs SP, Kurrer M, Rehrauer H, Thiele C, Kopf M. Induction of
the nuclear receptor PPAR-γ by the cytokine GM-CSF is critical for the differentiation of fetal monocytes into alveolar macrophages. Nat Immunol (2014)
15:1026–37. doi:10.1038/ni.3005
24. A-gonzalez N, Guillen JA, Gallardo G, Diaz M, Rosa JV, De Hernandez IH, et al.
The nuclear receptor LXRα controls the functional specialization of splenic
macrophages. Nat Immunol (2014) 14:831–9. doi:10.1038/ni.2622
25. Ingersoll MA, Spanbroek R, Lottaz C, Gautier EL, Frankenberger M, Hoffmann R, et al. Comparison of gene expression profiles between human and
mouse monocyte subsets. Blood (2010) 115:e10–9. doi:10.1182/blood-200907-235028
26. Cros J, Cagnard N, Woollard K, Patey N, Zhang S-Y, Senechal B, et al. Human
CD14dim monocytes patrol and sense nucleic acids and viruses via TLR7 and
TLR8 receptors. Immunity (2010) 33:375–86. doi:10.1016/j.immuni.2010.08.
012
27. Jakubzick C, Gautier EL, Gibbings SL, Sojka DK, Schlitzer A, Johnson TE,
et al. Minimal differentiation of classical monocytes as they survey steady-state
tissues and transport antigen to lymph nodes. Immunity (2013) 39:599–610.
doi:10.1016/j.immuni.2013.08.007
28. Mills CD, Thomas AC, Lenz LL, Munder M. Macrophage: SHIP of immunity.
Front Immunol (2014) 5:1–5. doi:10.3389/fimmu.2014.00620
29. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.4049/
jimmunol.164.12.6166
30. Mills CD, Ley K. M1 and M2 macrophages: the chicken and the egg of immunity. J Innate Immune (2014) 6:1716–26. doi:10.1159/000364945

Frontiers in Immunology | Molecular Innate Immunity

Macrophage ontogeny and polarization

31. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental
guidelines. Immunity (2014) 41:14–20. doi:10.1016/j.immuni.2014.06.008
32. Perry VH, Teeling J. Microglia and macrophages of the central nervous system: the contribution of microglia priming and systemic inflammation to
chronic neurodegeneration. Semin Immunopathol (2013) 35:601–12. doi:10.
1007/s00281-013-0382-8
33. Ginhoux F, Jung S. Monocytes and macrophages: developmental pathways and
tissue homeostasis. Nat Rev Immunol (2014) 14:392–404. doi:10.1038/nri3671
34. Kierdorf K, Erny D, Goldmann T, Sander V, Schulz C, Perdiguero EG,
et al. Microglia emerge from erythromyeloid precursors via Pu.1- and
Irf8-dependent pathways. Nat Neurosci (2013) 16:273–80. doi:10.1038/nn.
3318
35. Greter M, Lelios I, Pelczar P, Hoeffel G, Price J, Leboeuf M, et al. Stromaderived interleukin-34 controls the development and maintenance of langerhans cells and the maintenance of microglia. Immunity (2012) 37:1050–60.
doi:10.1016/j.immuni.2012.11.001
36. Wang Y, Szretter KJ, Vermi W, Gilfillan S, Rossini C, Cella M, et al. IL-34 is a
tissue-restricted ligand of CSF1R required for the development of Langerhans
cells and microglia. Nat Immunol (2012) 13:753–60. doi:10.1038/ni.2360
37. Miron VE, Franklin RJM. Macrophages and CNS remyelination. J Neurochem
(2014) 130:165–71. doi:10.1111/jnc.12705
38. Ajami B, Bennett JL, Krieger C, Tetzlaff W, Rossi FMV. Local self-renewal can
sustain CNS microglia maintenance and function throughout adult life. Nat
Neurosci (2007) 10:1538–43. doi:10.1038/nn2014
39. Olah M, Biber K, Vinet J, Boddeke HWGM. Microglia phenotype diversity. CNS Neurol Disord Drug Targets (2011) 10:108–18. doi:10.2174/
187152711794488575
40. Eggen BJL, Raj D, Hanisch U-K, Boddeke HWGM. Microglial phenotype and
adaptation. J Neuroimmune Pharmacol (2013) 8:807–23. doi:10.1007/s11481013-9490-4
41. Kettenmann H, Hanisch U, Noda M, Verkhratsky A. Physiology of microglia.
Physiol Rev (2011) 91:461–553. doi:10.1152/physrev.00011.2010
42. Boche D, Perry VH, Nicoll JAR. Review: activation patterns of microglia and
their identification in the human brain. Neuropathal Appl Neurobiol (2013)
39:3–18. doi:10.1111/nan.12011
43. Prinz M, Tay TL, Wolf Y, Jung S. Microglia: unique and common features with other tissue macrophages. Acta Neuropathol (2014) 128:319–31.
doi:10.1007/s00401-014-1267-1
44. Butovsky O, Jedrychowski MP, Moore CS, Cialic R, Amanda J, Gabriely G,
et al. Identification of a unique TGF-β dependent molecular and functional
signature in microglia. Nat Neurosci (2014) 17:131–43. doi:10.1038/nn.3599
45. Prinz M, Priller J. Microglia and brain macrophages in the molecular age:
from origin to neuropsychiatric disease. Nat Rev Neurosci (2014) 15:300–12.
doi:10.1038/nrn3722
46. Landsman L, Bar-On L, Zernecke A, Kim K-W, Krauthgamer R, Shagdarsuren
E, et al. CX3CR1 is required for monocyte homeostasis and atherogenesis by
promoting cell survival. Blood (2009) 113:963–72. doi:10.1182/blood-200807-170787
47. Donnelly DJ, Longbrake EE, Shawler TM, Kigerl KA, Lai W, Tovar CA, et al.
Deficient CX3CR1 signaling promotes recovery after mouse spinal cord injury
by limiting the recruitment and activation of Ly6Clo/iNOS+ macrophages. J
Neurosci (2011) 31:9910–22. doi:10.1523/JNEUROSCI.2114-11.2011
48. Wolf Y, Yona S, Kim K-W, Jung S. Microglia, seen from the CX3CR1 angle.
Front Cell Neurosci (2013) 7:26. doi:10.3389/fncel.2013.00026
49. Billadeau DD, Leibson PJ. ITAMs versus ITIMs: striking a balance during cell
regulation. J Clin Invest (2002) 109:161–8. doi:10.1172/JCI200214843
50. Linnartz B, Neumann H. Microglial activatory (immunoreceptor tyrosinebased activation motif)- and inhibitory (immunoreceptor tyrosine-based inhibition motif)-signaling receptors for recognition of the neuronal glycocalyx.
Glia (2013) 61:37–46. doi:10.1002/glia.22359
51. Lambertsen KL, Deierborg T, Gregersen R, Clausen BH, Wirenfeldt M, Nielsen
HH, et al. Differences in origin of reactive microglia in bone marrow chimeric
mouse and rat after transient global ischemia. J Neuropathol Exp Neurol (2011)
70:481–94. doi:10.1097/NEN.0b013e31821db3aa
52. Bechmann I, Priller J, Kovac A, Böntert M, Wehner T, Klett FF, et al. Immune
surveillance of mouse brain perivascular spaces by blood-borne macrophages.
Eur J Neurosci (2001) 14:1651–8. doi:10.1046/j.0953-816x.2001.01793.x

January 2015 | Volume 5 | Article 683 | 226

Dey et al.

53. Cherry JD, Olschowka JA, O’Banion MK. Neuroinflammation and M2
microglia: the good, the bad, and the inflamed. J Neuroinflammation (2014)
11:98. doi:10.1186/1742-2094-11-98
54. Varnum MM, Ikezu T. The classification of microglial activation
phenotypes on neurodegeneration and regeneration in Alzheimer’s disease
brain. Arch Immunol Ther Exp (2012) 60:251–66. doi:10.1007/s00005-0120181-2
55. Amor S, Peferoen LAN, Vogel DYS, Breur M, van der Valk P, Baker D, et al.
Inflammation in neurodegenerative diseases – an update. Immunology (2014)
142:151–66. doi:10.1111/imm.12233
56. Walker FR, Beynon SB, Jones KA, Zhao Z, Kongsui R, Cairns M, et al. Dynamic
structural remodelling of microglia in health and disease: a review of the
models, the signals and the mechanisms. Brain Behav Immun (2014) 37:1–14.
doi:10.1016/j.bbi.2013.12.010
57. Eggen BJL, Raj D, Hanisch U. Microglial phenotype and adaptation. J Neuroimmune Pharmacol (2013) 8:807–23. doi:10.1007/s11481-013-9490-4
58. Crain JM, Nikodemova M, Watters JJ. Microglia express distinct M1 and M2
phenotypic markers in the postnatal and adult central nervous system in male
and female mice. J Neurosci Res (2013) 91:1143–51. doi:10.1002/jnr.23242
59. Streit WJ, Conde JR. Microglia in the aging brain. J Neuropathol Exp Neurol
(2006) 65:199–203. doi:10.1097/01.jnen.0000202887.22082.63
60. Streit WJ, Xue Q-S. Microglial senescence. CNS Neurol Disord Drug Targets
(2013) 12:763–7. doi:10.2174/18715273113126660176
61. Rezai-Zadeh K, Gate D, Town T. CNS infiltration of peripheral immune
cells: d-day for neurodegenerative disease? J Neuroimmune Pharmacol (2009)
4:462–75. doi:10.1007/s11481-009-9166-2
62. Jiang Z, Jiang JX, Zhang G-X. Macrophages: a double-edged sword in experimental autoimmune encephalomyelitis. Immunol Lett (2014) 160:17–22.
doi:10.1016/j.imlet.2014.03.006
63. Buckman LB, Hasty AH, Flaherty DK, Buckman CT, Thompson MM, Matlock BK, et al. Obesity induced by a high-fat diet is associated with increased
immune cell entry into the central nervous system. Brain Behav Immun (2014)
35:33–42. doi:10.1016/j.bbi.2013.06.007
64. Tucsek Z, Toth P, Sosnowska D, Gautam T, Mitschelen M, Koller A, et al. Obesity in aging exacerbates blood-brain barrier disruption, neuroinflammation,
and oxidative stress in the mouse hippocampus: effects on expression of genes
involved in beta-amyloid generation and Alzheimer’s disease. J Gerontol A Biol
Sci Med Sci (2013) 69:1212–26. doi:10.1093/gerona/glt177
65. Yamasaki R, Lu H, Butovsky O, Ohno N, Rietsch AM, Cialic R, et al. Differential
roles of microglia and monocytes in the inflamed central nervous system. J Exp
Med (2014) 211:1533–49. doi:10.1084/jem.20132477
66. Jung S, Schwartz M. Non-identical twins – microglia and monocyte-derived
macrophages in acute injury and autoimmune inflammation. Front Immunol
(2012) 3:89. doi:10.3389/fimmu.2012.00089
67. Vainchtein ID, Vinet J, Brouwer N, Brendecke S, Biagini G, Biber K, et al. In
acute experimental autoimmune encephalomyelitis, infiltrating macrophages
are immune activated, whereas microglia remain immune suppressed. Glia
(2014) 62:1724–35. doi:10.1002/glia.22711
68. Ajami B, Bennett JL, Krieger C, McNagny KM, Rossi FMV. Infiltrating monocytes trigger EAE progression, but do not contribute to the resident microglia
pool. Nat Neurosci (2011) 14:1142–9. doi:10.1038/nn.2887
69. Jimenez S, Baglietto-Vargas D, Caballero C, Moreno-Gonzalez I, Torres
M, Sanchez-Varo R, et al. Inflammatory response in the hippocampus of
PS1M146L/APP751SL mouse model of Alzheimer’s disease: age-dependent
switch in the microglial phenotype from alternative to classic. J Neurosci (2008)
28:11650–61. doi:10.1523/JNEUROSCI.3024-08.2008
70. Yi C-X, Tschöp MH, Woods SC, Hofmann SM. High-fat-diet exposure
induces IgG accumulation in hypothalamic microglia. Dis Model Mech (2012)
5:686–90. doi:10.1242/dmm.009464
71. Goldmann T, Wieghofer P, Müller PF, Wolf Y, Varol D, Yona S, et al. A new type
of microglia gene targeting shows TAK1 to be pivotal in CNS autoimmune
inflammation. Nat Neurosci (2013) 16:1618–26. doi:10.1038/nn.3531
72. Van Ham TJ, Brady CA, Kalicharan RD, Oosterhof N, Kuipers J, Veenstra-Algra
A, et al. Intravital correlated microscopy reveals differential macrophage and
microglial dynamics during resolution of neuroinflammation. Dis Model Mech
(2014) 7:857–69. doi:10.1242/dmm.014886
73. De Heredia FP, Gómez-Martínez S, Marcos A. Obesity, inflammation
and the immune system. Proc Nutr Soc (2012) 71:332–8. doi:10.1017/
S0029665112000092

www.frontiersin.org

Macrophage ontogeny and polarization

74. Hotamisligil GS, Shargill S, Spiegelman BM. Adipose expression of tumor
necrosis factor – a: direct role in obesity-linked insulin resistance. Science
(1993) 259:87–91. doi:10.1126/science.7678183
75. Spiegelman BM, Choy L, Hotamisligil GS, Graves RA, Tontonoz P. Regulation of adipocyte gene expression in differentiation and syndromes of obesity/diabetes. J Biol Chem (1993) 268:6823–6.
76. Schipper HS, Prakken B, Kalkhoven E, Boes M. Adipose tissue-resident immune
cells: key players in immunometabolism. Trends Endocrinol Metab (2012)
23:407–15. doi:10.1016/j.tem.2012.05.011
77. Wernstedt Asterholm I, Tao C, Morley TS, Wang QA, Delgado-Lopez F, Wang
ZV, et al. Adipocyte inflammation is essential for healthy adipose tissue expansion and remodeling. Cell Metab (2014) 20:103–18. doi:10.1016/j.cmet.2014.
05.005
78. Chawla A, Nguyen KD, Goh YPS. Macrophage-mediated inflammation in
metabolic disease. Nat Rev Immunol (2011) 11:738–49. doi:10.1038/nri3071
79. Lumeng CN, DelProposto JB, Westcott DJ, Saltiel AR. Phenotypic switching of
adipose tissue macrophages with obesity is generated by spatiotemporal differences in macrophage subtypes. Diabetes (2008) 57:3239–46. doi:10.2337/
db08-0872
80. Wu D, Molofsky AB, Liang HE, Ricardo-Gonzalez RR, Jouihan HA, Bando JK,
et al. Eosinophils sustain adipose alternatively activated macrophages associated with glucose homeostasis. Science (2011) 332:243–7. doi:10.1126/science.
1201475
81. Prajeeth CK, Löhr K, Floess S, Zimmermann J, Ulrich R, Gudi V, et al. Effector
molecules released by Th1 but not Th17 cells drive an M1 response in microglia.
Brain Behav Immun (2014) 37:248–59. doi:10.1016/j.bbi.2014.01.001
82. Lumeng CN, Bodzin JL, Saltiel AR. Obesity induces a phenotypic switch
in adipose tissue macrophage polarization. J Clin Invest (2007) 117:175–84.
doi:10.1172/JCI29881
83. Nagareddy PR, Kraakman M, Masters SL, Stirzaker RA, Gorman DJ, Grant RW,
et al. Adipose tissue macrophages promote myelopoiesis and monocytosis in
obesity. Cell Metab (2014) 19:821–35. doi:10.1016/j.cmet.2014.03.029
84. Lynch L, Michelet X, Zhang S, Brennan PJ, Moseman A, Lester C, et al. Regulatory iNKT cells lack expression of the transcription factor PLZF and control
the homeostasis of Treg cells and macrophages in adipose tissue. Nat Immunol
(2014) 16:85–95. doi:10.1038/ni.3047
85. Gustafson B, Hammarstedt A, Andersson CX, Smith U. Inflamed adipose tissue:
a culprit underlying the metabolic syndrome and atherosclerosis. Arterioscler
Thromb Vasc Biol (2007) 27:2276–83. doi:10.1161/ATVBAHA.107.147835
86. Steinberg GR, Schertzer JD. AMPK promotes macrophage fatty acid oxidative
metabolism to mitigate inflammation: implications for diabetes and cardiovascular disease. Immunol Cell Biol (2014) 92:340–5. doi:10.1038/icb.2014.11
87. Liao X, Sharma N, Kapadia F, Zhou G, Lu Y, Hong H, et al. Krüppel-like factor 4 regulates macrophage polarization. J Clin Invest (2011) 121:2736–49.
doi:10.1172/JCI45444
88. Chawla A. Control of macrophage activation and function by PPARs. Circ Res
(2010) 106:1559–69. doi:10.1161/CIRCRESAHA.110.216523
89. Fernández-Quintela A, Churruca I, Portillo MP. The role of dietary fat in adipose tissue metabolism. Public Health Nutr (2007) 10:1126–31. doi:10.1017/
S1368980007000602
90. Charrière G, Cousin B, Arnaud E, André M, Bacou F, Penicaud L, et al.
Preadipocyte conversion to macrophage. Evidence of plasticity. J Biol Chem
(2003) 278:9850–5. doi:10.1074/jbc.M210811200
91. Chazenbalk G, Bertolotto C, Heneidi S, Jumabay M, Trivax B, Aronowitz J,
et al. Novel pathway of adipogenesis through cross-talk between adipose tissue macrophages, adipose stem cells and adipocytes: evidence of cell plasticity.
PLoS One (2011) 6:e17834. doi:10.1371/journal.pone.0017834
92. Koh YJ, Kang S, Lee HJ, Choi T-S, Lee HS, Cho C-H, et al. Bone marrow-derived
circulating progenitor cells fail to transdifferentiate into adipocytes in adult adipose tissues in mice. J Clin Invest (2007) 117:3684–95. doi:10.1172/JCI32504
93. Sera Y, Larue AC, Moussa O, Mehrotra M, James D, Williams CR, et al.
Hematpoietic stem cell origin of adipocytes. Exp Hematol (2010) 37:1108–20.
doi:10.1016/j.exphem.2009.06.008
94. Crossno JT, Majka SM, Grazia T, Gill RG, Klemm DJ. Rosiglitazone promotes
development of a novel adipocyte population from bone marrow-derived
circulating progenitor cells. J Clin Invest (2006) 116:3220–8. doi:10.1172/
JCI28510
95. Luche E, Cousin B, Garidou L, Serino M, Waget A, Barreau C, et al. Metabolic endotoxemia directly increases the proliferation of adipocyte precursors

January 2015 | Volume 5 | Article 683 | 227

Dey et al.

96.

97.

98.
99.

100.

101.

102.
103.

104.

105.
106.

107.

108.

109.

110.

111.

112.
113.

114.

115.

116.

at the onset of metabolic diseases through a CD14-dependent mechanism. Mol
Metab (2013) 2:281–91. doi:10.1016/j.molmet.2013.06.005
Poglio S, De Toni-Costes F, Arnaud E, Laharrague P, Espinosa E, Casteilla L,
et al. Adipose tissue as a dedicated reservoir of functional mast cell progenitors.
Stem Cells (2010) 28:2065–72. doi:10.1002/stem.523
Han J, Koh YJ, Moon HR, Ryoo HG, Cho C-H, Kim I, et al. Adipose tissue is
an extramedullary reservoir for functional hematopoietic stem and progenitor
cells. Blood (2010) 115:957–64. doi:10.1182/blood-2009-05-219923
Cousin B, Munoz O, Andre M, Fontanilles AM, Dani C, Cousin JL, et al. A role
for preadipocytes as macrophage-like cells. FASEB J (1999) 13:305–12.
Eto H, Ishimine H, Kinoshita K, Watanabe-Susaki K, Kato H, Doi K, et al.
Characterization of human adipose tissue-resident hematopoietic cell populations reveals a novel macrophage subpopulation with CD34 expression and
mesenchymal multipotency. Stem Cells Dev (2013) 22:985–97. doi:10.1089/
scd.2012.0442
Manteiga S, Choi K, Jayaraman A, Lee K. Systems biology of adipose tissue
metabolism: regulation of growth, signaling and inflammation. Wiley Interdiscip Rev Syst Biol Med (2013) 5:425–47. doi:10.1002/wsbm.1213
Strissel KJ, Stancheva Z, Miyoshi H, Perfield JW, Defuria J, Jick Z, et al.
Adipocyte death, adipose tissue remodeling, and obesity complications. Diabetes (2007) 56:2910–8. doi:10.2337/db07-0767
Lee Y, Thacker RI, Hall BE, Kong R, Granneman JG. Exploring the activated
adipogenic niche. Cell Cycle (2014) 13:184–90. doi:10.4161/cc.27647
Shaul ME, Bennett G, Strissel KJ, Greenberg AS, Obin MS. Dynamic, M2-Like
remodeling phenotypes of CD11c+ adipose tissue macrophages during highfat diet-induced obesity in mice. Diabetes (2010) 59:1171–81. doi:10.2337/
db09-1402
Kosteli A, Sugaru E, Haemmerle G, Martin JF, Lei J, Zechner R, et al. Weight loss
and lipolysis promote a dynamic immune response in murine adipose tissue.
J Clin Invest (2010) 120:3466–79. doi:10.1172/JCI42845
Red Eagle A, Chawla A. In obesity and weight loss, all roads lead to the mighty
macrophage. J Clin Invest (2010) 120:3437–40. doi:10.1172/JCI44721
Chávez-Sánchez L, Espinosa-Luna JE, Chávez-Rueda K, Legorreta-Haquet MV,
Montoya-Díaz E, Blanco-Favela F. Innate immune system cells in atherosclerosis. Arch Med Res (2014) 45:1–14. doi:10.1016/j.arcmed.2013.11.007
Cinti S, Mitchell G, Barbatelli G, Murano I, Ceresi E, Faloia E, et al. Adipocyte
death defines macrophage localization and function in adipose tissue of obese
mice and humans. J Lipid Res (2005) 46:2347–55. doi:10.1194/jlr.M500294JLR200
Amano SU, Cohen JL, Vangala P, Tencerova M, Nicoloro SM, Yawe JC, et al.
Local proliferation of macrophages contributes to obesity-associated adipose
tissue inflammation. Cell Metab (2014) 19:162–71. doi:10.1016/j.cmet.2013.
11.017
Haase J, Weyer U, Immig K, Klöting N, Blüher M, Eilers J, et al. Local proliferation of macrophages in adipose tissue during obesity-induced inflammation.
Diabetologia (2014) 57:562–71. doi:10.1007/s00125-013-3139-y
Shibuya M. Vascular endothelial growth factor and its receptor system: physiological functions in angiogenesis and pathological roles in various diseases.
J Biochem (2013) 153:13–9. doi:10.1093/jb/mvs136
Moens S, Goveia J, Stapor PC, Cantelmo AR, Carmeliet P. The multifaceted
activity of VEGF in angiogenesis – implications for therapy responses. Cytokine
Growth Factor Rev (2014) 25:473–82. doi:10.1016/j.cytogfr.2014.07.009
Sun K, Kusminski CM, Scherer PE. Adipose tissue remodeling and obesity.
J Clin Invest (2011) 121:2094–101. doi:10.1172/JCI45887
Elias I, Franckhauser S, Ferré T, Vilà L, Tafuro S, Muñoz S, et al. Adipose
tissue overexpression of vascular endothelial growth factor protects against
diet-induced obesity and insulin resistance. Diabetes (2012) 61:1801–13.
doi:10.2337/db11-0832
Sun K, Kusminski CM, Luby-Phelps K, Spurgin SB, An YA, Wang QA, et al.
Brown adipose tissue derived VEGF-A modulates cold tolerance and energy
expenditure. Mol Metab (2014) 3:474–83. doi:10.1016/j.molmet.2014.03.010
Nguyen KD, Qiu Y, Cui X, Goh YP, Mwangi J, David T, et al. Alternatively activated macrophages produce catecholamines to sustain adaptive thermogenesis.
Nature (2012) 480:104–8. doi:10.1038/nature10653
Bassaganya-Riera J, Misyak S, Guri AJ, Hontecillas R. PPAR gamma is highly
expressed in F4/80(hi) adipose tissue macrophages and dampens adipose-tissue
inflammation. Cell Immunol (2009) 258:138–46. doi:10.1016/j.cellimm.2009.
04.003

Frontiers in Immunology | Molecular Innate Immunity

Macrophage ontogeny and polarization

117. Stienstra R, Duval C, Keshtkar S, van der Laak J, Kersten S, Müller M. Peroxisome proliferator-activated receptor gamma activation promotes infiltration
of alternatively activated macrophages into adipose tissue. J Biol Chem (2008)
283:22620–7. doi:10.1074/jbc.M710314200
118. Odegaard JI, Ricardo-gonzalez RR, Goforth MH, Christine R, Subramanian
V, Mukundan L, et al. Macrophage-specific PPARgamma controls alternative activation and improves insulin resistance. Nature (2008) 447:1116–20.
doi:10.1038/nature05894
119. Bouhlel MA, Derudas B, Rigamonti E, Dièvart R, Brozek J, Haulon S,
et al. PPARgamma activation primes human monocytes into alternative M2
macrophages with anti-inflammatory properties. Cell Metab (2007) 6:137–43.
doi:10.1016/j.cmet.2007.06.010
120. Guri AJ, Hontecillas R, Bassaganya-Riera J. Abscisic acid synergizes with rosiglitazone to improve glucose tolerance, down-modulate macrophage accumulation in adipose tissue: possible action of the cAMP/PKA/PPAR γ axis. Clin Nutr
(2011) 29:646–53. doi:10.1016/j.clnu.2010.02.003
121. Satoh T, Kidoya H, Naito H, Yamamoto M, Takemura N, Nakagawa K, et al.
Critical role of Trib1 in differentiation of tissue-resident M2-like macrophages.
Nature (2013) 495:524–8. doi:10.1038/nature11930
122. Akira S, Misawa T, Satoh T, Saitoh T. Macrophages control innate inflammation. Diabetes Obes Metab (2013) 15(Suppl 3):10–8. doi:10.1111/dom.12151
123. Sica A, Invernizzi P, Mantovani A. Macrophage plasticity and polarization in
liver homeostasis and pathology. Hepatology (2014) 59:2034–42. doi:10.1002/
hep.26754
124. Balmer ML, Slack E, de Gottardi A, Lawson MAE, Hapfelmeier S, Miele
L, et al. The liver may act as a firewall mediating mutualism between the
host and its gut commensal microbiota. Sci Transl Med (2014) 6:237ra66.
doi:10.1126/scitranslmed.3008618
125. Dambach DM, Watson LM, Gray KR, Durham SK, Laskin DL. Role of CCR2 in
macrophage migration into the liver during acetaminophen-induced hepatotoxicity in the mouse. Hepatology (2002) 35:1093–103. doi:10.1053/jhep.2002.
33162
126. Zigmond E, Samia-Grinberg S, Pasmanik-Chor M, Brazowski E, Shibolet O,
Halpern Z, et al. Infiltrating monocyte-derived macrophages and resident
Kupffer cells display different ontogeny and functions in acute liver injury.
J Immunol (2014) 193:344–53. doi:10.4049/jimmunol.1400574
127. Holt AP, Haughton EL, Lalor PF, Filer A, Buckley CD, Adams DH. Liver myofibroblasts regulate infiltration and positioning of lymphocytes in human liver.
Gastroenterology (2009) 136:705–14. doi:10.1053/j.gastro.2008.10.020
128. You Q, Holt M, Yin H, Li G, Hu C-J, Ju C. Role of hepatic resident and infiltrating macrophages in liver repair after acute injury. Biochem Pharmacol (2013)
86:836–43. doi:10.1016/j.bcp.2013.07.006
129. Tarantino G, Saldalamacchia G, Conca P, Arena A. Non-alcoholic fatty liver
disease: further expression of the metabolic syndrome. J Gastroenterol Hepatol
l (2007) 22:293–303. doi:10.1111/j.1440-1746.2007.04824.x
130. Loomba R, Sanyal AJ. The global NAFLD epidemic. Nat Rev Gastroenterol
Hepatol (2013) 10:686–90. doi:10.1038/nrgastro.2013.171
131. Michelotti GA, Machado MV, Diehl AM. NAFLD, NASH and liver cancer. Nat
Rev Gastroenterol Hepatol (2013) 10:656–65. doi:10.1038/nrgastro.2013.183
132. Sun B, Karin M. Obesity, inflammation, and liver cancer. J Hepatol (2014)
56:704–13. doi:10.1016/j.jhep.2011.09.020
133. Endo M, Masaki T, Seike M, Yoshimatsu H. TNF-a induces hepatic steatosis
in mice by enhancing gene expression of sterol regulatory element binding
protein-1c. Exp Biol Med (2007) 232:614–21.
134. Pandey AK, Munjal N, Datta M. Gene expression profiling and network
analysis reveals lipid and steroid metabolism to be the most favored by TNFalpha in HepG2 cells. PLoS One (2010) 5:e9063. doi:10.1371/journal.pone.
0009063
135. Tosello-Trampont A-C, Landes SG, Nguyen V, Novobrantseva TI, Hahn YS.
Kupffer cells trigger nonalcoholic steatohepatitis development in diet-induced
mouse model through tumor necrosis factor-α production. J Biol Chem (2012)
287:40161–72. doi:10.1074/jbc.M112.417014
136. Musso G, Gambino R, Cassader M. Cholesterol metabolism and the pathogenesis of non-alcoholic steatohepatitis. Prog Lipid Res (2013) 52:175–91.
doi:10.1016/j.plipres.2012.11.002
137. Jeon T-I, Osborne TF. SREBPs: metabolic integrators in physiology and metabolism. Trends Endocrinol Metab (2012) 23:65–72. doi:10.1016/j.tem.2011.10.
004

January 2015 | Volume 5 | Article 683 | 228

Dey et al.

138. Shimomura I, Matsuda M, Hammer RE, Bashmakov Y, Brown MS, Goldstein
JL. Decreased IRS-2 and increased SREBP-1c lead to mixed insulin resistance
and sensitivity in livers of lipodystrophic and ob/ob mice. Mol Cell (2000)
6:77–86. doi:10.1016/S1097-2765(00)00009-5
139. Moon Y-A, Liang G, Xie X, Frank-Kamenetsky M, Fitzgerald K, Koteliansky V,
et al. The Scap/SREBP pathway is essential for developing diabetic fatty liver
and carbohydrate-induced hypertriglyceridemia in animals. Cell Metab (2012)
15:240–6. doi:10.1016/j.cmet.2011.12.017
140. Miele L, Valenza V, La Torre G, Montalto M, Cammarota G, Ricci R, et al.
Increased intestinal permeability and tight junction alterations in nonalcoholic
fatty liver disease. Hepatology (2009) 49:1877–87. doi:10.1002/hep.22848
141. Thuy S, Ladurner R,Volynets V,Wagner S, Strahl S, Königsrainer A, et al. Nonalcoholic fatty liver disease in humans is associated with increased plasma endotoxin and plasminogen activator inhibitor 1 concentrations and with fructose
intake. J Nutr (2008) 138:1452–5.
142. Suganami T, Tanimoto-Koyama K, Nishida J, Itoh M, Yuan X, Mizuarai S,
et al. Role of the toll-like receptor 4/NF-kappaB pathway in saturated fatty
acid-induced inflammatory changes in the interaction between adipocytes and
macrophages. Arterioscler Thromb Vasc Biol. (2007) 27:84–91. doi:10.1161/01.
ATV.0000251608.09329.9a
143. Yu M, Wang H, Ding A, Golenbock DT, Latz E, Czura CJ, et al. HMGB1 signals through toll-like receptor (TLR) 4 and TLR2. Shock (2006) 26:174–9.
doi:10.1097/01.shk.0000225404.51320.82
144. Aron-Wisnewsky J, Gaborit B, Dutour A, Clement K. Gut microbiota and
non-alcoholic fatty liver disease: new insights. Clin Microbiol Infect (2013)
19:338–48. doi:10.1111/1469-0691.12140
145. Soares J-B, Pimentel-Nunes P, Roncon-Albuquerque R, Leite-Moreira A. The
role of lipopolysaccharide/toll-like receptor 4 signaling in chronic liver diseases.
Hepatol Int (2010) 4:659–72. doi:10.1007/s12072-010-9219-x
146. Imajo K, Fujita K, Yoneda M, Nozaki Y, Ogawa Y, Shinohara Y, et al. Hyperresponsivity to low-dose endotoxin during progression to nonalcoholic steatohepatitis is regulated by leptin-mediated signaling. Cell Metab (2012) 16:44–54.
doi:10.1016/j.cmet.2012.05.012
147. Miura K, Kodama Y, Inokuchi S, Schnabl B, Aoyama T, Ohnishi H, et al. Toll-like
receptor 9 promotes steatohepatitis by induction of interleukin-1beta in mice.
Gastroenterology (2010) 139:323.e–34.e. doi:10.1053/j.gastro.2010.03.052
148. Pellicoro A, Ramachandran P, Iredale JP, Fallowfield JA. Liver fibrosis and repair:
immune regulation of wound healing in a solid organ. Nat Rev Immunol (2014)
14:181–94. doi:10.1038/nri3623

www.frontiersin.org

Macrophage ontogeny and polarization

149. Brancato SK, Albina JE. Wound macrophages as key regulators of repair: origin,
phenotype, and function. Am J Pathol (2011) 178:19–25. doi:10.1016/j.ajpath.
2010.08.003
150. Dooley S, ten Dijke P. TGF-β in progression of liver disease. Cell Tissue Res
(2012) 347:245–56. doi:10.1007/s00441-011-1246-y
151. Pesce JT, Ramalingam TR, Mentink-Kane MM, Wilson MS, El Kasmi KC,
Smith AM, et al. Arginase-1-expressing macrophages suppress Th2 cytokinedriven inflammation and fibrosis. PLoS Pathog (2009) 5:e1000371. doi:10.
1371/journal.ppat.1000371
152. Ramachandran P, Pellicoro A, Vernon MA, Boulter L, Aucott RL, Ali A, et al.
Differential Ly-6C expression identifies the recruited macrophage phenotype,
which orchestrates the regression of murine liver fibrosis. Proc Natl Acad Sci U
S A. (2012) 109:E3186–95. doi:10.1073/pnas.1119964109
153. Wan J, Benkdane M, Teixeira-Clerc F, Bonnafous S, Louvet A, Lafdil F, et al.
M2 Kupffer cells promote M1 Kupffer cell apoptosis: a protective mechanism against alcoholic and nonalcoholic fatty liver disease. Hepatology (2014)
59:130–42. doi:10.1002/hep.26607
154. Wan J, Benkdane M, Alons E, Lotersztajn S, Pavoine C. M2 Kupffer cells promote hepatocyte senescence: an IL-6-dependent protective mechanism against
alcoholic liver disease. Am J Pathol (2014) 184:1763–72. doi:10.1016/j.ajpath.
2014.02.014
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 12 September 2014; accepted: 17 December 2014; published online: 22 January
2015.
Citation: Dey A, Allen J and Hankey-Giblin PA (2015) Ontogeny and polarization
of macrophages in inflammation: blood monocytes versus tissue macrophages. Front.
Immunol. 5:683. doi: 10.3389/fimmu.2014.00683
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2015 Dey, Allen and Hankey-Giblin. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The
use, distribution or reproduction in other forums is permitted, provided the original
author(s) or licensor are credited and that the original publication in this journal is cited,
in accordance with accepted academic practice. No use, distribution or reproduction is
permitted which does not comply with these terms.

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REVIEW ARTICLE
published: 28 November 2014
doi: 10.3389/fimmu.2014.00614

Molecular mechanisms that influence the macrophage
M1–M2 polarization balance
Nan Wang 1,2 , Hongwei Liang 1,2 and Ke Zen 1,2 *
1
2

State Key Laboratory of Pharmaceutical Biotechnology, Nanjing University School of Life Sciences, Nanjing, China
Jiangsu Engineering Research Center for MicroRNA Biology and Biotechnology (JERC-MBB), Nanjing University School of Life Sciences, Nanjing, China

Edited by:
Laurel L. Lenz, University of Colorado
School of Medicine, USA
Reviewed by:
John P. Vasilakos, 3M Company, USA
Charles Dudley Mills, BioMedical
Consultants, USA
*Correspondence:
Ke Zen, State Key Laboratory of
Pharmaceutical Biotechnology,
Jiangsu Engineering Research Center
for MicroRNA Biology and
Biotechnology (JERC-MBB), Room
A229, School of Life Sciences, Xianlin
Campus of Nanjing University, No.
163, Xianlin Road, Qixia District,
Nanjing, Jiangsu 210046, China
e-mail: kzen@nju.edu.cn

As an essential component of innate immunity, macrophages have multiple functions in
both inhibiting or promoting cell proliferation and tissue repair. Diversity and plasticity are
hallmarks of macrophages. Classical M1 and alternative M2 activation of macrophages,
mirroring the Th1–Th2 polarization of T cells, represent two extremes of a dynamic changing state of macrophage activation. M1-type macrophages release cytokines that inhibit the
proliferation of surrounding cells and damage contiguous tissue, and M2-type macrophages
release cytokines that promote the proliferation of contiguous cells and tissue repair. M1–
M2 polarization of macrophage is a tightly controlled process entailing a set of signaling
pathways, transcriptional and posttranscriptional regulatory networks. An imbalance of
macrophage M1–M2 polarization is often associated with various diseases or inflammatory
conditions.Therefore, identification of the molecules associated with the dynamic changes
of macrophage polarization and understanding their interactions is crucial for elucidating
the molecular basis of disease progression and designing novel macrophage-mediated
therapeutic strategies.
Keywords: innate immune response, macrophage polarization, TLR, NLR, SOCS, microRNA

INTRODUCTION
As an essential component of innate immunity, macrophages are
capable of differentiating into protean varieties with a range of
function (1–3). In respond to various environmental cues (e.g.,
microbial products, damaged cells, activated lymphocytes) or
under different pathophysiologic conditions, macrophages can
acquire distinct functional phenotypes via undergoing different phenotypic polarization (4). Macrophage M1 and M2-type
responses describe the opposing activities of killing or repairing, and such polarized responses stimulate Th1- or Th2-like
responses in macrophages, respectively. First, M1 phenotype is
stimulated by microbial products or pro-inflammatory cytokines
[IFN-γ, TNF, or Toll-like receptor (TLR) ligands], and the typical characteristics of M1 macrophages include high antigen presentation, high production of IL-12 and IL-23, and high production of nitric oxide (NO) and reactive oxygen intermediates
(ROI) (5). In contrast, M2-type responses are the “resting” phenotype and are observed in healing-type circumstances without
infections. Such responses can also be further amplified by IL4, IL-10, or IL-13. M2 macrophages are characterized by the
upregulation of Dectin-1, DC-SIGN, mannose receptor, scavenger
receptor A, scavenger receptor B-1, CD163, CCR2, CXCR1, and
CXCR2 (6). Instead of generating NO or ROI, M2 macrophages
produce ornithine and polyamines through the arginase pathway (2, 7). In fact, from the functional point view, NO and
Ornithine, correlating to killing (M1) and repairing function (M2)
of macrophages, have been regarded by some investigators as the
most characteristic molecules of macrophages (8). Second, inflammatory M1 macrophages produce many other pro-inflammatory
cytokines like TNFα, IL-1, IL-6, IL-12, Type I IFN, CXCL1-3,

Frontiers in Immunology | Molecular Innate Immunity

CXCL-5, and CXCL8-10 (9), while M2 macrophages generate antiinflammatory cytokine such as IL-10 and very low level of proinflammatory cytokine such as IL-12 (10). Additional signatures
of M2 phenotype, such as YM1 (a member of the chitinase family) and FIZZ1 (found in inflammatory zone 1, RETNLA) are also
identified (11). Third, M1 macrophages promote Th1 response
and possess strong microbicidal and tumoricidal activity, while
M2 macrophages are involved in metazoan parasites containment
and promotion of Th2 response, tissue remodeling, immune tolerance, and tumor progression (12, 13). Additional information
about polarized activation of macrophages can be found in the
previous reviews (1, 14–16).
A coordinate action of various inflammatory modulators, signaling molecules, and transcription factors is involved in regulating macrophage polarization. At cellular level, although M1
and M2 macrophage activities exist without T or B cell influence
(17), specialized or polarized T cells (Th1, Th2, Tregs) do play a
role in macrophage polarized activation (1). Canonical IRF/STAT
signaling is a central pathway in modulating macrophage polarization. Activation of IRF/STAT signaling pathways by IFNs and
TLR signaling will skew macrophage function toward the M1 phenotype (via STAT1), while activation of IRF/STAT (via STAT6)
signaling pathways by IL-4 and IL-13 will skew macrophage function toward the M2 phenotype (9). Signals initiated by IL-10,
glucocorticoid hormones, apoptotic cell-released molecules, and
immune complexes can also profoundly affect macrophage functional statue (1). Macrophage polarization is also modulated
by local microenvironmental conditions such as hypoxia (18).
More importantly, M1–M2 polarization of macrophage is a highly
dynamic process and the phenotype of polarized macrophages

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can be reversed under physiological and pathological conditions
(19, 20). In the course of various pathophysiological settings, the
same signaling pathway can be involved in either M1 or M2 polarization of macrophages. The molecular mechanisms that govern
the phenotype switch of macrophages, however, remains incompletely understood. Moreover, imbalances of macrophage M1–M2
polarization are associated with various diseases. Disease conditions are frequently associated with polarization of macrophage
activation, with classically activated M1 macrophages implicated
in initiating and sustaining inflammation and M2 macrophages
associated with resolution of chronic inflammation (6). In the
past decade, a new class of small non-coding RNAs, termed as
microRNAs (miRNAs), have emerged as important regulators in
biological processes. An important role of miRNAs in modulating macrophage phenotypic polarization is demonstrated by
accumulating evidences in which an excessive or impaired inflammatory response of macrophages is found to be tightly linked to
the deregulation of miRNAs. In this review, we focus on recent
progress in understanding the molecular basis underlying the

FIGURE 1 | Mechanisms underlying the polarization of
macrophages. The major regulatory pathways of macrophage M1–M2
polarization are outlined. The crosstalk between the M1 and M2
macrophage polarizing pathways, particularly the balance between
activation of STAT1 and STAT3/STAT6, tightly regulates macrophage
polarization and activity. A predominance of NF-κB and STAT1 activation
promotes M1 macrophage polarization, resulting in cytotoxic and
tissue-damage proinflammatory functions. In contrast, a predominance
of STAT3 and STAT6 activation by IL-4/13 and IL-10 increases M2
macrophage polarization, associated with immune tolerance and tissue

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Macrophage M1–M2 polarization

dynamic macrophage polarization, including signaling pathways,
transcription factors and miRNAs.

IRF/STAT SIGNALING
As shown in Figure 1, IRF/STAT signaling is a central pathway
in controlling macrophage M1–M2 polarization. Toll-like receptor signaling, particularly TLR4 stimulated by lipopolysaccharide
(LPS) and other microbial ligands, drives macrophages to a preferentially M1 phenotype. Two adaptors, MyD88 and TRIF, mediate
the signaling downstream of TLR4. The signaling pathway through
the MyD88 adaptor results in the activation of a cascade of kinases,
including IRAK4, TRAF6, and IKKβ, which finally leads to the activation of nuclear factor kappa B (NF-κB). As a key transcription
factor related to macrophage M1 activation, NF-κB regulates the
expression of a large number of inflammatory genes including
TNFα, IL1B, cyclooxygenase 2 (COX2), IL-6, and IL12p40. NF-κB
activity is modulated via the activation of the inhibitor of kappa
B kinase (IKK) trimeric complex (two kinases, IKKα, IKKβ, and a
regulatory protein, IKKγ). When upstream signals converge at the

repairing. PPARδ (and PPARγ) control distinct aspects of M2 macrophage
activation and oxidative metabolism. KLF-4, a downstream of STAT6,
participates in the promotion of M2 macrophage functions by
suppressing the NF-κB/HIF-1α-dependent transcription. IL-4 induces not
only c-Myc, which controls the expression of a subset of M2-associated
genes but also the M2-polarizing IRF-4 axis to inhibit IRF5-mediated M1
polarization. IL-10 promotes M2 polarization through the induction of p50
NF-κB homodimer, c-Maf, and STAT3 activities. MicroRNAs such as
miR-155, miR-223, etc. are involved in modulating macrophage
polarization via targeting SOCS1, CEBP, and Pknox1, respectively.

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IKK complex, they first activate IKKβ via phosphorylation, and
activated IKKβ further phosphorylates the inhibitory molecule,
inhibitor of kappa B (I-κB). This results in the proteosomal degradation of I-κB and the release of NF-κB p65/p50 heterodimer from
the NF-κB/I-κB complex. The NF-κB p65/p50 heterodimer is then
translocated to the nucleus and binds to the promoters of inflammatory genes. The signaling through the TRIF adaptor pathway
activates the transcription factor interferon-responsive factor 3
(IRF3), leading to the expression and secretion of type I interferon, such as IFNα and IFNβ. Secreted type 1 interferons bind to
the type I interferon receptor (IFNAR) with consequent activation
of the transcription factor STAT1. It has been widely reported that
IRF3 and IRF5 are involved in regulating M1 polarization and M1associated gene induction (21, 22). IFN-stimulated genes include
chemokine CXCL9 and CXCL10 (23), which are characteristic of
classical M1 macrophage activation. In fact, macrophage polarization is tightly linked to the differential expression of various
TLRs on macrophages. The ratio of TLR4/TLR2 is significantly
higher in M1 macrophages compared to M2 macrophages (24),
while TLR4 deficiency promotes the alternative activation (M2)
of adipose tissue macrophages (ATMs) (25). TLR ligands, e.g.,
imiquimod and CpG, have been used as therapeutic treatments for
inflammatory diseases such as asthma by modulating macrophage
polarization.
Toll-like receptor and Toll-like receptor-induced cytokinereceptor cascades are broadly inhibited by tyrosine kinases Tyro3,
Axl, and Mer. IFNβ can activate the receptor for Axl, Tyro3, and
Mer and negatively regulate TLR signaling through induction of
SOCS1 and SOCS3 (26). A hyperactive signaling mediated by
Tyro3, Axl, and Mer receptor is suggested to induce immunosuppression in severe sepsis patients (26). Along the same lines,
chronic signaling through the TLR4 pathway has been shown
to induce various negative regulators like IRAK-M, ST2, SOCS1,
short version of MyD88 (MyD88sh) (27, 28) and SHIPs (29).
These negative regulators inhibit TLR-mediated signaling and
thus switch macrophages to an immunosuppressive, endotoxintolerant phenotype. A switching from an MyD88-dependent to
a TRIF-dependent TLR4 pathway in macrophages has also been
suggested to shift macrophage phenotype from an inflammatory
to anti-inflammatory, endotoxin-tolerant phenotype (30). Thus,
interplay of signaling molecules and transcription factors can
reverse the phenotype of macrophage polarization.
STAT-mediated activation of macrophages is regulated by
members of the suppressor of cytokine signaling (SOCS) family.
SOCS family members are inducible inhibitors of cytokine signals
and thus play a critical role in limiting inflammation responses.
SOCS proteins could be induced by cytokine signaling pathway,
and they in turn inhibit the cytokine signaling by several mechanisms. For example, IL-4 and IFN-γ, the latter in concert with
TLR stimulation, upregulate SOCS1 (31) and SOCS3 (32), which
in turn, inhibit the action of STAT1 and STAT3, respectively.
SOCS proteins can be also directly induced by TLR signaling.
In macrophages, SOCS proteins not only regulate the sensitivity of cells toward cytokines but also modulate signaling through
TLRs. Because SOCS3 is a downstream molecule of Notch signaling (33), it is likely that Notch signaling determines the M1 versus
M2 polarization of macrophages through SOCS3 (34). However,

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Macrophage M1–M2 polarization

the role of SOCS3 in modulating macrophage M1–M2 polarization is controversial. Although the unique expression of SOCS3
was reported to be essential for classic macrophage activation (32),
SOCS3 deficiency also promotes M1 macrophage polarization and
inflammation (35).
Macrophages can be driven to M2 phenotype by canonical M2 stimuli like IL-4, IL-13, and IL-10 (36, 37). As shown
in Figure 1, IL-4 and IL-13 polarize macrophages to M2 phenotype via activating STAT6 through the IL-4 receptor alpha
(IL-4Rα), whereas IL-10 promotes M2 phenotype via activating
STAT3 through receptor (IL-10R). In IL-4 and IL-13 pathway,
receptor binding of IL-4 activates JAK1 and JAK3 (38), leading
to STAT6 activation and translocation. Macrophage M2 phenotype is promoted by several transcription factors, including
peroxisome proliferator activated receptor γ (PPARγ) (39, 40)
and Krueppel-like factor 4 (KLF-4) (41). Myeloid-specific deficiency of either PPARγ or KLF-4 resulted in suppressed M2
polarization of macrophages, leading to accelerated lesion formation in apolipoprotein E-deficient (42) or low-density lipoprotein
receptor-knockout (43) mice. Moreover, ligation of PPARγ by specific PPARγ ligands resulted in a preferential M2 polarization
in mice and in human beings (40). Other transcription factors
involved in this process include c-Myc and IRF4. Transciptome
analysis of IL-4-stimulated cells consists of various enzymes and
transcription factors, including transglutaminase 2 (TGM2), mannose receptor, cholesterol hydroxylase CH25H, prostaglandinendoperoxide synthase PTGS1 (prostaglandin G/H synthase 1),
transcription factors IRF4, KLF-4, and the signaling modulators
CISH and SOCS1 (44). During severe respiratory syncytial virus
(RSV)-induced bronchiolitis, IL-4Rα/STAT6-dependent M2 differentiation of macrophages reduces inflammation and epithelial
damage in lungs (45).
PPARγ play an important role in modulating macrophage
M2 polarization induced by IL-4 or IL-13 (46). Studies using
PPARγ-deficient macrophages have shown the role of this nuclear
receptor in promoting M2 activation to protect mice from insulin
resistance (47). A similar role was also found for the PPARδ in
determination of macrophage polarization (48). Using myeloidspecific transcription factor KLF-4 knockout mice, Liao et al.
(41) demonstrated the role of KLF-4 in regulating M2 polarization of macrophages as well as in protecting mice from obesityinduced insulin resistance. In a similar fashion, IRF4 is also
involved in regulating the expression of genes associated with
macrophage M2 polarization (49). Collectively, all these findings
suggest that STAT6, PPARγ, KLF-4, and IRF4 may coordinate the
M2 polarization of macrophages.
IL-10R, a heterodimer of IL-10R1 and IL-10R2, is a receptor for
IL-10. Ligation of IL-10R with IL-10 results in the autophosphorylation of IL-10R, leading to the activation of the transcription
factor STAT3 and reduction of pro-inflammatory cytokine expression. In macrophages, IL-10 is also reported to respond to TLR
activation, glucocorticoid treatment, and C-type lectin signaling
(e.g., DC-SIGN and dectin 1 ligation). The components in IL10-induced macrophage transcriptome include specific Fc receptors, chemoattractants CXCL13 and CXCL4, recognition receptors
formyl peptide receptor 1 (FPR1), TLR1, TLR8, and macrophage
receptor with collagenous domain (50).

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HIF-1α AND HIF-2α
Macrophages can rapidly alter their metabolic and functional
state to adapt to the microenvironment of surrounding tissues.
Microenvironmental conditions in infected, inflamed, or damaged tissues are generally lack of oxygen and nutrients. When
macrophages are recruited into inflammatory sites, they encounter
the hypoxia condition, which can directly affect the macrophage
polarization. Hypoxia executes its effect on macrophages through
two isoforms of hypoxia-inducible factor (HIF), HIF-1α and HIF2
(51, 52). Gene expression profiling of macrophages and monocytes has identified profound changes in response to hypoxia (53,
54). Hypoxia strongly induces the expression of angiogenesisand metastasis-related genes such as VEGF, FGF2, MMP7, and
MMP9. Upregulation of those genes under hypoxia would lead
to more recruitment of macrophages into the hypoxic (avascular) areas in pathologies like atherosclerosis, obesity, and cancer
where they dampen the inflammation or promote tumor progression. In addition, pro-inflammatory cytokines like TNFα, IL-1β,
MIF, CCL3, and COX2, as well as M2 markers like IL-10 and
arginase 1 in macrophages, are also induced by hypoxia (55).
The crucial role of hypoxia in regulating macrophage inflammatory response has been confirmed in mice with myeloid cellspecific deletion of HIF-1α (56), in which HIF-1α was found to
be essential in regulating myeloid cell glycolytic capacity and survival and function in the inflammatory microenvironment. This
is in line with the finding that HIF-1α was induced by NF-κB
(57) and plays an important role in modulating macrophage
phagocytosis of bacteria under sepsis conditions (58). Moreover, recent studies also showed that HIF-1α can mediate the
effects of tumor-derived lactic acid (59) and cytokines (Oncostatin M and Eotaxin) (60) on promoting M2-like phenotype. In
contrast to these studies, recent study of myeloid-specific HIF2α deletion showed the role of HIF2 in mediating macrophage
inflammatory responses rather than HIF-1α (52). In contrast to
these studies, a recent (61) suggested that HIF-1α and HIF-2α
might also drive macrophage polarization by modulating NO
homeostasis in a cytokine-induced and transcription-dependent
fashion. Specifically, this study showed that inducible NO synthase gene and the arginase 1 gene in polarized macrophages are
specifically regulated by HIFs (61). Although HIF-1α and HIF-2α
displayed physiologically antagonistic functions, their antiphase
regulation allows them to coordinately regulate NO production
to guide macrophage polarization. Together, these findings suggest HIFs as an important regulator of macrophage polarization,
although a detailed dissection of whether the alteration of HIF isoform expression can switch macrophage phenotypes needs further
investigation.

OLIGOMERIZATION DOMAIN (NOD)-LIKE RECEPTORS
Stimulated by a diverse set of stimulus, including interferon-γ
(IFN-γ), LPS, and other TLR activators, macrophages are polarized toward to M1 state in which oxidative metabolites and proinflammatory cytokines are produced. Engagement of the respective receptors by these stimulus results in activation of the adapter
proteins such as MyD88, leading to sequential activation of kinases,
phosphorylation of transcription factors, and eventual genetic
program induction. Pro-inflammatory genes, including IFN-γ,

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Macrophage M1–M2 polarization

tumor necrosis factor-α (TNF-α), IL-1β, IL-18, chemokines, and
proteases, are subsequently produced. Further activation of the
M1 pathway occurs through the assembly of the NLR inflammasome and caspase-1 activation, which results in the conversion of
IL-1β and IL-18 into secreted active forms (62). With the NLRP3
inflammasome serving as a sensor of obesity-associated danger
signals, the progression of obesity can switch macrophages from
“M2-like” to“M1-like” cells (63). In macrophages, the activation of
NLR stimulates the cryptopyrin/NLRP3 inflammasome to induce
IL-1β and IL-18 production via caspase-1. Caspase-1 and IL-1β
are induced in adipose tissue with diet-induced obesity (DIO), and
Nlrp3- and caspase-1-deficient mice both demonstrate a resistance
to DIO-induced inflammation (63). The mechanism of this protective effect may be driven by the alteration of M1 activation of
ATMs, as Nlrp3-knockout mice show decreased M1 but increased
M2 gene expression in ATMs.
In addition to binding to TLRs, some pathogen-associated
molecular patterns (PAMPs) are also recognized by a family of
cytosolic nucleotide-binding receptors and NOD-like receptors
(NLRs) (64), another groups of PAMP receptors. Some NLRs
are involved in the recognition of microbial molecules and/or
endogenous factors released from tissue destruction. This recognition can lead to activation of caspase-1 (a pro-inflammatory
caspase), and subsequent proteolytic conversion of potent proinflammatory cytokines IL-1β and IL-18 from their precursors
pro-IL-1β and pro-IL-18, respectively. The proteolytic conversion
of IL-1β and IL-18 is mediated by a cytosolic caspase 1-activating
protein complex, termed as inflammasome (65).
As the most well-characterized members of the NLR family, NOD1 is ubiquitously expressed and NOD2 is restricted to
monocytes, macrophages, dendritic cells, and intestinal Paneth
cells (66). Both NOD1 and NOD2 induce NF-κB activation in a
TLR-independent manner (67). Structural analysis demonstrated
that NOD1 and NOD2 recognize different core motifs derived
from peptidoglycan (PGN), a component of bacterial cell walls.
NOD1 activity is triggered by γ-d-glutamyl-meso-diaminopimelic
acid, a unique PGN structures from all Gram-negative and some
Gram-positive bacteria (68). In contrast, NOD2 is activated by
muramyl dipeptide, a PGN motif in all Gram-positive and Gramnegative bacteria (69). Upon ligand recognition, NOD1 and NOD2
undergo conformational changes and self-oligomerization, which
is followed by the recruitment and activation of the serine threonine kinase RICK (RIP2, also known as RIPK2), an essential step
for the activation of NF-κB and MAPKs. The ubiquitination of
RICK is essential for NOD1/NOD2-mediated signaling because
removal of this modification by deubiquitinating enzyme A20
largely dampens NOD1/NOD2-induced NF-κB activation (70,
71). Although both NOD1 and NOD2 induce similar K63-linked
ubiquitination of RICK for NF-κB activation and upregulation
of various inflammatory mediators, NOD2 signaling appears to
preferentially utilize the E3 ligase TRAF6 and NOD1-mediated
signaling is mainly associated with TRAF2 and TRAF5. Nevertheless, the role of NOD1 and NOD2 in activating NF-κB-dependent
inflammatory responses is not limited to the recognition of PGN
motifs. Recent study by Keestra et al. (72) reported that that NOD1
can sense activation state of small Rho GTPases. In this study,
NOD1 signaling pathway was triggered by RAC1 and CDC42

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activated by bacterial delivery or ectopic expression of SopE, a
virulence factor of the enteric pathogen Salmonella.

GRANULOCYTE-MACROPHAGE COLONY-STIMULATING
FACTOR
As the most recently discovered cytokine involved in regulation
of macrophage polarization, granulocyte-macrophage colonystimulating factor (GM-CSF) is produced by a variety of cells
including macrophages and parenchyma cells. The main functions
of GM-CSF include regulating the proliferation and differentiation of functional hematopoietic cells. The GM-CSF receptor
forms a dodecamer structure (73) and recruits JAK2, leading to the
activation of STAT5, extracellular signal-regulated kinase (ERK),
V-Akt murine thymoma viral oncogene homolog 1 (AKT), and the
nuclear translocation of NF-κB and IRF5 (21). Many of these regulators are part of the IFN-γ and TLR signaling pathways. GM-CSF
enhances macrophage antigen presentation, complement- and
antibody-mediated phagocytosis, microbicidal capacity, leukocyte
chemotaxis, and adhesion. GM-CSF induces cytokine production
of IL-6, IL-8, G-CSF, M-CSF, TNF, and IL-1β in monocytes and
macrophages, although the degree of cytokine induction by GMCSF is less than that by LPS. Global gene expression analyses
of macrophages differentiated from GM-CSF-treated monocytes
showed that GM-CSF upregulated 340 genes and downregulated
190 genes in macrophages. Macrophage-specific genes including
CD14, CD163, C5R1, and FcγR1A, and several cell surface adhesion molecules, cytokine receptors were induced by GM-CSF (74).
In this study, a high-resolution transcriptome profiling of human
macrophages by RNA sequencing was employed to discover novel
marker genes unique for human macrophages. A similar strategy
has been used to obtain a high-resolution transcriptome profile of
human macrophages under M1 (or M1-like) and M2 (or M2-like)
polarization conditions, resulting in a more comprehensive understanding of the transcriptome of human macrophages (75). The
GM-CSF deficient mice have normal numbers of macrophages
in many tissues but display an impaired maturation of alveolar
macrophages and develop pulmonary alveolar proteinosis (76). In
human beings, mutations in the GM-CSF receptor, especially in
the common beta chain, lead to alveolar macrophage dysfunction,
proteinosis, and malignancy (77, 78).

MicroRNAs
MicroRNAs (miRNAs), a class of 19–24 nt non-coding RNAs
that induce gene silencing at the posttranscriptional level,
have emerged as an important regulatory mechanism for gene
expression in many immune cells including monocytes and
macrophages (79, 80). Functional miRNAs associated with polarized macrophages have been identified (81). While these functional
miRNAs like miR-155 and miR-146 are induced by a variety of
inflammatory stimuli, including LPS, TNFα, and IL-1β, they are
instrumental in attenuating TLR4/IL-1R signaling pathways in
monocytes and macrophages (79, 82, 83). These findings allow
us to postulate that miRNAs may contribute to the switching of
inflammatory macrophages to an immunosuppressive phenotype,
needed for resolution. For instance, miR-146, miR-125b, miR-155,
and miR-9 have been shown to be induced by LPS, and in turn,
these miRNAs inhibit TLR4/IL-1R signaling through regulation

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of IRAK-1, TRAF6, IKKe, p50NF-jB, and TNFα at transcriptional
and posttranscriptional level (79, 82–86). Our recent study has also
shown that a panel of miRNAs including miR-17, miR-20a, and
miR-106a are stimulated by LPS through c-Myc pathway, and these
miRNAs collectively reduces the expression level of macrophage
differentiation related marker, signal-regulatory protein α (SIRPα)
(87). It has been reported that miR-98 and miR-21 inhibit the
expression of inflammatory genes in monocytes and macrophages
via controlling IL-10 level (88, 89). These findings strongly argue
that miRNAs can regulate macrophage phenotype in the course of
various diseases, for example, during endotoxin tolerance (30, 90).
In the efforts to delineate the role of miRNAs in macrophage
activation in inflammatory diseases, Ponomarev et al. (91) found
that miR-124 promotes microglia quiescence and suppresses EAE
by deactivating macrophages in a C/EBPα-PU.1-dependent manner. This is one of the few studies in which a specific miRNA
is found to regulate macrophage plasticity, although it remains
unclear how C/EBPα suppresses macrophage M2 polarization.
Zhang and co-workers (92) also reported that miR-223 modulates
obesity-associated adipose tissue inflammation through regulating macrophage activation. In the study, they found that miR-223
was upregulated in LPS-treated macrophages but downregulated
in IL-4-treated bone marrow-derived macrophages (BMDMs). In
agreement with the observation of differential expression of miR223 in various macrophages, the miR-223-deficient macrophages
were hypersensitive to LPS stimulation, whereas such macrophages
exhibited a delayed responses to IL-4 compared with controls.
Moreover, miR-223-deficient mice exhibited an increased adipose tissue inflammatory response but a decreased adipose tissue
insulin signaling. They further identified Pknox1 as a genuine
target of miR-223. Although Pknox1 as a miR-223 target in regulating macrophage polarization was validated by gain-of-function
and loss-of-function analyses in BMDMs, it remains unclear how
Pknox1 further regulates macrophage polarization.
A recent work by Banerjee et al. (93) demonstrated that
let-7c could regulate bactericidal and phagocytic activities
of macrophages, two functional phenotypes implicated in
macrophage polarization. In the study, they found that let-7c was
expressed at a higher level in M2-type macrophages than in M1type macrophages. When M2-type macrophages were re-polarized
to M1-type macrophages or M1-type macrophages converted to
M2-type macrophages, let-7c expression level was decreased or
increased, respectively. As LPS stimulation reduced let-7c expression in M2 macrophages, let-7c might play an inhibitory role
in modulating macrophage inflammatory responses. In line with
this, upregulation of let-7c in macrophages diminished M1 phenotype but promoted M2 phenotype polarization. Their study
further identified that let-7c targeted C/EBP-δ, a key transcriptional factor in macrophage pro-inflammatory response to TLR4
stimulation (94, 95).
The modulation of macrophage polarization by miR-155 has
also been recently reported (96, 97). The expression of miR-155
was found to be repressed in naive macrophages or LPS-stimulated
Akt2-/- macrophages. In this process, miR-155 targets transcriptional factor C/EBP-β, a hallmark of M2 macrophages. C/EBP-β
can regulate Arg1 and its level is increased upon Akt2 ablation.
Overexpression or depletion of miR-155 drove macrophages to

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M1 or M2 phenotype, respectively, confirming that miR-155 plays
a central role in regulating Akt-dependent M1/M2 polarization
of macrophages. It has also reported that miR-155 can directly
block IL-13-induced macrophage M2 phenotype via suppressing
the expression of IL-13Rα1 (96). As an oncomiR, miR-155 also targets SHIP1 to promote TNFα-dependent tumor cell growth (98).
Through overexpression of miR-155, we successfully re-polarized
tumor-associated macrophages (TAMs) into pro-inflammatory
M1 macrophages (97). Taken together, these studies support the
hypothesis that miR-155 is a key molecule in causing macrophage
polarization toward M1-type activity.

M1–M2 PHENOTYPE SWITCH
Macrophage differentiation is highly dynamic. Responding to
microenviromental cues macrophages can rapidly switch from
one phenotype to the other. In fact, activation of NF-κB or IRF
family members in macrophages by TLR4 or other TLRs can
drive macrophage to either M1 or M2 polarization under various
pathological conditions (99–105). Accumulating evidences have
shown that the spatiotemporal activation of NF-κB is a key regulator of the plasticity of macrophages observed in the courses
of various disease progressions. For example, during the early
phase of tumorigenesis, NF-κB activation in M1 macrophages
is critical for cancer-related inflammation. However, at the late
phase of tumorigenesis, macrophages are re-programed to TAM
or M2-like macrophages displaying low NF-κB activation but
increased immunosuppressive capacity (106). A similar situation of macrophage polarization is observed at different stages
along the progression of sepsis, in which NF-κB activation in
M1 macrophages drives the initial overt inflammatory phase,
while during the late phase of endotoxin tolerance, macrophages
are polarized to an anti-inflammatory, tumor growth-promoting
(M2) phenotype, and display an impaired NF-κB activation
(107). The studies on RSV infection also show that polarization of macrophages is complicated process and the phenotype
of macrophage activation can be varied at the different stage
along disease progression. As the most significant cause of lower
respiratory tract infection in infants and young children, RSV
infection is found to be associated with a mixed “Th1” and “Th2”
cytokine storm. At the initial stage of RSV infection, RAV induces
the expression of various anti-viral genes like IFN-β in airway
epithelial cells, and then promotes the expression of many NF-κBdependent pro-inflammatory genes in macrophages through stimulating TLR4, TLR2, TLR3, and retinoic acid-induce gene I (RIGI), driving macrophages toward anti-viral, pro-inflammatory M1
phenotype. However, to maintain a mild but persisting infection,
RSV also induces alveolar macrophages to produce IL-4 and IL13 that contribute to macrophage M2 polarization and disease
resolution through IL-4Rα/STAT6-, TLR4-, and IFN-β-dependent
signaling pathways (45).
Although under certain conditions like parasite infections
and allergy, the functional phenotypes of macrophages in vivo
largely mirror those of canonical M1 and M2-polarized states,
macrophage populations often express mixed phenotypes in the
course of various disease settings. Indeed, macrophages with
combinations of M1 and M2 markers can be found in neurodegenerative disorders (108), atherosclerotic plaques (109), and some

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Macrophage M1–M2 polarization

murine tumors (110). Therefore, the contribution of coexisting
macrophages with different phenotypes, the impact of dynamic
changes of macrophage plasticity on diseases, and the molecular
networks orchestrating the switch of macrophage phenotype are
required to be analyzed for a full understanding of the M1–M2
paradigm of macrophage polarization.

FUTURE DIRECTIONS
Tremendous progress has been made in defining the molecular networks underlying M1–M2-polarized activation of
macrophages. Molecular determinants of M1–M2 polarization
include members of the PPAR, KLF, IRF, STAT, NF-κB, and HIF
families, and miRNAs. However, new molecules that regulate
macrophage M1–M2 polarization may still remain unidentified. A
novel class of large intergenic non-coding RNAs, termed as lincRNAs, has been recently shown to be involved in both activation and
repression of immune response genes (111). Among thousands of
lincRNAs identified in the mammalian genome, 159 lincRNAs was
found to be differentially expressed following innate activation of
THP1 macrophages (112). In these differentially expressed lincRNAs, linc1992 was found to specifically bind to heterogenous
nuclear ribonucleoprotein L (hnRNPL) and form a linc1992hnRNPL complex that regulates TNFα gene transcription. The
role of lincRNAs in modulation of macrophage polarization, however, has not been reported so far but certainly needs to be further
studied.
Different from the irreversible phenotypic changes seen in
lymphocytes after exposure to polarizing cytokines, macrophage
polarization is transient and plastic. In order to adapt to
the microenvironmental conditions of surrounding tissues,
macrophages can rapidly switch their phenotypes. For example,
M2 macrophages can be re-polarized into macrophages with M1
phenotype following exposure to TLR ligands or IFNγ or overexpression of miR-155 (113, 114), whereas M1 macrophages can be
reprogramed to express various genes of M2 macrophage by treating macrophages with reagents that increase IL-10 level (115, 116).
Therefore, further exploring the dynamic process of macrophage
polarization and the mechanisms that govern this process not only
is important for our understanding of the M1–M2 paradigm of
macrophage polarization but also provides new therapeutic strategies for various diseases including cancers via targeting imbalances
of macrophage polarization.

ACKNOWLEDGMENTS
This work was supported by grants from the National Basic
Research Program of China (973 Program, 2012CB517603 and
2011CB504803), the National Natural Science Foundation of
China (No. 30988003, 30225037, 30471991, 30570731), and the
Natural Science Foundation of Jiangsu Province (No. BK2011013).

REFERENCES
1. Biswas SK, Mantovani A. Macrophage plasticity and interaction with lymphocyte subsets: cancer as a paradigm. Nat Immunol (2010) 11(10):889–96.
doi:10.1038/ni.1937
2. Mantovani A, Sozzani S, Locati M, Allavena P, Sica A. Macrophage polarization: tumor-associated macrophages as a paradigm for polarized M2 mononuclear phagocytes. Trends Immunol (2002) 23(11):549–55. doi:10.1016/S14714906(02)02302-5

November 2014 | Volume 5 | Article 614 | 235

Wang et al.

3. Mosser DM, Edwards JP. Exploring the full spectrum of macrophage activation.
Nat Rev Immunol (2008) 8(12):958–69. doi:10.1038/nri2448
4. O’Shea JJ, Paul WE. Mechanisms underlying lineage commitment and plasticity of helper CD4+ T cells. Science (2010) 327(5969):1098–102. doi:10.1126/
science.1178334
5. Verreck FA, de Boer T, Langenberg DM, Hoeve MA, Kramer M,Vaisberg E, et al.
Human IL-23-producing type 1 macrophages promote but IL-10-producing
type 2 macrophages subvert immunity to (myco)bacteria. Proc Natl Acad Sci U
S A (2004) 101(13):4560–5. doi:10.1073/pnas.0400983101
6. Martinez FO, Helming L, Gordon S. Alternative activation of macrophages: an
immunologic functional perspective. Annu Rev Immunol (2009) 27:451–83.
doi:10.1146/annurev.immunol.021908.132532
7. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism
and functions. Immunity (2010) 32(5):593–604. doi:10.1016/j.immuni.2010.
05.007
8. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
9. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122(3):787–95. doi:10.1172/JCI59643
10. Mosser DM. The many faces of macrophage activation. J Leukoc Biol (2003)
73(2):209–12. doi:10.1189/jlb.0602325
11. Raes G, Van den Bergh R, De Baetselier P, Ghassabeh GH, Scotton C, Locati
M, et al. Arginase-1 and Ym1 are markers for murine, but not human, alternatively activated myeloid cells. J Immunol (2005) 174(11):6561. doi:10.4049/
jimmunol.174.11.6561
12. Gordon S, Taylor PR. Monocyte and macrophage heterogeneity. Nat Rev
Immunol (2005) 5(12):953–64. doi:10.1038/nri1733
13. Mantovani A, Germano G, Marchesi F, Locatelli M, Biswas SK. Cancerpromoting tumor-associated macrophages: new vistas and open questions. Eur
J Immunol (2011) 41(9):2522–5. doi:10.1002/eji.201141894
14. Biswas SK, Mantovani A. Orchestration of metabolism by macrophages. Cell
Metab (2012) 15(4):432–7. doi:10.1016/j.cmet.2011.11.013
15. Biswas SK, Chittezhath M, Shalova IN, Lim JY. Macrophage polarization
and plasticity in health and disease. Immunol Res (2012) 53(1–3):11–24.
doi:10.1007/s12026-012-8291-9
16. Labonte AC, Tosello-Trampont AC, Hahn YS. The role of macrophage polarization in infectious and inflammatory diseases. Mol Cells (2014) 37(4):275–85.
doi:10.14348/molcells.2014.2374
17. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
18. Escribese MM, Casas M, Corbi AL. Influence of low oxygen tensions on
macrophage polarization. Immunobiology (2012) 217(12):1233–40. doi:10.
1016/j.imbio.2012.07.002
19. Saccani A, Schioppa T, Porta C, Biswas SK, Nebuloni M, Vago L, et al.
p50 nuclear factor-kappaB overexpression in tumor-associated macrophages
inhibits M1 inflammatory responses and antitumor resistance. Cancer Res
(2006) 66(23):11432–40. doi:10.1158/0008-5472.CAN-06-1867
20. Guiducci C, Vicari AP, Sangaletti S, Trinchieri G, Colombo MP. Redirecting
in vivo elicited tumor infiltrating macrophages and dendritic cells towards
tumor rejection. Cancer Res (2005) 65(8):3437–46.
21. Krausgruber T, Blazek K, Smallie T,Alzabin S, Lockstone H, Sahgal N, et al. IRF5
promotes inflammatory macrophage polarization and TH1-TH17 responses.
Nat Immunol (2011) 12(3):231–8. doi:10.1038/ni.1990
22. Fleetwood AJ, Dinh H, Cook AD, Hertzog PJ, Hamilton JA. GM-CSF- and MCSF-dependent macrophage phenotypes display differential dependence on
type I interferon signaling. J Leukoc Biol (2009) 86(2):411–21. doi:10.1189/jlb.
1108702
23. Donlin LT, Jayatilleke A, Giannopoulou EG, Kalliolias GD, Ivashkiv LB. Modulation of TNF-Induced Macrophage Polarization by Synovial Fibroblasts.
J Immunol (2014) 193(5):2373–83. doi:10.4049/jimmunol.1400486
24. Sauer RS, Hackel D, Morschel L, Sahlbach H, Wang Y, Mousa SA, et al.
Toll like receptor (TLR)-4 as a regulator of peripheral endogenous opioidmediated analgesia in inflammation. Mol Pain (2014) 10(1):10. doi:10.1186/
1744-8069-10-10
25. Orr JS, Puglisi MJ, Ellacott KL, Lumeng CN, Wasserman DH, Hasty
AH. Toll-like receptor 4 deficiency promotes the alternative activation of
adipose tissue macrophages. Diabetes (2012) 61(11):2718–27. doi:10.2337/
db11-1595

Frontiers in Immunology | Molecular Innate Immunity

Macrophage M1–M2 polarization

26. Rothlin CV, Ghosh S, Zuniga EI, Oldstone MB, Lemke G. TAM receptors are pleiotropic inhibitors of the innate immune response. Cell (2007)
131(6):1124–36. doi:10.1016/j.cell.2007.10.034
27. Kobayashi K, Hernandez LD, Galan JE, Janeway CA Jr., Medzhitov R, Flavell
RA. IRAK-M is a negative regulator of Toll-like receptor signaling. Cell (2002)
110(2):191–202. doi:10.1016/S0092-8674(02)00827-9
28. Liew FY, Xu D, Brint EK, O’Neill LA. Negative regulation of toll-like receptormediated immune responses. Nat Rev Immunol (2005) 5(6):446–58. doi:10.
1038/nri1630
29. Rauh MJ, Ho V, Pereira C, Sham A, Sly LM, Lam V, et al. SHIP represses the generation of alternatively activated macrophages. Immunity (2005) 23(4):361–74.
doi:10.1016/j.immuni.2005.09.003
30. Biswas SK, Lopez-Collazo E. Endotoxin tolerance: new mechanisms, molecules
and clinical significance. Trends Immunol (2009) 30(10):475–87. doi:10.1016/
j.it.2009.07.009
31. Whyte CS, Bishop ET, Ruckerl D, Gaspar-Pereira S, Barker RN, Allen JE, et al.
Suppressor of cytokine signaling (SOCS)1 is a key determinant of differential macrophage activation and function. J Leukoc Biol (2011) 90(5):845–54.
doi:10.1189/jlb.1110644
32. Liu Y, Stewart KN, Bishop E, Marek CJ, Kluth DC, Rees AJ, et al. Unique expression of suppressor of cytokine signaling 3 is essential for classical macrophage
activation in rodents in vitro and in vivo. J Immunol (2008) 180(9):6270–8.
doi:10.4049/jimmunol.180.9.6270
33. Narayana Y, Balaji KN. NOTCH1 up-regulation and signaling involved in
Mycobacterium bovis BCG-induced SOCS3 expression in macrophages. J Biol
Chem (2008) 283(18):12501–11. doi:10.1074/jbc.M709960200
34. Wang YC, He F, Feng F, Liu XW, Dong GY, Qin HY, et al. Notch signaling determines the M1 versus M2 polarization of macrophages in antitumor
immune responses. Cancer Res (2010) 70(12):4840–9. doi:10.1158/0008-5472.
CAN-10-0269
35. Qin H, Holdbrooks AT, Liu Y, Reynolds SL, Yanagisawa LL, Benveniste EN.
SOCS3 deficiency promotes M1 macrophage polarization and inflammation.
J Immunol (2012) 189(7):3439–48. doi:10.4049/jimmunol.1201168
36. O’Farrell AM, Liu Y, Moore KW, Mui AL. IL-10 inhibits macrophage activation and proliferation by distinct signaling mechanisms: evidence for
Stat3-dependent and -independent pathways. EMBO J (1998) 17(4):1006–18.
doi:10.1093/emboj/17.4.1006
37. Lang R, Patel D, Morris JJ, Rutschman RL, Murray PJ. Shaping gene expression in activated and resting primary macrophages by IL-10. J Immunol (2002)
169(5):2253–63. doi:10.4049/jimmunol.169.5.2253
38. Nelms K, Keegan AD, Zamorano J, Ryan JJ, Paul WE. The IL-4 receptor: signaling mechanisms and biologic functions. Annu Rev Immunol (1999) 17:701–38.
doi:10.1146/annurev.immunol.17.1.701
39. Chawla A. Control of macrophage activation and function by PPARs. Circ Res
(2010) 106(10):1559–69. doi:10.1161/CIRCRESAHA.110.216523
40. Bouhlel MA, Derudas B, Rigamonti E, Dievart R, Brozek J, Haulon S,
et al. PPARgamma activation primes human monocytes into alternative
M2 macrophages with anti-inflammatory properties. Cell Metab (2007)
6(2):137–43. doi:10.1016/j.cmet.2007.06.010
41. Liao X, Sharma N, Kapadia F, Zhou G, Lu Y, Hong H, et al. Kruppel-like factor 4 regulates macrophage polarization. J Clin Invest (2011) 121(7):2736–49.
doi:10.1172/JCI45444
42. Sharma N, Lu Y, Zhou G, Liao X, Kapil P, Anand P, et al. Myeloid Kruppellike factor 4 deficiency augments atherogenesis in ApoE-/- mice – brief report.
Arterioscler Thromb Vasc Biol (2012) 32(12):2836–8. doi:10.1161/ATVBAHA.
112.300471
43. Babaev VR, Yancey PG, Ryzhov SV, Kon V, Breyer MD, Magnuson MA, et al.
Conditional knockout of macrophage PPARgamma increases atherosclerosis
in C57BL/6 and low-density lipoprotein receptor-deficient mice. Arterioscler
Thromb Vasc Biol (2005) 25(8):1647–53. doi:10.1161/01.ATV.0000173413.
31789.1a
44. Martinez FO, Helming L, Milde R, Varin A, Melgert BN, Draijer C, et al. Genetic
programs expressed in resting and IL-4 alternatively activated mouse and
human macrophages: similarities and differences. Blood (2013) 121(9):e57–69.
doi:10.1182/blood-2012-06-436212
45. Shirey KA, Pletneva LM, Puche AC, Keegan AD, Prince GA, Blanco JC, et al.
Control of RSV-induced lung injury by alternatively activated macrophages
is IL-4R alpha-, TLR4-, and IFN-beta-dependent. Mucosal Immunol (2010)
3(3):291–300. doi:10.1038/mi.2010.6

November 2014 | Volume 5 | Article 614 | 236

Wang et al.

46. Luzina IG, Keegan AD, Heller NM, Rook GA, Shea-Donohue T, Atamas SP. Regulation of inflammation by interleukin-4: a review of “alternatives.” J Leukoc
Biol (2012) 92(4):753–64. doi:10.1189/jlb.0412214
47. Odegaard JI, Ricardo-Gonzalez RR, Goforth MH, Morel CR, Subramanian
V, Mukundan L, et al. Macrophage-specific PPARgamma controls alternative
activation and improves insulin resistance. Nature (2007) 447(7148):1116–20.
doi:10.1038/nature05894
48. Mukundan L, Odegaard JI, Morel CR, Heredia JE, Mwangi JW, RicardoGonzalez RR, et al. PPAR-delta senses and orchestrates clearance of apoptotic cells to promote tolerance. Nat Med (2009) 15(11):1266–72. doi:10.1038/
nm.2048
49. Eguchi J, Wang X, Yu S, Kershaw EE, Chiu PC, Dushay J, et al. Transcriptional
control of adipose lipid handling by IRF4. Cell Metab (2011) 13(3):249–59.
doi:10.1016/j.cmet.2011.02.005
50. Park-Min KH, Antoniv TT, Ivashkiv LB. Regulation of macrophage phenotype by long-term exposure to IL-10. Immunobiology (2005) 210(2–4):77–86.
doi:10.1016/j.imbio.2005.05.002
51. Imtiyaz HZ, Simon MC. Hypoxia-inducible factors as essential regulators of
inflammation. Curr Top Microbiol Immunol (2010) 345:105–20. doi:10.1007/
82_2010_74
52. Imtiyaz HZ, Williams EP, Hickey MM, Patel SA, Durham AC, Yuan LJ,
et al. Hypoxia-inducible factor 2alpha regulates macrophage function in
mouse models of acute and tumor inflammation. J Clin Invest (2010)
120(8):2699–714. doi:10.1172/JCI39506
53. Bosco MC, Puppo M, Santangelo C, Anfosso L, Pfeffer U, Fardin P, et al.
Hypoxia modifies the transcriptome of primary human monocytes: modulation of novel immune-related genes and identification of CC-chemokine
ligand 20 as a new hypoxia-inducible gene. J Immunol (2006) 177(3):1941–55.
doi:10.4049/jimmunol.177.3.1941
54. Fang HY, Hughes R, Murdoch C, Coffelt SB, Biswas SK, Harris AL, et al.
Hypoxia-inducible factors 1 and 2 are important transcriptional effectors
in primary macrophages experiencing hypoxia. Blood (2009) 114(4):844–59.
doi:10.1182/blood-2008-12-195941
55. Murdoch C, Lewis CE. Macrophage migration and gene expression in response
to tumor hypoxia. Int J Cancer (2005) 117(5):701–8. doi:10.1002/ijc.21422
56. Cramer T, Yamanishi Y, Clausen BE, Forster I, Pawlinski R, Mackman N, et al.
HIF-1alpha is essential for myeloid cell-mediated inflammation. Cell (2003)
112(5):645–57. doi:10.1016/S0092-8674(03)00154-5
57. Rius J, Guma M, Schachtrup C, Akassoglou K, Zinkernagel AS, Nizet V,
et al. NF-kappaB links innate immunity to the hypoxic response through
transcriptional regulation of HIF-1alpha. Nature (2008) 453(7196):807–11.
doi:10.1038/nature06905
58. Nizet V, Johnson RS. Interdependence of hypoxic and innate immune
responses. Nat Rev Immunol (2009) 9(9):609–17. doi:10.1038/nri2607
59. Colegio OR, Chu NQ, Szabo AL, Chu T, Rhebergen AM, Jairam V, et al. Functional polarization of tumour-associated macrophages by tumour-derived lactic acid. Nature (2014) 513(7519):559–63. doi:10.1038/nature13490
60. Tripathi C, Tewari BN, Kanchan RK, Baghel KS, Nautiyal N, Shrivastava R,
et al. Macrophages are recruited to hypoxic tumor areas and acquire a proangiogenic M2-Polarized phenotype via hypoxic cancer cell derived cytokines
Oncostatin M and Eotaxin. Oncotarget (2014) 5(14):5350–68.
61. Takeda N, O’Dea EL, Doedens A, Kim JW, Weidemann A, Stockmann C,
et al. Differential activation and antagonistic function of HIF-{alpha} isoforms in macrophages are essential for NO homeostasis. Genes Dev (2010)
24(5):491–501. doi:10.1101/gad.1881410
62. Kofler J, Wiley CA. Microglia: key innate immune cells of the brain. Toxicol
Pathol (2011) 39(1):103–14. doi:10.1177/0192623310387619
63. Vandanmagsar B, Youm YH, Ravussin A, Galgani JE, Stadler K, Mynatt RL,
et al. The NLRP3 inflammasome instigates obesity-induced inflammation and
insulin resistance. Nat Med (2011) 17(2):179–88. doi:10.1038/nm.2279
64. Tschopp J, Martinon F, Burns K. NALPs: a novel protein family involved in
inflammation. Nat Rev Mol Cell Biol (2003) 4(2):95–104. doi:10.1038/nrm1019
65. Martinon F, Burns K, Tschopp J. The inflammasome: a molecular platform
triggering activation of inflammatory caspases and processing of proIL-beta.
Mol Cell (2002) 10(2):417–26. doi:10.1016/S1097-2765(02)00599-3
66. Inohara N, Chamaillard M, McDonald C, Nuñez G. NOD-LRR proteins: role
in host-microbial interactions and inflammatory disease. Ann Rev Biochem
(2005) 74:355–83. doi:10.1146/annurev.biochem.74.082803.133347

www.frontiersin.org

Macrophage M1–M2 polarization

67. Shaw MH, Reimer T, Kim YG, Nunez G. NOD-like receptors (NLRs): bona
fide intracellular microbial sensors. Curr Opin Immunol (2008) 20(4):377–82.
doi:10.1016/j.coi.2008.06.001
68. Chamaillard M, Hashimoto M, Horie Y, Masumoto J, Qiu S, Saab L, et al.
An essential role for NOD1 in host recognition of bacterial peptidoglycan
containing diaminopimelic acid. Nat Immunol (2003) 4(7):702–7. doi:10.1038/
ni945
69. Girardin SE, Boneca IG, Viala J, Chamaillard M, Labigne A, Thomas G, et al.
Nod2 is a general sensor of peptidoglycan through muramyl dipeptide (MDP)
detection. J Biol Chem (2003) 278(11):8869–72. doi:10.1074/jbc.C200651200
70. Hasegawa M, Fujimoto Y, Lucas PC, Nakano H, Fukase K, Nunez G, et al.
A critical role of RICK/RIP2 polyubiquitination in Nod-induced NF-kappaB
activation. EMBO J (2008) 27(2):373–83. doi:10.1038/sj.emboj.7601962
71. Hitotsumatsu O, Ahmad RC, Tavares R, Wang M, Philpott D, Turer EE, et al.
The ubiquitin-editing enzyme A20 restricts nucleotide-binding oligomerization domain containing 2-triggered signals. Immunity (2008) 28(3):381–90.
doi:10.1016/j.immuni.2008.02.002
72. Keestra AM, Winter MG, Auburger JJ, Frassle SP, Xavier MN, Winter SE, et al.
Manipulation of small Rho GTPases is a pathogen-induced process detected
by NOD1. Nature (2013) 496(7444):233–7. doi:10.1038/nature12025
73. Hansen G, Hercus TR, McClure BJ, Stomski FC, Dottore M, Powell J, et al. The
structure of the GM-CSF receptor complex reveals a distinct mode of cytokine
receptor activation. Cell (2008) 134(3):496–507. doi:10.1016/j.cell.2008.05.053
74. Lehtonen A, Ahlfors H, Veckman V, Miettinen M, Lahesmaa R, Julkunen
I. Gene expression profiling during differentiation of human monocytes to
macrophages or dendritic cells. J Leukoc Biol (2007) 82(3):710–20. doi:10.1189/
jlb.0307194
75. Beyer M, Mallmann MR, Xue J, Staratschek-Jox A, Vorholt D, Krebs W, et al.
High-resolution transcriptome of human macrophages. PLoS One (2012)
7(9):e45466. doi:10.1371/journal.pone.0045466
76. Dranoff G, Jaffee E, Lazenby A, Golumbek P, Levitsky H, Brose K, et al. Vaccination with irradiated tumor cells engineered to secrete murine granulocytemacrophage colony-stimulating factor stimulates potent, specific, and longlasting anti-tumor immunity. Proc Natl Acad Sci U S A (1993) 90(8):3539–43.
doi:10.1073/pnas.90.8.3539
77. Dirksen U, Nishinakamura R, Groneck P, Hattenhorst U, Nogee L, Murray R,
et al. Human pulmonary alveolar proteinosis associated with a defect in GMCSF/IL-3/IL-5 receptor common beta chain expression. J Clin Invest (1997)
100(9):2211–7. doi:10.1172/JCI119758
78. Dirksen U, Hattenhorst U, Schneider P, Schroten H, Gobel U, Bocking
A, et al. Defective expression of granulocyte-macrophage colony-stimulating
factor/interleukin-3/interleukin-5 receptor common beta chain in children
with acute myeloid leukemia associated with respiratory failure. Blood (1998)
92(4):1097–103.
79. Baltimore D, Boldin MP, O’Connell RM, Rao DS, Taganov KD. MicroRNAs:
new regulators of immune cell development and function. Nat Immunol (2008)
9(8):839–45. doi:10.1038/ni.f.209
80. Squadrito ML, Etzrodt M, De Palma M, Pittet MJ. MicroRNA-mediated control of macrophages and its implications for cancer. Trends Immunol (2013)
34(7):350–9. doi:10.1016/j.it.2013.02.003
81. Graff JW, Dickson AM, Clay G, McCaffrey AP, Wilson ME. Identifying functional microRNAs in macrophages with polarized phenotypes. J Biol Chem
(2012) 287(26):21816–25. doi:10.1074/jbc.M111.327031
82. O’Connell RM, Rao DS, Chaudhuri AA, Boldin MP, Taganov KD, Nicoll J, et al.
Sustained expression of microRNA-155 in hematopoietic stem cells causes a
myeloproliferative disorder. J Exp Med (2008) 205(3):585–94. doi:10.1084/jem.
20072108
83. Taganov KD, Boldin MP, Chang KJ, Baltimore D. NF-kappaB-dependent induction of microRNA miR-146, an inhibitor targeted to signaling proteins of
innate immune responses. Proc Natl Acad Sci U S A (2006) 103(33):12481–6.
doi:10.1073/pnas.0605298103
84. Barish GD, Yu RT, Karunasiri M, Ocampo CB, Dixon J, Benner C, et al. Bcl-6
and NF-kappaB cistromes mediate opposing regulation of the innate immune
response. Genes Dev (2010) 24(24):2760–5. doi:10.1101/gad.1998010
85. Tili E, Michaille JJ, Cimino A, Costinean S, Dumitru CD, Adair B, et al. Modulation of miR-155 and miR-125b levels following lipopolysaccharide/TNF-alpha
stimulation and their possible roles in regulating the response to endotoxin
shock. J Immunol (2007) 179(8):5082–9. doi:10.4049/jimmunol.179.8.5082

November 2014 | Volume 5 | Article 614 | 237

Wang et al.

86. Bazzoni F, Rossato M, Fabbri M, Gaudiosi D, Mirolo M, Mori L, et al.
Induction and regulatory function of miR-9 in human monocytes and neutrophils exposed to proinflammatory signals. Proc Natl Acad Sci U S A (2009)
106(13):5282–7. doi:10.1073/pnas.0810909106
87. Zhu D, Pan C, Li L, Bian Z, Lv Z, Shi L, et al. MicroRNA-17/20a/106a modulate macrophage inflammatory responses through targeting signal-regulatory
protein alpha. J Allergy Clin Immunol (2013) 132(2):426e–36e. doi:10.1016/j.
jaci.2013.02.005
88. Liu Y, Chen Q, Song Y, Lai L, Wang J, Yu H, et al. MicroRNA-98 negatively
regulates IL-10 production and endotoxin tolerance in macrophages after
LPS stimulation. FEBS Lett (2011) 585(12):1963–8. doi:10.1016/j.febslet.2011.
05.029
89. Sheedy FJ, Palsson-McDermott E, Hennessy EJ, Martin C, O’Leary JJ, Ruan
Q, et al. Negative regulation of TLR4 via targeting of the proinflammatory
tumor suppressor PDCD4 by the microRNA miR-21. Nat Immunol (2010)
11(2):141–7. doi:10.1038/ni.1828
90. Nahid MA, Satoh M, Chan EK. Mechanistic role of microRNA-146a in
endotoxin-induced differential cross-regulation of TLR signaling. J Immunol
(2011) 186(3):1723–34. doi:10.4049/jimmunol.1002311
91. Ponomarev ED, Veremeyko T, Barteneva N, Krichevsky AM, Weiner HL.
MicroRNA-124 promotes microglia quiescence and suppresses EAE by deactivating macrophages via the C/EBP-alpha-PU.1 pathway. Nat Med (2011)
17(1):64–70. doi:10.1038/nm.2266
92. Zhuang G, Meng C, Guo X, Cheruku PS, Shi L, Xu H, et al. A novel
regulator of macrophage activation: miR-223 in obesity-associated adipose tissue inflammation. Circulation (2012) 125(23):2892–903. doi:10.1161/
CIRCULATIONAHA.111.087817
93. Banerjee S, Xie N, Cui H, Tan Z, Yang S, Icyuz M, et al. MicroRNA let7c regulates macrophage polarization. J Immunol (2013) 190(12):6542–9.
doi:10.4049/jimmunol.1202496
94. Lu YC, Kim I, Lye E, Shen F, Suzuki N, Suzuki S, et al. Differential role
for c-Rel and C/EBPbeta/delta in TLR-mediated induction of proinflammatory cytokines. J Immunol (2009) 182(11):7212–21. doi:10.4049/jimmunol.
0802971
95. Litvak V, Ramsey SA, Rust AG, Zak DE, Kennedy KA, Lampano AE, et al. Function of C/EBPdelta in a regulatory circuit that discriminates between transient and persistent TLR4-induced signals. Nat Immunol (2009) 10(4):437–43.
doi:10.1038/ni.1721
96. Martinez-Nunez RT, Louafi F, Sanchez-Elsner T. The interleukin 13 (IL-13)
pathway in human macrophages is modulated by microRNA-155 via direct
targeting of interleukin 13 receptor alpha1 (IL13Ralpha1). J Biol Chem (2011)
286(3):1786–94. doi:10.1074/jbc.M110.169367
97. Cai X, Yin Y, Li N, Zhu D, Zhang J, Zhang CY, et al. Re-polarization of tumorassociated macrophages to pro-inflammatory M1 macrophages by microRNA155. J Mol Cell Biol (2012) 4(5):341–3. doi:10.1093/jmcb/mjs044
98. Pedersen IM, Otero D, Kao E, Miletic AV, Hother C, Ralfkiaer E, et al.
Onco-miR-155 targets SHIP1 to promote TNFalpha-dependent growth of B
cell lymphomas. EMBO Mol Med. (2009) 1(5):288–95. doi:10.1002/emmm.
200900028
99. Doyle SL, Jefferies CA, O’Neill LA. Bruton’s tyrosine kinase is involved in
p65-mediated transactivation and phosphorylation of p65 on serine 536
during NFkappaB activation by lipopolysaccharide. J Biol Chem (2005)
280(25):23496–501. doi:10.1074/jbc.C500053200
100. Doyle SL, Jefferies CA, Feighery C, O’Neill LA. Signaling by Toll-like receptors 8
and 9 requires Bruton’s tyrosine kinase. J Biol Chem (2007) 282(51):36953–60.
doi:10.1074/jbc.M707682200
101. Horwood NJ, Mahon T, McDaid JP, Campbell J, Mano H, Brennan FM,
et al. Bruton’s tyrosine kinase is required for lipopolysaccharide-induced
tumor necrosis factor alpha production. J Exp Med (2003) 197(12):1603–11.
doi:10.1084/jem.20021845
102. Horwood NJ, Page TH, McDaid JP, Palmer CD, Campbell J, Mahon T, et al.
Bruton’s tyrosine kinase is required for TLR2 and TLR4-induced TNF, but not
IL-6, production. J Immunol (2006) 176(6):3635–41. doi:10.4049/jimmunol.
176.6.3635
103. Ni Gabhann J, Hams E, Smith S, Wynne C, Byrne JC, Brennan K, et al. Btk regulates macrophage polarization in response to lipopolysaccharide. PLoS One
(2014) 9(1):e85834. doi:10.1371/journal.pone.0085834

Frontiers in Immunology | Molecular Innate Immunity

Macrophage M1–M2 polarization

104. Lee KG, Xu S, Kang ZH, Huo J, Huang M, Liu D, et al. Bruton’s tyrosine kinase
phosphorylates Toll-like receptor 3 to initiate antiviral response. Proc Natl Acad
Sci U S A (2012) 109(15):5791–6. doi:10.1073/pnas.1119238109
105. Schlaepfer E, Rochat MA, Duo L, Speck RF. Triggering TLR2, 3, 4, 5 and 8 reinforces the restrictive nature of M1- and M2-polarized macrophages to HIV.
J Virol (2014) 88(17):9769–81. doi:10.1128/JVI.01053-14
106. Biswas SK, Gangi L, Paul S, Schioppa T, Saccani A, Sironi M, et al. A distinct and
unique transcriptional program expressed by tumor-associated macrophages
(defective NF-kappaB and enhanced IRF-3/STAT1 activation). Blood (2006)
107(5):2112–22. doi:10.1182/blood-2005-01-0428
107. Porta C, Rimoldi M, Raes G, Brys L, Ghezzi P, Di Liberto D, et al. Tolerance and
M2 (alternative) macrophage polarization are related processes orchestrated by
p50 nuclear factor kappaB. Proc Natl Acad Sci U S A (2009) 106(35):14978–83.
doi:10.1073/pnas.0809784106
108. Garofalo RS, Orena SJ, Rafidi K, Torchia AJ, Stock JL, Hildebrandt AL, et al.
Severe diabetes, age-dependent loss of adipose tissue, and mild growth deficiency in mice lacking Akt2/PKB beta. J Clin Invest (2003) 112(2):197–208.
doi:10.1172/JCI16885
109. Kadl A, Meher AK, Sharma PR, Lee MY, Doran AC, Johnstone SR, et al. Identification of a novel macrophage phenotype that develops in response to atherogenic phospholipids via Nrf2. Circ Res (2010) 107(6):737–46. doi:10.1161/
CIRCRESAHA.109.215715
110. Umemura N, Saio M, Suwa T, Kitoh Y, Bai J, Nonaka K, et al. Tumorinfiltrating myeloid-derived suppressor cells are pleiotropic-inflamed monocytes/macrophages that bear M1- and M2-type characteristics. J Leukoc Biol
(2008) 83(5):1136–44. doi:10.1189/jlb.0907611
111. Carpenter S, Aiello D, Atianand MK, Ricci EP, Gandhi P, Hall LL, et al. A long
noncoding RNA mediates both activation and repression of immune response
genes. Science (2013) 341(6147):789–92. doi:10.1126/science.1240925
112. Li Z, Chao TC, Chang KY, Lin N, Patil VS, Shimizu C, et al. The long
noncoding RNA THRIL regulates TNFalpha expression through its interaction with hnRNPL. Proc Natl Acad Sci U S A (2014) 111(3):1002–7.
doi:10.1073/pnas.1313768111
113. Mylonas KJ, Nair MG, Prieto-Lafuente L, Paape D, Allen JE. Alternatively
activated macrophages elicited by helminth infection can be reprogrammed
to enable microbial killing. J Immunol (2009) 182(5):3084–94. doi:10.4049/
jimmunol.0803463
114. Stout RD, Jiang C, Matta B, Tietzel I,Watkins SK, Suttles J. Macrophages sequentially change their functional phenotype in response to changes in microenvironmental influences. J Immunol (2005) 175(1):342–9. doi:10.4049/jimmunol.
175.1.342
115. Hyam SR, Lee IA, Gu W, Kim KA, Jeong JJ, Jang SE, et al. Arctigenin ameliorates
inflammation in vitro and in vivo by inhibiting the PI3K/AKT pathway and
polarizing M1 macrophages to M2-like macrophages. Eur J Pharmacol (2013)
708(1–3):21–9. doi:10.1016/j.ejphar.2013.01.014
116. Jang SE, Hyam SR, Han MJ, Kim SY, Lee BG, Kim DH. Lactobacillus brevis
G-101 ameliorates colitis in mice by inhibiting NF-kappaB, MAPK and AKT
pathways and by polarizing M1 macrophages to M2-like macrophages. J Appl
Microbiol (2013) 115(3):888–96. doi:10.1111/jam.12273
Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 14 August 2014; accepted: 14 November 2014; published online: 28 November
2014.
Citation: Wang N, Liang H and Zen K (2014) Molecular mechanisms that influence the macrophage M1–M2 polarization balance. Front. Immunol. 5:614. doi:
10.3389/fimmu.2014.00614
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Wang , Liang and Zen. This is an open-access article distributed
under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s)
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OPINION ARTICLE
published: 28 July 2014
doi: 10.3389/fimmu.2014.00357

SOCS proteins in macrophage polarization and function
Heather M. Wilson*
Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK
*Correspondence: h.m.wilson@abdn.ac.uk
Edited by:
Laurel L. Lenz, National Jewish Health, USA
Reviewed by:
Masato Kubo, Tokyo University of Science, Japan
Heiko Mühl, University Hospital Goethe University Frankfurt, Germany
Keywords: suppressor of cytokine signaling proteins, macrophage, M1, M2, inflammation

INTRODUCTION
Macrophages were initially described as
“big eaters” due to their phagocytic nature.
It is now clear that macrophages have
many diverse functions not only in innate
immunity and tissue homeostasis but also
in metabolism, development, and regeneration. Macrophage functions are driven largely by tissue-derived and pathogenic microenvironmental stimuli that
help them adapt to changing conditions within tissues and tailor an appropriate response. The heterogeneity of
macrophages has resulted in their classification into subtypes based on their phenotype and function (1). One major classification, based on function, is M1 and M2
macrophages, with destructive and healing
properties, respectively (2, 3). As imbalances between M1 and M2 states have been
observed in a number of diseases, an understanding of the molecular mechanisms, signaling pathways, and transcription factors
controlling their polarization has obvious
therapeutic implications. Recent studies
have established strong potential for suppressor of cytokine signaling (SOCS) proteins to regulate M1 and M2 macrophage
polarization (4–7). Here, the focus will be
on the evidence for this, and the consequences of altered SOCS expressions on
macrophage function in health and disease. Overall it is proposed that a high
SOCS1 to SOCS3 ratio could be a potential marker for M2 macrophages while
high SOCS3 expression is associated with
M1 cells.

SOCS PROTEINS
Suppressor of cytokine signaling proteins
are a family of intracellular cytokineinducible proteins, consisting of eight
members (CIS and SOCS1–SOCS7) (8, 9).

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SOCS1 and SOCS3 are most widely
characterized regarding their roles in
shaping M1 and M2 macrophage polarization (4–6). They show low expression
in resting macrophages, but are rapidly
induced on activation. All SOCS family
proteins contain an Src homology 2 (SH2)
domain, a variable length amino-terminal
domain and a conserved carboxy-terminal
SOCS box motif that interacts with
ubiquitin–ligase machinery (8, 9). SOCS
are induced by a variety of stimuli that
cause M1 and M2 activation, including
cytokines, toll-like receptor (TLR) ligands, angiotensin II, immune complexes,
and high glucose (9). The most studied
signaling pathway regulated by SOCS is
JAK/STAT activation. SOCS negatively
regulate JAK/STAT signaling through association with key phosphorylated tyrosine
residues on JAK proteins and/or cytokine
receptors, and by degradation of signaling
molecules mediated via the ubiquitin–
proteasome pathway (8, 9). SOCS1 and
SOCS3 contain a kinase inhibitory region
(KIR) that directly suppresses JAK tyrosine kinase activity. SOCS proteins also
influence ERK (10), PI3K (11), Notch (12),
MAPK (13), and NF-κB (14) signaling
cascades that directs M1 and M2 functions.
SOCS1

SOCS1 regulates M1-macrophage activation by inhibiting the interferon
gamma-induced JAK2/STAT1 pathway and
TLR/NF-κB signaling (9, 15) (Figure 1). To
suppress the latter pathway, SOCS1 binds
to the p65 subunit of NF-κB and the TLR
adaptor molecule Mal/TIRAP as well as
IRAK, facilitating its ubiquitin-mediated
proteolysis via ubiquitin ligases recruited
by the SOCS box (8, 14–17). SOCS1
indirectly inhibits TLR4 signaling through

secondary mechanisms targeting IRF3 and
IFN-β induced JAK/STAT pathways (18,
19). Thus, SOCS1 mediates a negative feedback mechanism during TLR4 signaling,
via control of both MyD88-dependent and
MyD88-independent signaling. SOCS1deficient mice succumb to severe systemic
autoimmune and inflammatory disease
(14, 16) and their M1-macrophages display
an increased capacity to kill intracellular
bacterial pathogens, presumably due to
unrestrained IFN-γ/STAT1 and p65 signaling. In line with this, SOCS1 knockout
or knockdown M1-activated macrophages
show enhanced levels of IL-6, IL-12, MHC
class II, and nitric oxide suggesting SOCS1
sustains the properties of M1 macrophages
at a less destructive level to prevent overshooting inflammatory responses (4, 18).
This explains why SOCS1 promoter hypermethylation, which results in loss of
SOCS1 expression leads to enhanced secretion of lipopolysaccharide (LPS)-induced
pro-inflammatory cytokines (20). Micro
RNA-155 (miR-155) is a critical regulator
of innate immunity and TLR signaling
(21–23); miR-155 targets and degrades
SOCS1 in M1-activated macrophages (21),
thus miR-155 induction during activation serves to maximize and extend the
inflammatory process.
SOCS1 also regulates M2 macrophage
polarization. Expression of macrophage
SOCS1, but not SOCS3, is strongly upregulated in an M2 polarizing environment in vitro and in vivo, where it has
an important role in acquisition of M2
functional characteristics, such as a high
arginase I/low inducible nitric oxide synthase (iNOS) expression ratio (4). Strikingly, this contrasts with macrophages infiltrating an in vivo inflamed M1-activating
environment, where macrophages with

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SOCS proteins and macrophage polarization

FIGURE 1 | Role of SOCS1 and SOCS3 in macrophage activation. STAT1 and NFκB drive M1
polarization and SOCS1 can inhibit these pathways. SOCS3 can regulate TLR signaling and inhibits
IL-6-induced STAT3 activation and SMAD3 and PI3K activity to action an appropriate destructive effect.
STAT3, STAT6, and PI3K can drive M2 activation and SOCS3 inhibits STAT3 and PI3K. Pathways that
trigger SOCS1 in macrophages include STAT1 and NFκB, while SOCS3 expression can be induced by
STAT3, NFκB, NOTCH1, PI3K, and MAP kinase activation.

enhanced SOCS3 but not SOCS1 expression are prominent (5). This suggests
that exclusive upregulation of SOCS1, or
indeed, a high SOCS1/SOCS3 expression
ratio, has potential as a useful and additional in vivo biomarker for M2 (see
later). Arginase I expression, as an M2
macrophage marker, can be mediated via
activation of either STAT6 (24) or PI3K
(25). SOCS1 is important in controlling PI3K activity, supporting a mechanism for regulating arginase I expression in M2 cells; SOCS1 also regulates
STAT6 phosphorylation (26). Following
activation, SOCS1 knockdown or SOCS1deficient macrophages show a reciprocal upregulation of SOCS3 expression.
SOCS3 inhibits PI3K activation (27), and
so the expression of high SOCS1 and
low SOCS3 in M2 macrophages could
result in greater PI3K activity and more
arginase I induction in these cells. An
elevated expression of SOCS1 is important for the arginase I-induced suppressive nature of M2 macrophages that
attenuate lymphocyte proliferation (28).
Moreover, siRNA-mediated knockdown of
SOCS1 results in the induction of iNOS
in IL-4-pretreated cells stimulated with
IFN/LPS (4). Thus, SOCS1 regulates the
iNOS/arginase I expression ratio in both
M1 and M2 macrophages and helps finetune key signaling pathways to mount an
appropriate response to changes within the
microenvironment.

SOCS2

An important role for SOCS2 in driving M2 polarization and limiting M1
polarization has been shown, with IL4 activation of macrophages, resulting
in enhanced SOCS2 expression (27).
Macrophages from SOCS2-/- mice display
increased secretion of IFN-γ, IL-1β, and
TNF-α in response to LPS in parallel to
an increased pro-inflammatory cytokine
mRNA expression (29). These BMDMs
have higher basal levels of p65–NF-κB
compared with macrophages from wildtype mice (29). In another study, SOCS2deficient macrophages were hyper responsive to IFN-γ, produced more NO and
dealt with infection more efficiently (30).
SOCS2 has also been described as a feedback inhibitor of TLR-induced activation
in dendritic cells (31).
SOCS3

In contrast with SOCS2, a key role
for SOCS3 in M1 polarization is proposed (Figure 1). The majority of
macrophages activated within an in vivo
pro-inflammatory conditioning environment show strong upregulation of SOCS3
expression and this cell population
co-express the M1 marker, iNOS (5,
6). Without SOCS3, both human and
rodent macrophages have a reduced ability to develop pro-inflammatory features but instead display immunoregulatory characteristics (5, 6). Notably, mice

Frontiers in Immunology | Molecular Innate Immunity

with a targeted deletion of SOCS3 in
macrophages and neutrophils demonstrate
a reduced IL-12 response and succumb
to toxoplasmosis (32). SOCS3 binds to
and inhibits gp130-related cytokine receptors and consequently this abrogates IL-6induced STAT1 gene expression and IL-6induced STAT3 anti-inflammatory effects
(33–35). Therefore, in SOCS3-deficient
macrophages, IL-6 signals in a similar manner to the immunosuppressive cytokine
IL-10, through prolonged STAT3 activation and dampening of LPS signaling (33).
As a result, mice deficient in SOCS3 in
myeloid cells are resistant to endotoxic
shock (35) with reduced production of
pro-inflammatory cytokines. However, one
report in the same mice suggests SOCS3
deficiency promotes M1 macrophage activation in spite of enhanced STAT3 activation (7). The reasons for this discrepancy in findings are unclear but could
relate to differences in dose and purity
of the LPS used in the different studies,
as well as and the genes and time-points
analyzed after macrophage activation (7,
35). Moreover, the conflicting results for
the role of SOCS3 in M1 polarization in
isolated macrophages in vitro (5–7) could
result from the different technologies and
species used (siRNA-mediated knockdown
in rat and human macrophages, which
avoids the risk of compensatory effects
of other SOCS genes (5, 6) versus cells
from macrophage-specific SOCS3 knockout mice) (7). Resolving these issues should
establish the importance of SOCS3 in modulating macrophage function in vivo.
Studies
of
SOCS3-deficient
macrophages confirm that SOCS3 positively regulates TLR4 signaling and
M1 activation by inhibition of IL-6Rmediated STAT3 activation, as well as
TGF-β-mediated SMAD3 activation,
which is critical for the negative regulation
of TLR-induced TNF-α and IL-6 production (5, 6, 33, 36). Since SOCS3 blocks PI3K
that feeds and inhibits TLR responses, this
could be an alternative mechanism by
which SOCS3 augments TLR signaling
in M1 macrophages (6). Forced activation of Notch signaling enhances both
M1 polarization and anti-tumor activity via SOCS3 induction (12). In line
with this, macrophage-specific SOCS3
knockout animals are resistant to tumor
transplantation due to reduced secretion

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of tumor-promoting TNF-α and IL-6,
together with elevated MCP2/CCL8 that is
anti-tumorigenic (37).
Regulation of SOCS3 in innate cells
influences downstream T cell fates. The
presence of SOCS3 in macrophages is
important in fine-tuning downstream T
effector cell priming due to both influences in expression of presenting molecules and altered secretion of T cell polarizing cytokines (6, 7). Mouse SOCS3deficient dendritic cells display an analogous reduced potential to drive T effector
cell responses and a tolerogenic phenotype as a result of enhanced TGFβ production and expansion of Foxp3-positive
regulatory T cells (38). These dendritic
cells reduce the severity of experimental autoimmune disease. Therefore, regulation of intracellular signaling pathways by
SOCS3 in innate cells is critical for the decision of adaptive responses such as T cell
fates. The depletion of macrophage SOCS3
in a clinical situation would thus be predicted to dampen both pro-inflammatory
innate and adaptive immune responses.
The above studies suggest that
macrophage SOCS3 is associated with
M1 macrophages and pro-inflammatory
responses and is a potential therapeutic
target in inflammatory diseases. However,
a word of caution should be introduced as
this may not be the case in all inflammatory conditions. In diseases, where STAT3
activation exerts a profound inflammatory
and pathogenic response (39, 40) then the
effects of SOCS3 targeting may not be
beneficial. For example, in an IL-1/STAT3
model of chronic arthritis where SOCS3
was deleted in hematopoietic and endothelial cells, animals exhibited more severe
disease. Thus, the pathology needs first to
be assessed before SOCS3 manipulation as
a therapy is considered (37).

MACROPHAGE SOCS EXPRESSION
AND PATHOLOGY
The heightened expression of macrophage
SOCS1 and SOCS3 proteins have been
demonstrated in many pathologies in vivo
where this has been proposed, through the
molecular mechanisms described above, to
enhance or inhibit pathogenesis.
SOCS AND GLOMERULONEPHRITIS

Macrophages are an important feature in glomerulonephritis pathology.

www.frontiersin.org

SOCS proteins and macrophage polarization

Macrophages
infiltrating
inflamed
glomeruli in experimental models are
rapidly polarized to express either SOCS1
or SOCS3, but rarely both, with most
exclusively expressing SOCS3 (5, 6). The
proportion of these SOCS3-expressing
macrophages correlates strongly with the
severity of immune-mediated injury. Local
delivery of IL-4 to inflamed glomeruli
has a major effect on reducing the number of SOCS3-expressing glomerular
macrophages, and this is reflected by
a decrease in the severity of nephritis,
supporting a role for SOCS3 in driving
M1-mediated injury (5).
SOCS AND ATHEROSCLEROSIS

Human atherosclerotic plaques exhibit a
high expression of macrophage SOCS1 and
SOCS3 in unstable inflammatory shoulder regions as compared to stable fibrous
area (41). SOCS1 and SOCS3 expression
is increased in aortic lesion macrophages
from apoE(−/−) mice (42). In human tissue, the percentages of SOCS1-positive,
M2 macrophages are decreased in morphologically stable atherosclerotic plaques,
whereas percentages of SOCS3-positive,
iNOS positive, macrophages are increased
in unstable, rupture-prone plaques, suggesting targeting macrophage SOCS3
would be beneficial to dampen inflammation and plaque vulnerability (43). The differing expression ratio of SOCS1:SOCS3
in atherosclerotic plaques again suggests
that the ratio could be an indicator
of the inflammatory status of human
macrophages in vivo. SOCS1 was atheroprotective in mouse models (44) while
the absence of macrophage SOCS3 of
apoE(−/−) mice attenuates disease, confirming a causal link between macrophage
SOCS3 and atherosclerosis (45).

inflammation associated with IBD. PPARγ
expression is upregulated in M2 but not
M1 macrophages. In macrophages lacking PPARγ, a significant upregulation of
SOCS3 was noted and this could be important if treating IBD with PPARγ agonists (47).
SOCS AND TUMORS

In human tumors, SOCS3 expression
identifies macrophages with enhanced
tumor killing, whereas SOCS1 expressing macrophages (M2) favor tumor survival (48). Macrophage-specific deletion
of SOCS1 leads to reduced susceptibility to melanoma growth and colon carcinogenesis through increased anti-tumor
responses (49) and a switch to M1 polarization of tumor-associated macrophage.
In contrast, mice with a macrophagespecific deletion of SOCS3, subcutaneously
implanted with melanoma cells, did not
show a difference in tumor size, although
the number of metastasis increased in
these mice (37). These SOCS3-deficient
macrophages produce less IL-6 and TNF-α
upon stimulation with tumor lysates due
to aberrant STAT3 activity, again showing
a positive link of SOCS and macrophage
polarization (37).
SOCS AND OBESITY

SOCS3 restrains macrophage responses to
IL-6 and leptin that are systemically upregulated in obesity (50). SOCS1 inhibits
insulin signaling and macrophage cytokine
secretion, resulting in insulin sensitivity
in spite of an obese state (17). Moreover, an increase in SOCS1 expression
in mouse macrophages inhibits LPS- and
palmitate-induced TLR4 signaling and in
so doing prevents systemic inflammation
and hepatic insulin resistance (17).

SOCS AND INFLAMMATORY BOWEL DISEASE

CONCLUSION AND PERSPECTIVES

Beneath the gut epithelia, lamina propria macrophages phagocytose bacteria
and maintain an M2 phenotype in the
steady state. Approximately 10% of these
macrophages express SOCS3 in healthy
individuals, whereas in inflammatory
bowel disease (IBD) patients this increases
to 40%, again suggesting SOCS3 expression relates to M1-activated macrophages
(46). Peroxisome proliferator-activated
receptor-γ (PPARγ) agonists demonstrate efficacy in ameliorating intestinal

Given the broad role of SOCS in regulating macrophage functions in health and
disease, the modulation of macrophagespecific SOCS1 and SOCS3 expression provides new opportunities for therapeutic
manipulation of immune and inflammatory responses. However, it is not only
macrophages that are affected by SOCS
proteins. Other cell types upregulate and
react to SOCS proteins to shape cellular
functions. Targeting SOCS specifically in
macrophages is therefore important as an

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Wilson

efficient means of changing the inflammatory response.

REFERENCES
1. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat
Rev Immunol (2011) 11:723–37. doi:10.1038/
nri3073
2. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill
AM. M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol (2000) 164:6166–73. doi:10.
4049/jimmunol.164.12.6166
3. Sica A, Mantovani A. Macrophage plasticity and
polarization: in vivo veritas. J Clin Invest (2012)
122:787–95. doi:10.1172/JCI59643
4. Whyte CS, Bishop ET, Rückerl D, Gaspar-Pereira S,
Barker RN, Allen JE, et al. Suppressor of cytokine
signaling (SOCS)1 is a key determinant of differential macrophage activation and function.
J Leukoc Biol (2011) 90:845–54. doi:10.1189/jlb.
1110644
5. Liu Y, Stewart KN, Bishop ET, Marek CM, Kluth
DC, Rees AJ, et al. Unique expression of suppressor of cytokine signaling 3 is essential for classical macrophage activation in rodents in vitro
and in vivo. J Immunol (2008) 180:6270–8. doi:
10.4049/jimmunol.180.9.6270
6. Arnold CE, Whyte CS, Gordon P, Barker RN,
Rees AJ, Wilson HM. A critical role for suppressor of cytokine signalling 3 in promoting M1
macrophage activation and function in vitro and
in vivo. Immunology (2014) 141:96–110. doi:10.
1111/imm.12173
7. Qin H, Holdbrooks AT, Liu Y, Reynolds SL, Yanagisawa LL, Benveniste EN. SOCS3 deficiency promotes M1 macrophage polarization and inflammation. J Immunol (2012) 189:3439–48. doi:10.
4049/jimmunol.1201168
8. Strebovsky J, Walker P, Dalpke AH. Suppressor of
cytokine signaling proteins as regulators of innate
immune signaling. Front Biosci (2012) 17:1627–39.
9. Yoshimura A, Naka T, Kubo M. SOCS proteins, cytokine signaling, and immune regulation.
Nat Rev Immunol (2007) 7:454–65. doi:10.1038/
nri2093
10. Madonna S, Scarponi C, De Pità O, Albanesi C.
Suppressor of cytokine signaling 1 inhibits IFNgamma inflammatory signaling in human keratinocytes by sustaining ERK1/2 activation. FASEB
J (2008) 22:3287–97. doi:10.1096/fj.08-106831
11. Lu Y, Fukuyama S, Yoshida R, Kobayashi T, Saeki K,
Shiraishi H, et al. Loss of SOCS3 gene expression
converts STAT3 function from anti-apoptotic to
pro-apoptotic. J Biol Chem (2006) 281:36683–90.
doi:10.1074/jbc.M607374200
12. Wang YC, He F, Feng F, Liu XW, Dong GY, Qin
HY. Notch signaling determines the M1 versus
M2 polarization of macrophages in antitumor
immune responses. Cancer Res (2010) 70:4840–9.
doi:10.1158/0008-5472.CAN-10-0269
13. Puhr M, Santer FR, Neuwirt H, Marcias G, Hobisch
A, Culig Z. SOCS-3 antagonises the proliferative
and migratory effects of fibroblast growth factor-2
in prostate cancer by inhibition of p44/p42 MAPK
signalling. Endocr Relat Cancer (2010) 17:525–38.
doi:10.1677/ERC-10-0007

SOCS proteins and macrophage polarization

14. Nakagawa R, Naka T, Tsutsui H, Fujimoto M,
Kimura A, Abe T, et al. SOCS-1 participates in
negative regulation of LPS responses. Immunity
(2002) 17:677–87. doi:10.1016/S1074-7613(02)
00449-1
15. Ryo A, Suizu F, Yoshida Y, Perrem K, Liou YC,
Wulf G, et al. Regulation of NF-kappaB signaling by Pin1-dependent prolyl isomerization
and ubiquitin-mediated proteolysis of p65/RelA.
Mol Cell (2003) 12:1413–26. doi:10.1016/S10972765(03)00490-8
16. Kinjyo I, Hanada T, Inagaki-Ohara K, Mori H,
Aki D, Ohishi M, et al. SOCS1/JAB is a negative
regulator of LPS-induced macrophage activation.
Immunity (2002) 17:583–91. doi:10.1016/S10747613(02)00446-6
17. Sachithanandan N, Graham KL, Galic S, Honeyman JE, Fynch SL, Hewitt KA, et al. Macrophage
deletion of SOCS1 increases sensitivity to LPS and
palmitic acid and results in systemic inflammation and hepatic insulin resistance. Diabetes (2011)
60:2023–31. doi:10.2337/db11-0259
18. Baetz A, Frey M, Heeg K, Dalpke AH. Suppressor of cytokine signaling (SOCS) proteins indirectly regulate toll-like receptor signaling in innate
immune cells. J Biol Chem (2004) 279:54708–15.
doi:10.1074/jbc.M410992200
19. Gingras S, Parganas E, de Pauw A, Ihle JN, Murray PJ. Re-examination of the role of suppressor
of cytokine signaling 1 (SOCS1) in the regulation
of toll-like receptor signaling. J Biol Chem (2004)
279:54702–7. doi:10.1074/jbc.M411043200
20. Cheng C, Huang C, Ma TT, Bian EB, He Y, Zhang
L, et al. SOCS1 hypermethylation mediated by
DNMT1 is associated with lipopolysaccharideinduced inflammatory cytokines in macrophages.
Toxicol Lett (2014) 225:488–97. doi:10.1016/j.
toxlet.2013.12.023
21. O’Connell RM, Taganov KD, Boldin MP, Cheng
G, Baltimore D. MicroRNA-155 is induced during
the macrophage inflammatory response. Proc Natl
Acad Sci U S A (2007) 104:1604–9. doi:10.1073/
pnas.0610731104
22. Ruggiero T, Trabucchi M, De Santa F, Zupo S, Harfe
BD, McManus MT, et al. LPS induces KH-type
splicing regulatory protein-dependent processing
of microRNA-155 precursors in macrophages.
FASEB J (2009) 23:2898–908. doi:10.1096/fj.09131342
23. Androulidaki A, Iliopoulos D, Arranz A, Doxaki
C, Schworer S, Zacharioudaki V. The kinase Akt1
controls macrophage response to lipopolysaccharide by regulating microRNAs. Immunity (2009)
31:220–31. doi:10.1016/j.immuni.2009.06.024
24. Pauleau AL, Rutschman R, Lang R, Pernis A,
Watowich SS, Murray PJ. Enhancer-mediated control of macrophage-specific arginase I expression. J Immunol (2004) 172:7565–73. doi:10.4049/
jimmunol.172.12.7565
25. Sly LM, Rauh MJ, Kalesnikoff J, Song CH, Krystal G. LPS-induced upregulation of SHIP is essential for endotoxin tolerance. Immunity (2004)
21:227–39. doi:10.1016/j.immuni.2004.07.010
26. Dickensheets HN, Vazquez N, Sheikh F, Gingras S, Murray PJ, Ryan JJ, et al. Suppressor of
cytokine signaling-1 is an IL-4-inducible gene in
macrophages and feedback inhibits IL-4 signaling.

Frontiers in Immunology | Molecular Innate Immunity

Genes Immun (2007) 8:21–7. doi:10.1038/sj.gene.
6364352
27. O’Connor JC, Sherry CL, Guest CB, Freund GG. Type 2 diabetes impairs insulin
receptor substrate-2-mediated phosphatidylinositol 3-kinase activity in primary macrophages
to induce a state of cytokine resistance to IL4 in association with overexpression of suppressor of cytokine signaling-3. J Immunol (2007)
178:6886–93. doi:10.4049/jimmunol.178.11.6886
28. Pesce JT, Ramalingam TR, Mentink-Kane MM,
Wilson MS, El Kasmi KC, Smith AM, et al.
Arginase-1-expressing macrophages suppress Th2
cytokine-driven inflammation and fibrosis. PLoS
Pathog (2009) 5:e1000371. doi:10.1371/journal.
ppat.1000371
29. Esper L, Roman-Campos D, Lara A, Brant F, Castro LL, Barroso A, et al. Role of SOCS2 in modulating heart damage and function in a murine
model of acute Chagas disease. Am J Pathol (2012)
181:130–40. doi:10.1016/j.ajpath.2012.03.042
30. Zadjali F, Santana-Farre R, Vesterlund M, Carow
B, Mirecki-Garrido M, Hernandez-Hernandez I,
et al. In SOCS2 KO mice, there was a reduction in
both parasitemia and the expression of interferonγ (IFN-γ), tumor necrosis factor-α (TNF-α), IL-6,
IL-10, SOCS1, and SOCS3 in the spleen. FASEB J
(2012) 26:3282–91. doi:10.1096/fj.12-205583
31. Posselt G, Schwarz H, Duschl A, HorejsHoeck J. Suppressor of cytokine signaling 2
is a feedback inhibitor of TLR-induced activation in human monocyte-derived dendritic
cells. J Immunol (2011) 187:2875–84. doi:10.4049/
jimmunol.1003348
32. Whitmarsh RJ, Gray CM, Gregg B, Christian DA,
May MJ, Murray PJ, et al. A critical role for SOCS3
in innate resistance to Toxoplasma gondii. Cell Host
Microbe (2011) 10:224–36. doi:10.1016/j.chom.
2011.07.009
33. Lang R, Pauleau AL, Parganas E, Takahashi Y,
Mages J, Ihle JN. SOCS3 regulates the plasticity
of gp130 signaling. Nat Immunol (2003) 4:546–50.
doi:10.1038/ni932
34. Croker BA, Krebs DL, Zhang JG, Wormald S, Willson TA, Stanley EG, et al. Negatively regulates IL6 signaling in vivo. Nat Immunol (2003) 4:540–5.
doi:10.1038/ni931
35. Yasukawa H, Ohishi M, Mori H, Murakami M, Chinen T, Aki D. IL-6 induces an anti-inflammatory
response in the absence of SOCS3 in macrophages.
Nat Immunol (2003) 4:551–6. doi:10.1038/ni938
36. Liu X, Zhang Y, Yu Y, Yang X, Cao X. SOCS3
promotes TLR4 response in macrophages by
feedback inhibiting TGF-beta1/Smad3 signaling.
Mol Immunol (2008) 45:1405–13. doi:10.1016/j.
molimm.2007.08.018
37. Hiwatashi K, Tamiya T, Hasegawa E, Fukaya T,
Hashimoto M, Kakoi K, et al. Suppression of
SOCS3 in macrophages prevents cancer metastasis
by modifying macrophage phase and MCP2/CCL8
induction. Cancer Lett (2011) 308:172–80. doi:10.
1016/j.canlet.2011.04.024
38. Matsumura Y, Kobayashi T, Ichiyama K, Yoshida
R, Hashimoto M, Takimoto T, et al. Selective
expansion of foxp3-positive regulatory T cells and
immunosuppression by suppressors of cytokine
signaling 3-deficient dendritic cells. J Immunol

July 2014 | Volume 5 | Article 357 | 242

Wilson

(2007) 179:2170–9. doi:10.4049/jimmunol.179.4.
2170
39. Qin H, Yeh WI, De Sarno P, Holdbrooks AT, Liu
Y, Muldowney MT, et al. Signal transducer and
activator of transcription-3/suppressor of cytokine
signaling-3 (STAT3/SOCS3) axis in myeloid cells
regulates neuroinflammation. Proc Natl Acad Sci
U S A (2012) 109:5004–9. doi:10.1073/pnas.
1117218109
40. Wong PK, Egan PJ, Croker BA, O’Donnell K,
Sims NA, Drake S, et al. SOCS-3 negatively regulates innate and adaptive immune mechanisms
in acute IL-1-dependent inflammatory arthritis.
J Clin Invest (2006) 116:1571–81. doi:10.1172/
JCI25660
41. Ortiz-Munoz G, Martin-Ventura JL, HernandezVargas P, Mallavia B, Lopez-Parra V, Lopes-Franco
O, et al. Suppressors of cytokine signalling modulate JAK/STAT-mediated cell responses during atherosclerosis. Arterioscler Thromb Vasc Biol (2009)
29:525–31. doi:10.1161/ATVBAHA.108.173781
42. Tang J, Kozaki K, Farr AG, Martin PJ, Lindahl
P, Betsholtz C, et al. The absence of plateletderived growth factor-B in circulating cells promotes immune and inflammatory responses in
atherosclerosis-prone ApoE-/- mice. Am J Pathol
(2005) 167:901–12. doi:10.1016/S0002-9440(10)
62061-5
43. Shaikh S, Brittenden J, Lahiri R, Brown PA, Thies
F, Wilson HM. Macrophage subtypes in symptomatic carotid artery and femoral artery plaques.

www.frontiersin.org

SOCS proteins and macrophage polarization

Eur J Vasc Endovasc Surg (2012) 44:491–7. doi:10.
1016/j.ejvs.2012.08.005
44. Grothusen C, Schuett H, Hillmer A, Lumpe S,
Grote K, Ballmaier M. Role of suppressor of
cytokine signaling-1 in murine atherosclerosis.
PLoS One (2012) 7:e51608. doi:10.1371/journal.
pone.0051608
45. Yamamoto TYH, Koga M. Lacking the SOCS3 Gene
in macrophages prevents the development of atherosclerosis in ApoE deficient mice. Circ J (2007)
71:170.
46. Li Y, de Haar C, Peppelenbosch MP, van der Woude
CJ. SOCS3 in immune regulation of inflammatory
bowel disease and inflammatory bowel diseaserelated cancer. Cytokine Growth Factor Rev (2012)
23:127–38. doi:10.1016/j.cytogfr.2012.04.005
47. Hontecillas R, Horne WT, Climent M, Guri AJ,
Evans C, Zhang Y. Immunoregulatory mechanisms
of macrophage PPAR-γ in mice with experimental inflammatory bowel disease. Mucosal Immunol
(2011) 4:304–13. doi:10.1038/mi.2010.75
48. Heys SD, Stewart KN, McKenzie EJ, Miller
ID, Wong SY, Sellar G, et al. Characterisation
of tumour-infiltrating macrophages: impact on
response and survival in patients receiving primary
chemotherapy for breast cancer. Breast Cancer
Res Treat (2009) 135:539–48. doi:10.1007/s10549012-2190-6
49. Hashimoto M, Ayada T, Kinjyo I, Hiwatashi K,
Yoshida H, Okada Y, et al. Silencing of SOCS1
in macrophages suppresses tumor development

by enhancing antitumor inflammation. Cancer Sci
(2009) 100:730–6. doi:10.1111/j.1349-7006.2009.
01098.x
50. Scheller EL, Hankenson KD, Reuben JS, Krebsbach PH. Zoledronic acid inhibits macrophage
SOCS3 expression and enhances cytokine production. J Cell Biochem (2011) 112:3364–72. doi:10.
1002/jcb.23267
Conflict of Interest Statement: The author declares
that the research was conducted in the absence of any
commercial or financial relationships that could be
construed as a potential conflict of interest.
Received: 04 June 2014; paper pending published: 26 June
2014; accepted: 12 July 2014; published online: 28 July
2014.
Citation: Wilson HM (2014) SOCS proteins in
macrophage polarization and function. Front. Immunol.
5:357. doi: 10.3389/fimmu.2014.00357
This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology.
Copyright © 2014 Wilson. This is an open-access article
distributed under the terms of the Creative Commons
Attribution License (CC BY). The use, distribution or
reproduction in other forums is permitted, provided the
original author(s) or licensor are credited and that the
original publication in this journal is cited, in accordance
with accepted academic practice. No use, distribution or
reproduction is permitted which does not comply with
these terms.

July 2014 | Volume 5 | Article 357 | 243

PERSPECTIVE ARTICLE
published: 21 November 2014
doi: 10.3389/fimmu.2014.00554

Myeloid colony-stimulating factors as regulators of
macrophage polarization
Thomas A. Hamilton*, Chenyang Zhao, Paul G. Pavicic Jr. and Shyamasree Datta
Department of Immunology, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Robert Adam Harris, Karolinska
Institutet, Sweden
Cordula M. Stover, University of
Leicester, UK
*Correspondence:
Thomas A. Hamilton, Department of
Immunology, Lerner Research
Institute, Cleveland Clinic Foundation,
9500 Euclid Ave, Cleveland, OH
44195, USA
e-mail: hamiltt@ccf.org

The scope of functional heterogeneity in macrophages has been defined by two polarized
end states known as M1 and M2, which exhibit the proinflammatory activities necessary
for host defense and the tissue repair activities required for restoration of homeostasis,
respectively. Macrophage populations in different tissue locations exist in distinct phenotypic states across this M1/M2 spectrum and the development and abundance of individual
subsets result from the local and systemic action of myeloid colony-stimulating factors
(CSFs) including M-CSF and GM-CSF. These factors have relatively non-overlapping roles in
the differentiation and maintenance of specific macrophage subsets. Furthermore, there is
now evidence that CSFs may also regulate macrophage phenotype during challenge. Cell
culture studies from multiple laboratories demonstrate that macrophages developed in
the presence of GM-CSF exhibit amplified response to M1 polarizing stimuli while M-CSF
potentiates responses to M2 stimuli. As a consequence, these factors can be important
determinants of the magnitude and duration of both acute and chronic inflammatory pathology and may, therefore, be potential targets for therapeutic manipulation in specific human
disease settings.
Keywords: macrophage activation, macrophage colony-stimulating factor, granulocyte macrophage colonystimulating factor, inflammation, cytokines

THE MACROPHAGE POLARIZATION PARADIGM: CLASSICAL
(M1) AND ALTERNATIVE (M2) ACTIVATION PHENOTYPES
The mononuclear phagocyte system is known to exhibit substantial functional heterogeneity (1–3). Though the scope of such
heterogeneity is dramatically large, the prevailing concept of heterogeneity is organized around two polarized endpoints known as
classical and alternative activation, also often termed M1 and M2,
respectively (4–6). These polarized states represent the capacities
to initiate inflammatory response, carry out anti-microbial function, and promote Th1/Th17 adaptive immune responses for M1,
or, for M2, to phagocytize debris, promote wound healing, antagonize destructive inflammation, and suppress adaptive immunity.
These functional activities are mediated by molecular features that
include, for M1, the production of proinflammatory cytokines,
co-stimulatory molecules including CD80/86, effector enzymes
such as iNOS, and NADPH oxidase or, for M2, anti-inflammatory
cytokines, immunosuppressive arginase, and scavenger receptors
(CD163) (5, 6).
Importantly, there are multiple subsets of macrophages with
relatively non-overlapping functional responsibilities and this
additional heterogeneity must be superimposed upon the M1/M2
paradigm (2, 7–9). Resident macrophage populations exhibit
many common features but are not identical, often deriving
from distinct origins and showing a broad spectrum of functional potential (10). It is noteworthy that most resident tissue
macrophages are capable of self-replenishment and generally do
not derive from circulating monocytes (9, 11). There are two major
categories of monocytes found in the circulation. In the mouse,
these are characterized as Ly6C-hi/CCR2hi and Ly6Clo /CX3CR1hi .

Frontiers in Immunology | Molecular Innate Immunity

The former is likely the source of most infiltrating inflammatory
macrophages or dendritic cells, while the latter is a patrolling and
longer lived cell population that provides maintenance of the vascular endothelium (7). The understanding of macrophage subsets
and their developmental origin has advanced dramatically through
the application of fate mapping and other transgenic strategies
(2, 11–13). Moreover, the molecular features responsible for distinct functional activities (anti-microbial, inflammatory, reparative, etc.) have been defined through detailed analysis of isolated
cell populations and cell type-specific transgenic manipulation (5,
6, 14). These macrophage subpopulations exhibit many molecular
similarities that reflect their common myeloid origin but each has
very distinct responsibilities and their inter-conversion appears to
be limited under steady-state conditions (9). Moreover, each of
these subset populations can be induced to exhibit M1 or M2 like
functional polarization.
The concepts of classical and alternative activation or even M1
and M2 markedly oversimplify the spectrum of macrophage phenotypes that exist within vertebrate organisms (6, 15). The number
of individual phenotypes will, in part, depend upon the number of gene products that are measured and the degree to which
gene expression events are independent (16). Furthermore, the
spectrum of environmental cues encountered by macrophages is
highly complex both in number and exposure sequence. Hence,
the combinatorial spectrum of possible phenotypes is extremely
large. As a simplifying principle, the number of specific molecular endpoints used to define phenotype can be limited to those
activities and molecules requisite to the functions of interest (e.g.,
anti-microbial, reparative, etc.). This will enable comparison of

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macrophage populations within different physiologic and pathologic circumstances and evaluation of how the specific molecular characteristics may vary with cell subset and stimulus
environment.
Finally, it is also important to recognize that inflammation is a
dynamic process that is well recognized to proceed in stages with an
early proinflammatory function followed by a transition to healing and restoration of tissue homeostasis (1, 17, 18). While this
process includes the temporally distinct recruitment of multiple
cell populations that can provide different aspects of the evolving
functional features, there is also reason to believe that infiltrating inflammatory monocytes may cycle through different states of
functional activity that are, at a minimum, reminiscent of the M1
and M2 categories (19, 20).

REGULATION OF POLARIZATION
Classically and alternatively activated (M1 or M2) macrophage
phenotypes are defined by the specific molecular characteristics
induced in response to prototypic pro- and anti-inflammatory
stimuli [e.g., IFNγ, Toll-like receptor (TLR) agonists, IL-4/IL13, IL-10, etc.] (5, 15, 18). Such molecular correlates are now
used with increasing frequency to identify populations of M1
or M2 like macrophages in different physiologic settings. Thus,
M1 macrophages express high levels of IL-12 or IL-23, TNFα,
IL-1α/β, chemokines eliciting neutrophil, inflammatory monocyte, and proinflammatory T lymphocyte infiltration (CXCL1-3,
8, CCL2, CXCL9, and CXCL10), CD80/86, CD64, MARCO, iNOS,
and reactive oxygen species (hydroxyl radicals and H2 O2 ) induced
by M1 polarizing stimuli. Correlates of the M2 phenotype include
IL-10, TGFβ, arginase, YM1, FIZZ1 (in the mouse), the mannose receptor (CD206), and scavenger receptors, such as CD163
induced following exposure to IL-4, IL-13, TGFβ, IL-10, and other
agents.
In this context, M1 and M2 phenotypes are often presented as
relating to the actions of Th1 and Th2 cell subsets, respectively,
because of the Th1 and Th2 lymphokine-based modulation of
their defining features (15, 21). It is evident, however, that polarized macrophage function existed prior to the development of
adaptive immunity in evolution (22, 23). Moreover, mice with
T- and B-cell deficiency still possess the potential for polarized
function demonstrating that the M1 and M2 concept can exist
in the absence of adaptive immunity and its products (4). Nevertheless, the normal vertebrate immune system operates with
a full spectrum of immune cell populations and their products
clearly influence the macrophage polarization process. Indeed, it
is likely that macrophages will encounter both M1 and M2 polarizing stimuli simultaneously within the inflamed tissue microenvironment. The complexity of response to this may reasonably
explain the large spectrum of macrophage phenotypes encountered in cell populations in vivo and the variability in markers of
polarization.
Macrophage populations may also exhibit a predisposition for
polarization toward the M1 or M2 phenotype. For example, different strains of mice and rats have been shown to have skewed
patterns of activation potential that ultimately correlate with their
innate and adaptive immune functionality (4, 24, 25). Furthermore, it is well recognized that macrophages in different anatomic

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CSFs and macrophage polarization

or physiologic settings exhibit dramatically different capacities for
polarization. Importantly, there are agents that, by themselves, do
not induce expression of common polarization markers but which
can alter response to stimulation with classical M1 or M2 stimuli.
Because most, if not all macrophage populations, can respond to
either M1 or M2 stimuli, the tone of a response may be set by
differential regulation of sensitivity to polarizing stimuli. Hence,
agents that promote priming of macrophages for enhanced or
diminished response to classical or alternative activation are likely
to be important determinants of the character and temporal patterns of macrophage functional change in the course of response
to injury and infection.
The myeloid colony-stimulating factors (CSFs) M-CSF and
GM-CSF are known to modulate macrophage phenotype (26, 27).
While both agents were first identified as inducers of myeloid cell
differentiation and proliferation in cultured bone marrow progenitors, many studies illustrate their importance in the magnitude,
duration, and character of many forms of inflammatory response
(26–30). Though GM-CSF is associated with classical or M1 activation while M-CSF is linked with alternative or M2 activation,
neither factor is a potent stimulus of definitive polarization markers, when compared with prototypic polarizing stimuli (e.g., IFNγ,
TLRs, IL-4, IL-10, etc.) (31–33). Instead, GM-CSF and M-CSF
appear to induce a state in which macrophages are primed for M1
and M2 endpoints, respectively.

MOLECULAR AND CELLULAR PHENOTYPES PRODUCED BY
M-CSF AND GM-CSF IN VITRO
While receptor signaling mechanisms mediating responses to MCSF and GM-CSF are appreciated in basic detail (5, 6, 26),
understanding of how these agents modulate functional polarization remains obscure. The potential for CSFs to regulate the
responses of mature myeloid cell populations is well recognized.
For example, the capacity of GM-CSF, but not M-CSF, to generate DCs in culture clearly demonstrated distinct functional
roles (34). Verreck et al. initially demonstrated that GM-CSF or
M-CSF treatment of myeloid cells in culture was able to selectively alter the magnitude of M1 or M2 polarized phenotypes
following appropriate stimulation (31, 32). While GM-CSF cultured macrophages stimulated with LPS ± IFNγ produced large
amounts of IL-23 or IL-12 and little IL-10, M-CSF cultured
macrophages were unable to generate either IL-12 or IL-23 but
did produce significant amounts of IL-10 under the same conditions. GM-CSF-treated cells produced appreciably higher levels
of other proinflammatory cytokines including TNF, IL-18, IL-1β,
and IL-6 in comparison to those grown in M-CSF. Furthermore,
M-CSF-treated cells were more phagocytic but less competent in
antigen presentation when compared with GM-CSF treated cells.
For the most part, the growth factor-treated cells did not exhibit
the full M1 or M2 phenotypes in otherwise unstimulated state but
rather showed polarized sensitivity for corresponding response
to IFNγ/TLR signaling. These findings led the authors to conclude that GM-CSF and M-CSF were promoting the development
of monocyte-macrophages predisposed to exhibit differential M1
and M2 phenotypes, respectively. These findings were confirmed
and extended by Fleetwood et al. using bone marrow derived
macrophages from mice cultured in either GM-CSF or M-CSF

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Hamilton et al.

(33). Interestingly, though GM-CSF is frequently used to generate
DCs from bone marrow progenitors, the cells arising from such
cultures more closely resemble macrophages than dendritic cells
based upon whole genome profiling (35). Though there are many
similarities between human being and mouse macrophages in different polarized states, there also appear to be many differences
(14, 35–37). Despite these findings, there remain important questions about the strict relationship between these two factors and
the molecular/functional phenotype definitions for M1 and M2.
Thus, studies in both mouse and human cells show that the effects
of GM-CSF and M-CSF on gene expression do not map exactly
with M1 and M2 marker expression, even following stimulation
(35, 38, 39).
In the context of the hypothesis that macrophage predisposition
is an important determinant of polarized phenotype expression,
there are certainly other natural ligands that have the capacity to
alter macrophage sensitivity to specific polarization stimuli (40–
45). Because inflammatory responses in vivo will always occur in a
complex stimulus environment, these additional agents are likely
to co-operate with or antagonize the actions of the CSFs. PPARγ,
in particular, has been reported to be required for development
of alternatively activated macrophages in the context of insulin
resistance and metabolic inflammatory disease (46). The tyrosine
kinase receptor CD136 (RON, MST1R) can also modulate sensitivity for M2-like activators, in part by altering the sensitivity to TLR
stimulation (24, 47). In contrast to these agents, NOTCH and its
ligand RBP-J, are reported to promote M1-like responses via alterations in intracellular signaling factors including IRF8 and SOCS3
(40). Indeed, the sensitivity of myeloid cell populations to polarizing stimuli appears to be controlled in part by alterations in the
abundance or activity of the signaling pathway components that
mediate responses to pro- and anti-inflammatory stimuli (41–44,
48–50). Thus, signaling adaptors, protein kinases, protein phosphatases, and transcription factors including members of the IRF,
SOCS, Tec, and KLF families have all been implicated in controlling either M1 or M2 polarization. The mechanisms through which
stimulus sensitivity is altered by ligand/receptor pairs, such as the
CSFs, as well as others mentioned above remains to be fully elucidated but intracellular signaling factors are likely to be important
targets.

ROLE OF M-CSF AND GM-CSF IN MACROPHAGE
FUNCTIONAL POLARIZATION IN VIVO
M-CSF and GM-CSF have distinct effects on the development and
expansion of myeloid cell populations in different anatomic settings (26, 27). Based upon studies of mice with targeted deletion
of ligand and/or receptor genes, M-CSF is known to be required
for the production and maintenance of many (though not all)
tissue macrophage populations (51). In this regard, distinctions
between receptor and ligand deficient mice revealed the existence
of a second ligand (IL-34), which is now known to be necessary
for the maintenance of a subset of tissue macrophage populations
(microglia and Langerhans cells) (52). M-CSF and IL-34 function in homeostatic maintenance of tissue-resident macrophage
populations through promoting viability and proliferation and
both drive predisposition to M2 character (53). While M-CSF

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CSFs and macrophage polarization

is found in the serum of healthy individuals and can be produced constitutively by epithelia, fibroblasts, endothelial cells, and
by macrophages themselves, its expression can be elevated by
inflammatory conditions in many cells including macrophages
as well as T and B lymphocytes (27). In contrast to M-CSF or
IL-34, GM-CSF deficiency has little impact on steady-state tissue macrophage populations with the exception of those found in
the lung (26, 27). While GM-CSF is believed to be important for
the development of infiltrating inflammatory DCs, recent findings show that M-CSF but not GM-CSF sensitivity is requisite for
these cell populations (13). Importantly, GM-CSF, unlike M-CSF,
is not detectable in most tissues at rest but expression is frequently
induced during inflammatory or immune stimulation in many
tissues and cell types (26, 27, 54).
While studies using animals with global or cell-type restricted
deficiencies in M-CSF or GM-CSF ligand/receptor function do
provide insight into their relative contributions to macrophage
phenotypes during inflammatory responses in vivo, the interpretation of such experiments should be viewed with caution
due to the impact of such deficiencies on development and/or
abundance of specific myeloid subsets. These studies are effectively complemented by transiently manipulating M- or GM-CSF
levels using specific ligand delivery or ligand/receptor antagonism. Results from multiple studies indicate that both M-CSF
and GM-CSF can modulate the magnitude and character of
inflammatory response in multiple tissue specific disease models (2, 26, 54–56). These include autoimmune encephalomyelitis
(MS), atherosclerosis, arthritis, nephritis, lung inflammation, and
cancer. In most cases, however, the mechanisms through which
these endpoints are achieved have not been elucidated. Certainly,
both factors have the capacity to promote survival and/or expansion of macrophage populations both systemically and in specific tissue locations and the decreased number of macrophages
observed with CSF antagonism would likely result in reduced
intensity and/or duration of disease (26). Multiple approaches
including delivery of M-CSF, antibody-mediated depletion of
the ligand, antibody-induced antagonism of receptor, or the
use of receptor tyrosine kinase inhibitors have provided evidence supporting both positive and inhibitory roles for M-CSF
in inflammatory diseases (26, 55). Interestingly, several recent
reports demonstrate that antagonistic targeting of M-CSF can
have appreciable benefit in tumor therapy as a consequence of
altering the tumor-associated macrophage phenotype from M2 to
M1 (49, 57–59).
There are several important considerations that can provide
some speculative insight into the role for these CSFs in regulating the nature of macrophage polarization in vivo. First are
the patterns of CSF expression within tissues both at rest and
during inflammatory responses. While the expression of both
factors can be amplified during response to various forms of
tissue injury, it is apparent that M-CSF is produced constitutively in many tissues and is critical for the maintenance of
resident tissue populations throughout the body. GM-CSF, in
contrast, is rarely detectable except at times of injury and does
not appear to be a critical determinant of macrophage numbers with a few exceptions (i.e., the lung). Second, resident

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macrophage populations are generally found to exhibit an M2like phenotype under resting conditions, consistent with the need
to minimize tissue damaging inflammatory reaction (10). This
is certainly consistent with the ability of M-CSF to predispose
toward an M2 phenotype in cell culture experiments involving both human monocytes and mouse bone marrow-derived
macrophages. Third, GM-CSF has been shown to be a critical
determinant of inflammatory injury in several model systems (60–
62). Particularly, T-cell derived GM-CSF was recently shown to
be critical for disease phenotype in experimental autoimmune
encephalomyelitis, generated by IL-23 action on Th17 cells (60,
61). In this instance, GM-CSF selectively targeted infiltrating
macrophages within the CNS. Finally, several studies examining responses to M-CSF and GM-CSF in vitro show that the
M1/M2 phenotype can be reversibly modulated by GM-CSF/MCSF exposure in cell culture and GM-CSF predisposition may be
dominant (33, 39, 63) (Hamilton, unpublished). This is also supported by increased bioavailability (half-life) of GM-CSF, which
might also contribute to its dominance relative to M-CSF. We
would suggest then that M-CSF provides the default condition
and will promote an M2 (healing) phenotype both at rest and in
the absence of other forms of stimulation. Induced expression of
GM-CSF (e.g., during adaptive T-cell driven immune responses)
will provide the mechanism for retaining or re-expressing an M1
phenotype when conditions require (e.g., continued infection or
injury). It is clear, however, that the specific effects of M-CSF
and GM-CSF on macrophage polarization in cell culture models are unlikely to fully predict their effects in vivo. This reflects
complexities associated with the variable nature of inflammatory
injury, stimulus exposure, and distinct features of specific tissue
microenvironments.

UNANSWERED QUESTIONS
Macrophage heterogeneity or phenotype polarization is an area
of high current interest and the impact of myeloid CSFs on this
process in vivo is evident but poorly understood. Hence, we pose
the following outstanding questions regarding the roles that MCSF and GM-CSF may play, particularly through modulating
sensitivity to M1 and M2 promoting stimuli with the expectation
that answers will help to clarify the process and provide insights
to therapeutic application. (1) What mechanisms are involved in
skewing responses to polarizing stimuli? Can we identify specific
CSF-induced patterns of gene expression that are requisite to generating macrophages predisposed for more potent responses to
cytokines and pattern recognition receptors? Can we correlate outcomes obtained in vitro with those obtained in vivo? (2) What are
the sources and timing of M-CSF and GM-CSF expression within
specific tissues during different forms of inflammatory response?
Which myeloid cell populations are the targets of the CSFs and how
is this co-ordinated with the need to enhance or diminish specific
aspects of function over the full course of inflammatory response?
(3) Finally, we must begin to consider not only the mechanisms
through which M-CSF and GM-CSF operate but also how these
stimuli are integrated with the host of other agents encountered
within inflammatory settings that also have marked influence on
M1/M2 skewing?

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CSFs and macrophage polarization

REFERENCES
1. Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol (2011) 11(11):723–37. doi:10.1038/nri3073
2. Wynn TA, Chawla A, Pollard JW. Macrophage biology in development, homeostasis and disease. Nature (2013) 496(7446):445–55. doi:10.1038/nature12034
3. Gordon S, Taylor PR. Monocyte and macrophage heterogeneity. Nat Rev
Immunol (2005) 5(12):953–64. doi:10.1038/nri1733
4. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
5. Martinez FO, Gordon S. The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000Prime Rep (2014) 6:13. doi:10.12703/P6-13
6. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122(3):787–95. doi:10.1172/JCI59643
7. Geissmann F, Jung S, Littman DR. Blood monocytes consist of two principal subsets with distinct migratory properties. Immunity (2003) 19(1):71–82.
doi:10.1016/S1074-7613(03)00174-2
8. Auffray C, Sieweke MH, Geissmann F. Blood monocytes: development, heterogeneity, and relationship with dendritic cells. Annu Rev Immunol (2009)
27:669–92. doi:10.1146/annurev.immunol.021908.132557
9. Geissmann F, Manz MG, Jung S, Sieweke MH, Merad M, Ley K. Development of
monocytes, macrophages, and dendritic cells. Science (2010) 327(5966):656–61.
doi:10.1126/science.1178331
10. Davies LC, Jenkins SJ, Allen JE, Taylor PR. Tissue-resident macrophages. Nat
Immunol (2013) 14(10):986–95. doi:10.1038/ni.2705
11. Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, et al. Tissueresident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity (2013) 38(4):792–804.
doi:10.1016/j.immuni.2013.04.004
12. Ginhoux F, Greter M, Leboeuf M, Nandi S, See P, Gokhan S, et al. Fate mapping
analysis reveals that adult microglia derive from primitive macrophages. Science
(2010) 330(6005):841–5. doi:10.1126/science.1194637
13. Greter M, Helft J, Chow A, Hashimoto D, Mortha A, Agudo-Cantero J, et al.
GM-CSF controls nonlymphoid tissue dendritic cell homeostasis but is dispensable for the differentiation of inflammatory dendritic cells. Immunity (2012)
36(6):1031–46. doi:10.1016/j.immuni.2012.03.027
14. Lawrence T, Natoli G. Transcriptional regulation of macrophage polarization: enabling diversity with identity. Nat Rev Immunol (2011) 11(11):750–61.
doi:10.1038/nri3088
15. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
16. Adams DO, Hamilton TA. Molecular basis of macrophage activation: diversity
and its origins. In: Lewis CE, O’D McGee J, editors. The Natural Immune System,
Volume II: The Macrophage. New York, NY: Oxford University Press (1992). p.
75–114.
17. Lichtnekert J, Kawakami T, Parks WC, Duffield JS. Changes in macrophage
phenotype as the immune response evolves. Curr Opin Pharmacol (2013)
13(4):555–64. doi:10.1016/j.coph.2013.05.013
18. Mantovani A, Biswas SK, Galdiero MR, Sica A, Locati M. Macrophage plasticity
and polarization in tissue repair and remodelling. J Pathol (2013) 229(2):176–85.
doi:10.1002/path.4133
19. Holt MP, Cheng L, Ju C. Identification and characterization of infiltrating
macrophages in acetaminophen-induced liver injury. J Leukoc Biol (2008)
84(6):1410–21. doi:10.1189/jlb.0308173
20. Arnold L, Henry A, Poron F, Baba-Amer Y, van Rooijen N, Plonquet A, et al.
Inflammatory monocytes recruited after skeletal muscle injury switch into
antiinflammatory macrophages to support myogenesis. J Exp Med (2007)
204(5):1057–69. doi:10.1084/jem.20070075
21. Martinez FO, Helming L, Gordon S. Alternative activation of macrophages:
an immunologic functional perspective. Annu Rev Immunol (2009) 27:451–83.
doi:10.1146/annurev.immunol.021908.132532
22. Dzik JM. The ancestry and cumulative evolution of immune reactions. Acta
Biochim Pol (2010) 57(4):443–66.
23. Stefater JA III, Ren S, Lang RA, Duffield JS. Metchnikoff ’s policemen:
macrophages in development, homeostasis and regeneration. Trends Mol Med
(2011) 17(12):743–52. doi:10.1016/j.molmed.2011.07.009

November 2014 | Volume 5 | Article 554 | 247

Hamilton et al.

24. Chaudhuri A, Wilson NS, Yang B, Paler Martinez A, Liu J, Zhu C, et al.
Host genetic background impacts modulation of the TLR4 pathway by RON
in tissue-associated macrophages. Immunol Cell Biol (2013) 91(7):451–60.
doi:10.1038/icb.2013.27
25. Andersson A, Kokkola R, Wefer J, Erlandsson-Harris H, Harris RA. Differential macrophage expression of IL-12 and IL-23 upon innate immune activation defines rat autoimmune susceptibility. J Leukoc Biol (2004) 76(6):1118–24.
doi:10.1189/jlb.0704385
26. Hamilton JA, Achuthan A. Colony stimulating factors and myeloid cell biology
in health and disease. Trends Immunol (2013) 34(2):81–9. doi:10.1016/j.it.2012.
08.006
27. Hamilton JA. Colony-stimulating factors in inflammation and autoimmunity.
Nat Rev Immunol (2008) 8(7):533–44. doi:10.1038/nri2356
28. Vogel DY, Glim JE, Stavenuiter AW, Breur M, Heijnen P, Amor S, et al. Human
macrophage polarization in vitro: maturation and activation methods compared. Immunobiology (2014) 219(9):695–703. doi:10.1016/j.imbio.2014.05.002
29. Page AV, Liles WC. Colony-stimulating factors in the prevention and management of infectious diseases. Infect Dis Clin North Am (2011) 25(4):803–17.
doi:10.1016/j.idc.2011.07.007
30. Akagawa KS. Functional heterogeneity of colony-stimulating factor-induced
human monocyte-derived macrophages. Int J Hematol (2002) 76(1):27–34.
doi:10.1007/BF02982715
31. Verreck FA, de Boer T, Langenberg DM, Hoeve MA, Kramer M, Vaisberg E, et al.
Human IL-23-producing type 1 macrophages promote but IL-10-producing
type 2 macrophages subvert immunity to (myco)bacteria. Proc Natl Acad Sci U
S A (2004) 101(13):4560–5. doi:10.1073/pnas.0400983101
32. Verreck FA, de Boer T, Langenberg DM, van der Zanden L, Ottenhoff TH.
Phenotypic and functional profiling of human proinflammatory type-1 and
anti-inflammatory type-2 macrophages in response to microbial antigens
and IFN-gamma- and CD40L-mediated costimulation. J Leukoc Biol (2006)
79(2):285–93. doi:10.1189/jlb.0105015
33. Fleetwood AJ, Lawrence T, Hamilton JA, Cook AD. Granulocyte-macrophage
colony-stimulating factor (CSF) and macrophage CSF-dependent macrophage
phenotypes display differences in cytokine profiles and transcription factor
activities: implications for CSF blockade in inflammation. J Immunol (2007)
178(8):5245–52. doi:10.4049/jimmunol.178.8.5245
34. Inaba K, Inaba M, Romani N, Aya H, Deguchi M, Ikehara S, et al. Generation
of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colony-stimulating factor. J Exp Med
(1992) 176(6):1693–702. doi:10.1084/jem.176.6.1693
35. Lacey DC, Achuthan A, Fleetwood AJ, Dinh H, Roiniotis J, Scholz GM, et al.
Defining GM-CSF- and macrophage-CSF-dependent macrophage responses by
in vitro models. J Immunol (2012) 188(11):5752–65. doi:10.4049/jimmunol.
1103426
36. Beyer M, Mallmann MR, Xue J, Staratschek-Jox A, Vorholt D, Krebs W,
et al. High-resolution transcriptome of human macrophages. PLoS One (2012)
7(9):e45466. doi:10.1371/journal.pone.0045466
37. Xue J, Schmidt SV, Sander J, Draffehn A, Krebs W, Quester I, et al. Transcriptomebased network analysis reveals a spectrum model of human macrophage activation. Immunity (2014) 40(2):274–88. doi:10.1016/j.immuni.2014.01.006
38. Ishii M, Wen H, Corsa CA, Liu T, Coelho AL, Allen RM, et al. Epigenetic
regulation of the alternatively activated macrophage phenotype. Blood (2009)
114(15):3244–54. doi:10.1182/blood-2009-04-217620
39. Mia S, Warnecke A, Zhang XM, Malmstrom V, Harris RA. An optimized protocol for human M2 macrophages using M-CSF and IL-4/IL-10/TGF-beta
yields a dominant immunosuppressive phenotype. Scand J Immunol (2014)
79(5):305–14. doi:10.1111/sji.12162
40. Xu H, Zhu J, Smith S, Foldi J, Zhao B, Chung AY, et al. Notch-RBP-J signaling
regulates the transcription factor IRF8 to promote inflammatory macrophage
polarization. Nat Immunol (2012) 13(7):642–50. doi:10.1038/ni.2304
41. Lee B, Qiao L, Lu M, Yoo HS, Cheung W, Mak R, et al. C/EBPalpha regulates
macrophage activation and systemic metabolism. Am J Physiol Endocrinol Metab
(2014) 306(10):E1144–54. doi:10.1152/ajpendo.00002.2014
42. Qin H, Holdbrooks AT, Liu Y, Reynolds SL, Yanagisawa LL, Benveniste EN.
SOCS3 deficiency promotes M1 macrophage polarization and inflammation.
J Immunol (2012) 189(7):3439–48. doi:10.4049/jimmunol.1201168
43. Festuccia WT, Pouliot P, Bakan I, Sabatini DM, Laplante M. Myeloid-specific
rictor deletion induces M1 macrophage polarization and potentiates

Frontiers in Immunology | Molecular Innate Immunity

CSFs and macrophage polarization

in vivo pro-inflammatory response to lipopolysaccharide. PLoS One (2014)
9(4):e95432. doi:10.1371/journal.pone.0095432
44. Tao B, Jin W, Xu J, Liang Z, Yao J, Zhang Y, et al. Myeloid-specific disruption
of tyrosine phosphatase Shp2 promotes alternative activation of macrophages
and predisposes mice to pulmonary fibrosis. J Immunol (2014) 193(6):2801–11.
doi:10.4049/jimmunol.1303463
45. Olefsky JM, Glass CK. Macrophages, inflammation, and insulin resistance. Annu
Rev Physiol (2010) 72:219–46. doi:10.1146/annurev-physiol-021909-135846
46. Odegaard JI, Ricardo-Gonzalez RR, Goforth MH, Morel CR, Subramanian
V, Mukundan L, et al. Macrophage-specific PPARgamma controls alternative
activation and improves insulin resistance. Nature (2007) 447(7148):1116–20.
doi:10.1038/nature05894
47. Sharda DR, Yu S, Ray M, Squadrito ML, De Palma M, Wynn TA, et al. Regulation
of macrophage arginase expression and tumor growth by the Ron receptor tyrosine kinase. J Immunol (2011) 187(5):2181–92. doi:10.4049/jimmunol.1003460
48. Liao X, Sharma N, Kapadia F, Zhou G, Lu Y, Hong H, et al. Kruppel-like factor 4 regulates macrophage polarization. J Clin Invest (2011) 121(7):2736–49.
doi:10.1172/JCI45444
49. Ni Gabhann J, Hams E, Smith S, Wynne C, Byrne JC, Brennan K, et al. Btk
regulates macrophage polarization in response to lipopolysaccharide. PLoS One
(2014) 9(1):e85834. doi:10.1371/journal.pone.0085834
50. Satoh T, Takeuchi O, Vandenbon A, Yasuda K, Tanaka Y, Kumagai Y, et al.
The Jmjd3-Irf4 axis regulates M2 macrophage polarization and host responses
against helminth infection. Nat Immunol (2010) 11(10):936–44. doi:10.1038/
ni.1920
51. Dai XM, Zong XH, Sylvestre V, Stanley ER. Incomplete restoration of colonystimulating factor 1 (CSF-1) function in CSF-1-deficient Csf1op/Csf1op mice
by transgenic expression of cell surface CSF-1. Blood (2004) 103(3):1114–23.
doi:10.1182/blood-2003-08-2739
52. Greter M, Lelios I, Pelczar P, Hoeffel G, Price J, Leboeuf M, et al. Stromaderived interleukin-34 controls the development and maintenance of Langerhans cells and the maintenance of microglia. Immunity (2012) 37(6):1050–60.
doi:10.1016/j.immuni.2012.11.001
53. Foucher ED, Blanchard S, Preisser L, Garo E, Ifrah N, Guardiola P, et al. IL34 induces the differentiation of human monocytes into immunosuppressive
macrophages. Antagonistic effects of GM-CSF and IFNgamma. PLoS One (2013)
8(2):e56045. doi:10.1371/journal.pone.0056045
54. Shi Y, Liu CH, Roberts AI, Das J, Xu G, Ren G, et al. Granulocyte-macrophage
colony-stimulating factor (GM-CSF) and T-cell responses: what we do and don’t
know. Cell Res (2006) 16(2):126–33. doi:10.1038/sj.cr.7310017
55. Hume DA, MacDonald KP. Therapeutic applications of macrophage colonystimulating factor-1 (CSF-1) and antagonists of CSF-1 receptor (CSF-1R) signaling. Blood (2012) 119(8):1810–20. doi:10.1182/blood-2011-09-379214
56. Lenzo JC, Turner AL, Cook AD, Vlahos R, Anderson GP, Reynolds EC, et al.
Control of macrophage lineage populations by CSF-1 receptor and GM-CSF
in homeostasis and inflammation. Immunol Cell Biol (2012) 90(4):429–40.
doi:10.1038/icb.2011.58
57. Pyonteck SM, Akkari L, Schuhmacher AJ, Bowman RL, Sevenich L, Quail DF,
et al. CSF-1R inhibition alters macrophage polarization and blocks glioma progression. Nat Med (2013) 19(10):1264–72. doi:10.1038/nm.3337
58. Gabrilovich DI, Ostrand-Rosenberg S, Bronte V. Coordinated regulation of
myeloid cells by tumours. Nat Rev Immunol (2012) 12(4):253–68. doi:10.1038/
nri3175
59. Sluijter M, van der Sluis TC, van der Velden PA, Versluis M, West BL, van der
Burg SH, et al. Inhibition of CSF-1R supports T-cell mediated melanoma therapy. PLoS One (2014) 9(8):e104230. doi:10.1371/journal.pone.0104230
60. Codarri L, Gyulveszi G, Tosevski V, Hesske L, Fontana A, Magnenat L, et al.
RORgammat drives production of the cytokine GM-CSF in helper T cells,
which is essential for the effector phase of autoimmune neuroinflammation.
Nat Immunol (2011) 12(6):560–7. doi:10.1038/ni.2027
61. El-Behi M, Ciric B, Dai H, Yan Y, Cullimore M, Safavi F, et al. The encephalitogenicity of T(H)17 cells is dependent on IL-1- and IL-23-induced production of the cytokine GM-CSF. Nat Immunol (2011) 12(6):568–75. doi:10.1038/
ni.2031
62. Campbell IK, van Nieuwenhuijze A, Segura E, O’Donnell K, Coghill E, Hommel M, et al. Differentiation of inflammatory dendritic cells is mediated by
NF-kappaB1-dependent GM-CSF production in CD4 T cells. J Immunol (2011)
186(9):5468–77. doi:10.4049/jimmunol.1002923

November 2014 | Volume 5 | Article 554 | 248

Hamilton et al.

63. Xu W, Zhao X, Daha MR, van Kooten C. Reversible differentiation of pro- and
anti-inflammatory macrophages. Mol Immunol (2012) 53(3):179–86. doi:10.
1016/j.molimm.2012.07.005

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 29 August 2014; accepted: 18 October 2014; published online: 21 November
2014.

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CSFs and macrophage polarization

Citation: Hamilton TA, Zhao C, Pavicic PG Jr and Datta S (2014) Myeloid colonystimulating factors as regulators of macrophage polarization. Front. Immunol. 5:554.
doi: 10.3389/fimmu.2014.00554
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Hamilton, Zhao, Pavicic and Datta. This is an open-access article
distributed under the terms of the Creative Commons Attribution License (CC BY).
The use, distribution or reproduction in other forums is permitted, provided the original
author(s) or licensor are credited and that the original publication in this journal is cited,
in accordance with accepted academic practice. No use, distribution or reproduction is
permitted which does not comply with these terms.

November 2014 | Volume 5 | Article 554 | 249

PERSPECTIVE ARTICLE
published: 31 October 2014
doi: 10.3389/fimmu.2014.00546

Regulation of macrophage polarization by RON receptor
tyrosine kinase signaling
Amitabha Chaudhuri *
MedGenome Inc., San Francisco, CA, USA

Edited by:
Laurel L. Lenz, University of Colorado
School of Medicine, USA
Reviewed by:
Christine Skerka, Leibniz Institute for
Natural Product Research and
Infection Biology, Germany
Miki Nakao, Kyushu University, Japan
*Correspondence:
Amitabha Chaudhuri , MedGenome
Inc., QB3@953, 953 Indiana Street,
San Francisco, CA 94107, USA
e-mail: amitc@medgenome.com

The M1 and M2 states of macrophage polarization are the two extremes of a physiologic/phenotypic continuum that is dynamically influenced by environmental signals. The
M1/M2 paradigm is an excellent framework to understand and appreciate some of the
diverse functions that macrophages perform. Molecular analysis of mouse and human
macrophages indicated that they gain M1 and M2-related functions after encountering
specific ligands in the tissue environment. In this perspective, I discuss the function of
recepteur d’origine nantais (RON) receptor tyrosine kinase in regulating the M2-like state
of macrophage activation Besides decreasing pro-inflammatory cytokine production in
response to toll-like receptor-4 activation, macrophage-stimulating protein strongly suppresses nitric oxide synthase and at the same time upregulates arginase, which is the rate
limiting enzyme in the ornithine biosynthesis pathway. Interestingly, RON signaling preserved some of the characteristics of the M1 state, while still promoting the hallmarks of
M2 polarization. Therefore, therapeutic modulation of RON activity can shift the activation
state of macrophages between acute and chronic inflammatory states.
Keywords: macrophages, polarization, RON signaling, tumor promotion, immune therapy

INTRODUCTION
Macrophages perform the essential function of preserving tissue
homeostasis following infection or tissue damage in all animals in
the absence of B and T cells. Macrophages originate from bone
marrow-derived monocytic cells through a process of differentiation, directed by the activation of specific transcription factors
(1–3). Circulating monocytes are recruited to tissues, where they
differentiate into functionally distinct subsets of cells with distinct
phenotypic characteristics. In response to the tissue microenvironment, these cells can either produce pro-inflammatory cytokines to
kill the offending foreign pathogen and polarize T-cells to mount
an adaptive immune response, or participate in tissue repair by
increasing their phagocytic activity and producing growth factors
for tissue healing and regeneration. The cellular plasticity is a hall
mark of macrophages and the complex signaling pathways that
contribute to these biochemical and functional differentiation is
beginning to be understood (4, 5).
The two states of macrophage activation, “Classical” (M1) and
“Alternative” (M2) occupy two extremes of a phenotypic continuum in which macrophages respond to secreted factors to evoke
distinct functional responses (6, 7). These functional responses
are regulated by a combination of signaling pathway modulators
and transcription factors. As an example, combination of IFN-γ
with toll-like receptor (TLR) pathway activation produces a complete M1 phenotype in macrophages mediated by the activation of
STAT1 and NF-κB transcription factors. By contrast, IFN-γ alone
causes a partial M1 response mediated by STAT1 transcription factor at sites of infection. Similarly, macrophage M2 phenotype and
response are fine-tuned at tissue-specific sites by the activation of
distinct sets of chemicals, such as IL-10 in the gut, or IL-4 and fatty
acids in the adipose tissues (8, 9).

Frontiers in Immunology | Molecular Innate Immunity

The activation of macrophages (AAM) into M1- or M2-type is
dictated by the cytokine milieu of the tissue microenvironment.
Monocytes are primed to differentiate in response to macrophage
colony stimulating factor (M-CSF) or by granulocyte macrophage
colony stimulating factor (GM-CSF) (2, 3). Further priming is dictated by a balance between IFN-γ and IL-4, the former pushing the
macrophages into an M1 state and the latter into an M2 state. The
primed macrophages receive additional signals in the form of TLR
stimulation to display the full complement of classical and alternative activation functions. Whereas, M1-primed macrophages
produce pro-inflammatory cytokines and inducible nitric oxide
synthase (iNOS) upon TLR activation, M2-primed macrophages
produce arginase, IL-10, and growth factors such as transforming growth factor-β (TGF-β), vascular endothelial growth factor (VEGF), and epidermal growth factor (EGF) among others.
Interestingly, these macrophage phenotypes can be reversed and
brought back to the state of naïve macrophages by growing them in
the absence of any priming factors for a couple of days, indicating
reversibility of the response (10, 11).

INVOKING THE M1 PHENOTYPE OF MACROPHAGES
M1 polarization can be evoked by treating naïve macrophages
with a combination of IFN-γ and LPS. Macrophages express a
variety of microbial pattern recognition receptors (PRRs), such as
TLRs that recognize pathogen – or danger associated molecular
patterns to clear the offending signal (12, 13). A large number of
inflammatory cytokines including TNF-α, IFN-γ IL-12p40, IL-6
are produced rapidly after TLR-4 activation. The delayed TLR-4
response is triggered by the recruitment of TIR-containing adaptor protein (TRIF) and TRIF-related adaptor molecule (TRAM) to
induce IFN-β and trigger the interferon response (14). Together,

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the NF-κB, p38 MAPK, and IFN-β pathways regulate the output of the TLR-4 signaling. A key enzyme of arginine metabolism, iNOS metabolizes arginine to make nitric oxide (NO),
a potent anti-microbial agent. Production of NO is a hallmark
of M1 macrophages and IFN-γ regulates NO production via
transcriptional upregulation of iNOS (4, 15).
Built within these signaling circuits are negative feedback loops
that circumscribe the intensity and duration of the LPS response.
Proteins, such as dual specificity phosphatases (DUSP) and suppressor of cytokine signaling (SOCS) are induced to dephosphorylate upstream activators of the MAPK pathway and inhibit
signaling downstream of the interferon receptor, respectively (6,
16). As part of body’s defense mechanism, M1 polarization is critical to mount an effective innate immune response against the
offending pathogen. However, the body can sustain extensive tissue
damage if the pro-inflammatory responses are allowed to persist.
Therefore, these multiple feedback loops are turned on downstream of TLR-4 and cytokine-signaling pathways that quickly
reduce the output from these pro-inflammatory signaling circuits
(17, 18).

INVOKING THE M2 PHENOTYPE OF MACROPHAGES
Tissue resident macrophages assume an M2 phenotype by default.
This phenotype, also defined as alternative AAM or M2-type of
macrophages can be induced by IL-4 + IL-13 and by other signaling molecules (6, 9). In this state, macrophages metabolize
arginine into ornithine by the expression of arginase-1 that diverts
arginine from the production of NO and citrulline. (15). The M2
macrophages also produce growth factors and extracellular matrix
remodeling enzymes that promote processes related to tissue repair
and healing. Additionally, their phagocytic activity is increased to
help in clearing tissue debris. The M2 activity is sustained by factors produced by injured tissues such as TGF-β and adenosine
(4, 19).

SIGNALING PATHWAYS MEDIATING MACROPHAGE
POLARIZATION
The signaling circuitry leading to changes in gene expression
pattern during macrophage polarization is complex (16). Different subsets of tissue-specific macrophages are dependent on
different signaling pathways to polarize and sustain their polarized state. As an example, c-jun N-terminal kinase pathway
(JNK) is required for the adipose tissue-associated macrophages
to assume M1-phenotype (20). Polarization of macrophages by
the phosphatidyl inositol-3 kinase (PI3K) pathway is mediated
by the activation of AKT1 and AKT2 kinases. Genetic ablation experiments revealed that AKT1 and 2 regulate macrophage
M1 and M2 phenotype in a reciprocal pattern (21, 22). In
the absence of AKT1, macrophages produce pro-inflammatory
cytokines resembling the M1 phenotype, whereas in the absence
of AKT2 the cells express markers of M2 polarization such
as Arg-1, Fizz-1, and IL-10 (21, 23, 24). Interestingly, preliminary data support that this reciprocal regulation of macrophage
M1/M2 phenotypes is mediated by a micro-RNA, mir-155,
and its target transcriptional regulator CAAT-enhanced binding protein-β play an important role (23). JAK/STAT pathway
downstream to IFN-γ is a strong inducer of M1 polarization,

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Macrophage polarization in health and disease

although to reach the full spectrum of the M1-phenotypic state,
dual activation of the TLR-4 and IFN-γ pathways are required
(25, 26).
Signals that promote M2 polarization are diverse and a variety of molecules from cytokines to growth factors can influence
this transition (6). The IL-4/IL-13 combination is a physiological
mediator of the M2 state that impinges on the transcription factor
STAT-6 to induce cell surface expression of M2 markers and metabolic reprograming (9). In the absence of IL-4/IL-13, M-CSF and
IL-10 can push macrophages to assume an M2 phenotype mediated by the transcription factor STAT-3 and SP-1, respectively.
In this state, the macrophages become highly phagocytic, produce
growth factors that promote repair of wound or tissue damage, and
promote Th-2 immune response. Fcγ receptors in combination
with LPS promotes Th-2 response, upregulates antigen presentation, turns off IL-12, and induces IL-10 production by activating
the Syk and PI3K pathways that cross-talk with TLR signaling.
Finally, glucocorticoids promote macrophage adherence, spreading, phagocytosis, induction of complement proteins, and secretion of IL-10 by directly engaging the macrophage transcription
machinery.
Integrating the function of M1 and M2 macrophages in a
physiological setting raises several questions:
1. Are the M1 and M2 macrophage states mutually exclusive in a
tissue environment?
2. Can a cell transition from one state to the other directly, or are
there other intermediate states?
3. Can a macrophage assume characteristics of both M1 and
M2 states? How do they arise? Do they represent a fleeting
intermediate, or can cells in this state be stabilized?
Answers to these questions are not fully known. However, M1and M2-polarized macrophages are found as mixed populations
in the tissue microenvironment. Depending on the inflammatory stimuli, one state may dominate over the other. Both M1
and M2 states are functionally and phenotypically heterogeneous.
The translatability of macrophage M1/M2 polarization in normal homeostasis and in diseases have remained elusive beyond
the fact that M1 macrophages favor bacterial and viral elimination, whereas M2 macrophages give protection against helminthes
and other parasites and participate in tissue repair. Observations
in vitro cannot always be readily applied to in vivo situations and
many of the macrophage responses discussed above are yet to find
validation in vivo (27).

HOW FUNCTIONAL STATES OF MACROPHAGES ARE
MODULATED BY GROWTH FACTOR SIGNALING
So far in our discussion, the macrophage phenotypic and functional states have been described in the context of inflammation
or interaction between pathogens and immune cells. However,
M2-polarized macrophages also perform essential functions in
the resolution of tissue inflammation, remodeling of the tissue
microenvironment during wound healing and repair of tissue
damage. In the next section, the effect of growth factor signaling on macrophage polarization, and its implication in human
cancer is discussed.

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with MSP alone induced activation of MAPK and PI3K pathways,
but failed to induce any of the hall marks of M2 polarization such
as expression of arginase-1, scavenger receptors, or IL-10 (34).
When MSP was combined with LPS stimulation, macrophages
exhibited hallmarks of M2-polarized state (Figure 1B). Further,
global gene expression analysis and measurement of cytokines in
the conditioned media indicated that RON signaling had minimal
effect on TLR-4-mediated early NF-κB activation – the effect being
significantly pronounced at later time points (34).
Taken together, these observations support that RON signaling has dual effect on macrophages. On the one hand, it enhances
macrophage motility and survival without any input from TLR signaling, but on the other hand, it significantly modifies the TLR-4
signaling output when LPS is present along with MSP. Interestingly, the reprograming of the TLR-4 signaling circuits by RON is
sensitive to mice genetic background (34).

IMPACT OF GENETIC BACKGROUND AND DIFFERENTIAL
EFFECT OF RON ON MACROPHAGE POLARIZATION

FIGURE 1 | Recepteur d’origine nantais (RON) promotes macrophage
M2 polarization. (A) Structure of RON and MSP proteins. (B) RON
signaling alters macrophage phenotype and restores the balance between
M1- and M2-polarized states.

Among the growth factor receptors known to modulate
macrophage behavior and function is the receptor tyrosine kinase
“recepteur d’origine nantais” (RON) (28, 29). The ligand for RON,
macrophage-stimulating protein (MSP) regulates macrophage
motility and its phagocytic activity (28). MSP is produced by
the liver and circulates in an inactive form in the serum. MSP
activation occurs as a result of proteolytic processing of ProMSP resulting in active MSP. Several trypsin like proteases such
as matriptase, hepsin, and hepatocyte growth factor-A (HGF-A)
cleave inactive pro-MSP into an active form (30) (Figure 1A).
These proteases are known to be activated at sites of inflammation
and can be a source of active MSP that can turn on RON signaling
on macrophages and epithelial cells at these sites.
Genetic ablation of RON kinase activity in mice leads to viable
and fertile progenies with no apparent developmental defects.
However, studies showing that RON knockout (RON-KO) mice
are sensitive to LPS challenge suggested that RON signaling negatively regulates downstream effects of TLR-4 activation (31, 32).
Further studies by many groups have led to a general model in
which RON signaling promotes some of the functional and phenotypic traits of M2-like macrophages, in particular, a strong
suppression of pro-inflammatory cytokine production in response
to LPS (33, 34) or LPS + IFN-γ (35), suppression of iNOS (36),
induction of arginase-1 (37, 38), and expression of scavenger
receptors (34). Interestingly, treatment of peritoneal macrophages

Frontiers in Immunology | Molecular Innate Immunity

Studies employing different mice strains have revealed that host
genetic background significantly influences the metabolic reprograming and behavior of macrophages when exposed to LPS or
IFN-γ (39). Mills et al. first reported that whereas macrophages
from C57Bl/6 mice produced citrulline and nitric oxide (NO) in
response to LPS or IFN-γ, those from BALB/c background produced ornithine (40). Both NO and ornithine are the products
of differential arginine metabolism mediated by the expression
of enzymes iNOS and arginase-1, respectively, and this “fork in
the arginine metabolism” is recognized as one of the hallmarks of
M1 and M2 polarization (4, 39). Gene expression profiles comparing bone marrow-derived macrophages from different mice
strains further revealed that in response to LPS, timing and intensity of expression of genes differed significantly between mice
strains (41).
Interestingly, RON signaling modulated the TLR-4 responses
of macrophages between C57Bl/6 (M1-polarized) and FVB (M2polarized) mice differently (34). Whereas MSP strongly suppressed LPS-induced production of pro-inflammatory cytokines
in macrophages from FVB background, the effect was minimal
in the C57Bl/6 background. A clue to the mechanism came from
analyzing the effect of RON signaling on LPS-induced gene expression in macrophages. LPS induced the transcriptional targets of
the NF-κB and MAPK pathways early on, and RON signaling had
no effect on most of the early response genes. The late response
genes, dominated by the transcriptional targets of interferon signaling, selected NF-κB target genes, and genes associated with
tissue repair and immune tolerance was modulated variably by
RON. RON suppressed most of the targets of the interferon pathway (50% of the downregulated genes at the later time point were
targets of interferon signaling), as well as few selected targets of the
NF-κB pathway (TNF-α), but enhanced the expression of tissue
repair (EGF, PDGF, MMP9) and immune tolerance genes (IL-10,
IL-19, CTLA-2A). The kinetics of IFN-β expression in response to
TLR-4 activation was rapid (1 h) in macrophages from FVB mice,
whereas it was significantly delayed (8 h) in C57Bl/6 background.
This early upregulation of IFN-β in FVB mice was blunted by
RON signaling resulting in a strong inhibition of the expression of

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the tumor (34). These observations support the hypothesis that
ablation of RON function in the innate immune compartment
accentuates tumor-specific T-cell responses.
In the last few years, tumor-associated macrophages have
received significant attention due to their pro-tumorigenic properties, such as producing tumor-promoting and pro-angiogenic
factors and suppression of the adaptive immune response within
the tumor microenvironment (43). Polarization of macrophages
into an M2-type is one of the mechanisms that subverts the
sentinel function of the innate immune cells and make them
pro-tumorigenic.
How can this immune-modulatory property of RON be reconciled with normal tissue homeostasis? Maintaining macrophages
in M1/M2-like polarized state is important under certain physiological conditions. For example, during wound healing or during
repair of damaged tissues, macrophages serve two important functions. First, it is ready to mount an immune response to eliminate
pathogens, if the wound site gets infected, and during the same
time limit the intensity and duration of the localized immune
response to prevent further tissue damage. Second, it needs to
produce growth-promoting and tissue-rebuilding factors to accelerate healing and reverse damage. In this context, the tissue repair
or wound healing pathways of macrophages are co-opted by the
tumor to promote its own survival against immune attack and
activation of RON in the tumor microenvironment may facilitate
this conversion.
FIGURE 2 | Overview of the impact of the RON pathway on M1 versus
M2 differentiation program in the context of TLR-4 signaling. Transcript
and protein levels of IFN-β and TNF-α were compiled from experimental
values. Protein or mRNA levels at each time point are expressed as
percentage of maximal expression (100%). We propose that RON signaling
in macrophages from FVB mice preserves M2 differentiation in the
presence of TLR-4 signaling, whereas C57Bl6 macrophages maintain
polarization toward M1 cells in the presence of RON signaling. Taken from
Chaudhuri et al. (34).

interferon regulatory factors (IRFs) and target genes of the interferon pathway at later time points. The IFN-β expression however,
was minimally affected by RON signaling in C57Bl/6 macrophages
partly because of the delayed expression of IFN-β in this strain
background, and partly as a result of high expression of TNF-α, IL12, and IL-6, which by themselves can modulate the expression of
IFN-β independently of LPS (34). The final outcome of the interplay between RON and LPS signaling in these two mice strains
resulted in the stabilization of M2-polarized state in FVB mice
even in the presence of a strong M1-polarizing signal, but failed
to alter the phenotype of macrophages from C57Bl/6 background
(Figure 2).
In addition to affecting the polarized behavior of macrophages,
RON kinase-deficient FVB mice formed less number of tumors,
which developed with a delayed kinetics in two models of chemical induced carcinogenesis (34, 42). This inhibitory effect was lost
in the C57Bl/6 RON-KO background (34). Additionally, depletion of CD8+ T-cells in a transplantable fibrosarcoma model in
FVB mice suppressed the rejection of tumors in the RON-KO
background suggesting that lack of RON in the innate immune
compartment facilitates generation of a CTL response against

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CONCLUSION AND THERAPEUTIC IMPLICATIONS
M1 and M2 polarization of macrophages is dynamically controlled by changes in the tissue microenvironment. These two
functional states participate in two important activities – protection against foreign pathogens and promotion of tissue restoration
and healing after injury. Therefore, tight regulation of these two
states is critical to the health of the organism (Figure 1B). Sustained activation of M1 state can lead to excessive tissue damage
as a result of excessive inflammation, whereas prolonged activation of the M2 state can cause chronic inflammation leading
to cancer. Therapeutic targeting of certain diseases may involve
artificial manipulation of macrophage polarization. As an example, inhibiting RON function in tumor-associated macrophages
can restore tumor immunity allowing enhanced efficacy of cancer immunotherapy drugs. Similarly, enhancing RON activity in
tissue-associated macrophages can lead to efficient wound healing
and restoration of tissue damage. However, such artificial manipulation of immune cell functions has to be tightly controlled to
prevent systemic damage to the organism.

REFERENCES
1. Auffray C, Sieweke MH, Geissmann F. Blood monocytes: development, heterogeneity, and relationship with dendritic cells. Annu Rev Immunol (2009)
27:669–92. doi:10.1146/annurev.immunol.021908.132557
2. Geissmann F, Manz MG, Jung S, Sieweke MH, Merad M, Ley K. Development of
monocytes, macrophages, and dendritic cells. Science (2010) 327(5966):656–61.
doi:10.1126/science.1178331
3. Terry RL, Miller SD. Molecular control of monocyte development. Cell Immunol
(2014). doi:10.1016/j.cellimm.2014.02.008
4. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10

October 2014 | Volume 5 | Article 546 | 253

Chaudhuri

5. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas.
J Clin Invest (2012) 122(3):787–95. doi:10.1172/JCI59643
6. Martinez FO, Gordon S. The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000Prime Rep (2014) 6:13. doi:10.12703/P6-13
7. Murray PJ, Allen JE, Biswas SK, Fisher EA, Gilroy DW, Goerdt S, et al.
Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity (2014) 41(1):14–20. doi:10.1016/j.immuni.2014.06.008
8. Ivashkiv LB. Epigenetic regulation of macrophage polarization and function.
Trends Immunol (2013) 34(5):216–23. doi:10.1016/j.it.2012.11.001
9. Van Dyken SJ, Locksley RM. Interleukin-4- and interleukin-13-mediated
alternatively activated macrophages: roles in homeostasis and disease. Annu
Rev Immunol (2013) 31:317–43. doi:10.1146/annurev-immunol-032712095906
10. Gordon S, Martinez FO. Alternative activation of macrophages: mechanism and
functions. Immunity (2010) 32(5):593–604. doi:10.1016/j.immuni.2010.05.007
11. Cassetta L, Cassol E, Poli G. Macrophage polarization in health and disease.
ScientificWorldJournal (2011) 11:2391–402. doi:10.1100/2011/213962
12. Schenten D, Medzhitov R. The control of adaptive immune responses by the
innate immune system. Adv Immunol (2011) 109:87–124. doi:10.1016/B978-012-387664-5.00003-0
13. Medzhitov R. Pattern recognition theory and the launch of modern innate
immunity. J Immunol (2013) 191(9):4473–4. doi:10.4049/jimmunol.1302427
14. Jenkins KA, Mansell A. TIR-containing adaptors in toll-like receptor signalling.
Cytokine (2010) 49(3):237–44. doi:10.1016/j.cyto.2009.01.009
15. Mills CD. Macrophage arginine metabolism to ornithine/urea or nitric
oxide/citrulline: a life or death issue. Crit Rev Immunol (2001) 21(5):399–425.
doi:10.1615/CritRevImmunol.v21.i5.10
16. Zhou D, Huang C, Lin Z, Zhan S, Kong L, Fang C, et al. Macrophage polarization and function with emphasis on the evolving roles of coordinated
regulation of cellular signaling pathways. Cell Signal (2014) 26(2):192–7.
doi:10.1016/j.cellsig.2013.11.004
17. Babon JJ, Nicola NA. The biology and mechanism of action of suppressor
of cytokine signaling 3. Growth Factors (2012) 30(4):207–19. doi:10.3109/
08977194.2012.687375
18. Peti W, Page R. Molecular basis of MAP kinase regulation. Protein Sci (2013)
22(12):1698–710. doi:10.1002/pro.2374
19. Wynn TA, Barron L. Macrophages: master regulators of inflammation and fibrosis. Semin Liver Dis (2010) 30(3):245–57. doi:10.1055/s-0030-1255354
20. Gadang V, Kohli R, Myronovych A, Hui DY, Perez-Tilve D, Jaeschke A. MLK3
promotes metabolic dysfunction induced by saturated fatty acid-enriched diet.
Am J Physiol Endocrinol Metab (2013) 305(4):E549–56. doi:10.1152/ajpendo.
00197.2013
21. Luyendyk JP, Schabbauer GA, Tencati M, Holscher T, Pawlinski R,
Mackman N. Genetic analysis of the role of the PI3K-Akt pathway in
lipopolysaccharide-induced cytokine and tissue factor gene expression in monocytes/macrophages. J Immunol (2008) 180(6):4218–26. doi:10.4049/jimmunol.
180.6.4218
22. Abad C, Gomariz R, Waschek J, Leceta J, Martinez C, Juarranz Y, et al. VIP in
inflammatory bowel disease: state of the art. Endocr Metab Immune Disord Drug
Targets (2012) 12(4):316–22. doi:10.2174/187153012803832576
23. Arranz A, Doxaki C, Vergadi E, Martinez de la Torre Y, Vaporidi K, Lagoudaki
ED, et al. Akt1 and Akt2 protein kinases differentially contribute to macrophage
polarization. Proc Natl Acad Sci U S A (2012) 109(24):9517–22. doi:10.1073/
pnas.1119038109
24. Xu F, Kang Y, Zhang H, Piao Z, Yin H, Diao R, et al. Akt1-mediated regulation
of macrophage polarization in a murine model of Staphylococcus aureus pulmonary infection. J Infect Dis (2013) 208(3):528–38. doi:10.1093/infdis/jit177
25. Hu X, Herrero C, Li WP, Antoniv TT, Falck-Pedersen E, Koch AE, et al. Sensitization of IFN-gamma Jak-STAT signaling during macrophage activation. Nat
Immunol (2002) 3(9):859–66. doi:10.1038/ni828
26. Lawrence T, Natoli G. Transcriptional regulation of macrophage polarization: enabling diversity with identity. Nat Rev Immunol (2011) 11(11):750–61.
doi:10.1038/nri3088
27. Murray PJ, Wynn TA. Obstacles and opportunities for understanding
macrophage polarization. J Leukoc Biol (2011) 89(4):557–63. doi:10.1189/jlb.
0710409
28. Leonard EJ, Skeel AH. Enhancement of spreading, phagocytosis and chemotaxis by macrophage stimulating protein (MSP). Adv Exp Med Biol (1979)
121B:181–94.

Frontiers in Immunology | Molecular Innate Immunity

Macrophage polarization in health and disease

29. Wang MH, Iwama A, Skeel A, Suda T, Leonard EJ. The murine stk gene product, a transmembrane protein tyrosine kinase, is a receptor for macrophagestimulating protein. Proc Natl Acad Sci U S A (1995) 92(9):3933–7. doi:10.1073/
pnas.92.25.11751
30. Ganesan R, Kolumam GA, Lin SJ, Xie MH, Santell L, Wu TD, et al. Proteolytic
activation of pro-macrophage-stimulating protein by hepsin. Mol Cancer Res
(2011) 9(9):1175–86. doi:10.1158/1541-7786.MCR-11-0004
31. Correll PH, Iwama A, Tondat S, Mayrhofer G, Suda T, Bernstein A. Deregulated
inflammatory response in mice lacking the STK/RON receptor tyrosine kinase.
Genes Funct (1997) 1(1):69–83. doi:10.1046/j.1365-4624.1997.00009.x
32. Waltz SE, Eaton L, Toney-Earley K, Hess KA, Peace BE, Ihlendorf JR, et al. RONmediated cytoplasmic signaling is dispensable for viability but is required to
limit inflammatory responses. J Clin Invest (2001) 108(4):567–76. doi:10.1172/
JCI200111881
33. Morrison AC, Wilson CB, Ray M, Correll PH. Macrophage-stimulating protein, the ligand for the stem cell-derived tyrosine kinase/RON receptor tyrosine
kinase, inhibits IL-12 production by primary peritoneal macrophages stimulated
with IFN-gamma and lipopolysaccharide. J Immunol (2004) 172(3):1825–32.
doi:10.4049/jimmunol.172.3.1825
34. Chaudhuri A, Wilson NS, Yang B, Paler Martinez A, Liu J, Zhu C, et al.
Host genetic background impacts modulation of the TLR4 pathway by RON
in tissue-associated macrophages. Immunol Cell Biol (2013) 91(7):451–60.
doi:10.1038/icb.2013.27
35. Liu QP, Fruit K, Ward J, Correll PH. Negative regulation of macrophage activation in response to IFN-gamma and lipopolysaccharide by the STK/RON
receptor tyrosine kinase. J Immunol (1999) 163(12):6606–13.
36. Chen YQ, Fisher JH, Wang MH. Activation of the RON receptor tyrosine kinase
inhibits inducible nitric oxide synthase (iNOS) expression by murine peritoneal exudate macrophages: phosphatidylinositol-3 kinase is required for RONmediated inhibition of iNOS expression. J Immunol (1998) 161(9):4950–9.
37. Morrison AC, Correll PH. Activation of the stem cell-derived tyrosine
kinase/RON receptor tyrosine kinase by macrophage-stimulating protein results
in the induction of arginase activity in murine peritoneal macrophages.
J Immunol (2002) 168(2):853–60. doi:10.4049/jimmunol.168.2.853
38. Sharda DR, Yu S, Ray M, Squadrito ML, De Palma M, Wynn TA, et al. Regulation
of macrophage arginase expression and tumor growth by the RON receptor tyrosine kinase. J Immunol (2011) 187(5):2181–92. doi:10.4049/jimmunol.1003460
39. Mills CD, Xu GY, Johnson KM, McAdoo DJ, Hulsebosch CE. AIDA reduces
glutamate release and attenuates mechanical allodynia after spinal cord injury.
Neuroreport (2000) 11(14):3067–70. doi:10.1097/00001756-200009280-00007
40. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
41. Wells CA, Ravasi T, Faulkner GJ, Carninci P, Okazaki Y, Hayashizaki Y, et al.
Genetic control of the innate immune response. BMC Immunol (2003) 4:5.
doi:10.1186/1471-2172-4-5
42. Chan EL, Peace BE, Collins MH, Toney-Earley K, Waltz SE. RON tyrosine kinase receptor regulates papilloma growth and malignant conversion
in a murine model of skin carcinogenesis. Oncogene (2005) 24(3):479–88.
doi:10.1038/sj.onc.1208909
43. Noy R, Pollard JW. Tumor-associated macrophages: from mechanisms to therapy. Immunity (2014) 41(1):49–61. doi:10.1016/j.immuni.2014.06.010
Conflict of Interest Statement: The author declares that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 20 August 2014; paper pending published: 22 September 2014; accepted: 13
October 2014; published online: 31 October 2014.
Citation: Chaudhuri A (2014) Regulation of macrophage polarization by RON receptor
tyrosine kinase signaling. Front. Immunol. 5:546. doi: 10.3389/fimmu.2014.00546
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Chaudhuri. This is an open-access article distributed under the
terms of the Creative Commons Attribution License (CC BY). The use, distribution or
reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
accepted academic practice. No use, distribution or reproduction is permitted which
does not comply with these terms.

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MINI REVIEW ARTICLE
published: 21 August 2014
doi: 10.3389/fimmu.2014.00402

Complement, C1q, and C1q-related molecules regulate
macrophage polarization
Suzanne S. Bohlson 1 *, Sean D. O’Conner 1 , Holly Jo Hulsebus 1 , Minh-Minh Ho 2 and Deborah A. Fraser 2
1
2

Department of Microbiology and Immunology, Des Moines University, Des Moines, IA, USA
Department of Biologicial Sciences, California State University Long Beach, Long Beach, CA, USA

Edited by:
Laurel L. Lenz, National Jewish
Health, USA
Reviewed by:
Cecilia Garlanda, Istituto Clinico
Humanitas, Italy
Daisuke Kamimura, Hokkaido
University, Japan
*Correspondence:
Suzanne S. Bohlson, Department of
Microbiology and Immunology, Des
Moines University, 3200 Grand
Avenue, Des Moines, IA 50312-4198,
USA
e-mail: suzanne.bohlson@dmu.edu

Complement is a critical system of enzymes, regulatory proteins, and receptors that regulates both innate and adaptive immune responses. Natural mutations in complement molecules highlight their requirement in regulation of a variety of human conditions including
infectious disease and autoimmunity. As sentinels of the immune system, macrophages
are specialized to respond to infectious microbes, as well as normal and altered self, and
dictate appropriate immune responses. Complement components such as anaphylatoxins
(C3a and C5a) and opsonins [C3b, C1q, mannan binding lectin (MBL)] influence macrophage
responses. While anaphylatoxins C3a and C5a trigger inflammasome activation, opsonins
such as C1q and related molecules (MBL and adiponectin) downregulate inflammasome
activation and inflammation, and upregulate engulfment of apoptotic cells consistent with
a pro-resolving or M2 macrophage phenotype. This review summarizes our current understanding of the influence of the complement system on macrophage polarization with an
emphasis on C1q and related molecules.
Keywords: macrophage, complement, C1q, adiponectin, inflammasome, cytokine, phagocytosis, efferocytosis

COMPLEMENT SYSTEM
The complement system comprises over 35 cell associated and soluble molecules, which play a critical role in our innate immune
response. Activation of complement begins with a recognition
step. Recognition proteins of the complement system include C1q,
mannan binding lectin (MBL), and ficolins (ficolin-1, -2, -3).
These proteins are innate pattern recognition receptors (PRRs)
and are capable of recognizing a wide range of structures including foreign organisms, either via binding directly to their pathogen
associated molecular patterns (PAMPs) or when coated with antibody in an immune-complex (C1q) [reviewed in Ref. (1)]. In addition, these PRRs also recognize and bind to structures associated
with cellular damage/debris such as apoptotic cell associated molecular patterns (ACAMPs) like phosphatidylserine, damage associated molecular patterns (DAMPs) like oxidation neo-epitopes,
and fibrillar protein structures (2–8). Activation of complement
via the classical pathway (C1q), lectin pathway (MBL/ficolins),
or alternative pathway (C3 “tickover” autoactivation/properdin)
begins a coordinated cascade of enzymatic cleavage events generating complement protein fragments that carry out effector
functions. These include opsonization for enhanced phagocytosis,
either directly or via production of C3b, triggering inflammation
through production of anaphylatoxins C3a and C5a, and lysis of
target cells through deposition of C5b-9, the membrane attack
complex (MAC), in the complement terminal pathway (Figure 1).
COMPLEMENT DEFICIENCIES IN HUMAN DISEASE

Genetic deficiencies in complement components highlight its
important role not only in clearance of pathogens but also in
removal of dying cells/cellular debris and prevention of autoimmunity. For example, genetic deficiencies in lectin, alternative and

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terminal pathway components like MBL, factor D, properdin, C3,
C5, C6, C7, C8, and C9, increase susceptibility to infections, in
particular, by encapsulated bacteria [reviewed in Ref. (9)]. The
strong link between late complement component deficiencies (C5–
9) and recurrent neisserial infections indicates a critical role for
C5b–9 MAC deposition in direct bactericidal defense. However,
deficiency in early classical pathway component C1q is strongly
associated with development of the autoimmune disease systemic
lupus erythematosus (SLE), likely due to impaired opsonophagocytosis, and compromised removal of immune complexes and
apoptotic cells [reviewed in Ref. (10)]. Weaker associations with
SLE are also seen with deficiencies in other classical pathway components C1r/s, C2, C4, and C3. Thus, complement opsonization
via C1q recognition plays a critical role in maintaining normal tissue homeostasis and prevention of autoimmunity. While excessive
or inappropriate complement activation is associated with almost
all inflammatory or inflammation-related diseases including cancer, Alzheimer’s disease (AD), and metabolic disease [reviewed in
Ref. (11, 12)], associations of these diseases with (generally very
rare) complement deficiencies in humans have not been reported.
However, polymorphisms in MBL2 are common, and provide
extensive evidence for links between MBL levels and cardiovascular
disease [reviewed in Ref. (13)].
COMPLEMENT RECEPTORS ON MACROPHAGES

Macrophages are key players in carrying out complement effector functions. Complement components including C1q, MBL,
C3b, C4b, C3a, and C5a interact with a variety of receptors
on macrophages, leading to modulation of cytokine production/inflammatory responses and increased opsonophagocytic
clearance of targets. For example, monocytes and macrophages

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FIGURE 1 | The interaction of macrophages with complement. Innate
immune pattern recognition receptors C1q, MBL, and ficolins recognize a
number of structures including immune complexes, damaged-self molecules
expressing damage associated molecular patterns (DAMPS), apoptotic cells
(ACAMPs), and pathogens (PAMPs), leading to activation of classical and
lectin pathways, respectively, and amplification via the alternative pathway.
Complement activation has three main effector functions: cytolysis via
membrane attack complex (MAC) formation, inflammation mediated by
anaphylatoxins C3a and C5a, and opsonization leading to phagocyte clearance
via C3b deposition. However, C1q, MBL, and ficolins also have non-cascade
activation related functions and are directly opsonic, leading to enhanced

express complement receptors CR1 (CD35), CR3 (CD11b/CD18),
and CR4 (CD11c/CD18). CR1 binds complement opsonins C1q,
C3b, and C4b, which are deposited on target cells/surfaces (14)
and reviewed in Ref. (15). Binding to CR1 promotes phagocytosis of targets, along with degradation of C3b to its inactive
fragment iC3b, preventing C5-convertase activity and thus inhibiting the complement terminal pathway. CR3 and CR4 bind iC3b
and promote the phagocytosis of targets. Genetic deficiencies
in these receptors are also a risk factor for the development of
SLE [reviewed in Ref. (10, 16)]. A subset of tissue macrophages
expresses the complement receptor CRIg. Gene expression of this
receptor is associated with activated macrophages (17), and the
protein is found in human liver Kupffer cells and in subsets
of various resident tissue macrophages including alveolar and

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Complement-dependent macrophage polarization

clearance of targets and phagocyte activation. Macrophages express a
number of receptors that recognize complement components. The gC1qR
and cC1qR bind the globular heads and collagen-like tail domains of C1q,
respectively. However, additional C1q receptors likely exist (C1qR), some of
which may also recognize the closely related collagen-like domains of other
defense collagen family members, MBL and ficolins. Additional complement
receptors on macrophages CR1, CR3, CR4, and CRIg recognize
C3b-opsonized targets either as intact C3b or its degradation fragment iC3b.
Macrophages also express receptors for complement activation fragments
C3a (C3aR) and C5a (C5aR1 and C5aR2). Thus, macrophages are key players
in carrying out the effector functions of complement activation.

synovial macrophages (18). CRIg binds to C3b, and its degradation product, iC3b, and was shown to be important in clearance
of C3b-opsonized pathogens from the circulation.
Beyond CR1, macrophages express other receptors capable of
binding C1q. These include the ubiquitously co-expressed molecules gC1qR and cC1qR. gC1qR binds to the globular heads of
C1q (19), whereas cC1qR (calreticulin) interacts with both the
collagen-like tail and globular head domains of C1q (4). C1q
opsonized targets are internalized more rapidly than in the absence
of C1q via interaction of the collagen-like tail with a receptor on
phagocytes (20). In addition, C1q bound to a variety of targets
modulates macrophage inflammatory responses via its collagenlike domain (5, 21). The collagen-like domain of C1q is a feature
shared with other so-called “defense collagens.” This family of

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Complement-dependent macrophage polarization

molecules includes MBL and ficolins, which also trigger enhanced
phagocytosis [reviewed in Ref. (1)] and modulate cytokine production (22). While the phagocytic receptor is not definitively
identified for all targets, involvement of the cC1qR has been implicated in the C1q- and MBL-mediated removal of apoptotic cells
by macrophages (6). An additional family of proteins, termed
C1q/TNF-related proteins (CTRPs), contains C-terminal globular domains homologous to C1q (23, 24). Adiponectin is perhaps
the best characterized member of the CTRP family, and interacts with macrophages, regulating inflammatory responses similar
to C1q (25). However, the receptors for the metabolic actions of
adiponectin were shown to be distinct from the C1qR, and include
AdipoR1 and AdiopR2 (26).

misleading and over simplified. In vivo, macrophages are constantly encountering various external signals leading to a very
fluid existence in terms of phenotypes (38). Even within the same
site and population, macrophages can express different and constantly changing phenotypes, termed macrophage plasticity (39).
Macrophage polarization is a result of a combination of external signals macrophages receive from their environment. Common signals that have been investigated include the Th1 cytokine
interferon-γ (IFN-γ) and various PAMPs, which are associated
with M1 polarization and the Th2 cytokines IL-4/IL-13, which are
associated with M2 polarization. Here, we will review some recent
work illuminating the role of the complement system in regulating
macrophage activation and polarization.

COMPLEMENT ANAPHYLATOXINS

COMPLEMENT ANAPHYLATOXINS REGULATE MACROPHAGE
ACTIVATION

Complement anaphylatoxins C3a and C5a are soluble complement
fragments produced from C3 or C5 by activation of the C3- or
C5-convertase enzyme complex, respectively. They carry out their
biological functions via interactions with three receptors. These
include the C3a receptor (C3aR) (27), which binds C3a but not its
degradation product C3a-desArg, the C5a receptor (C5aR1) (28),
which binds C5a, and C5a receptor-like 2 (C5aR2, C5L2) (29),
which binds C5a but has greater affinity for C5a-desArg. C5aR2
lacks signaling capabilities (30), and thus, is considered a decoy
receptor, capable of sequestering the bioavailability of C5a/C5adesArg and limiting their ability to activate via the C5aR1. C5aR2
may also bind C3a-desArg, but this is controversial (31). Cellular
expression of the anaphylatoxin receptors is widespread but particularly includes immune cells like monocytes and macrophages (32,
33). Interestingly, LPS, associated with M1 macrophage polarization increases gene expression of C5aR1 in macrophages (34) while
IL-4, associated with M2 macrophage polarization downregulates
C5aR1 expression (35).

MACROPHAGE POLARIZATION
Macrophages are grouped as M1 and M2 in accordance with
the Th1/Th2 nomenclature, and this terminology describes two
macrophage phenotypes: the pro-inflammatory/classically activated macrophage (M1) and the pro-resolving/alternatively activated macrophage (M2). The expression of a variety of genes
has been associated with macrophage polarization, most notably,
the machinery required for enzymatic breakdown of arginine in
pro-resolving and pro-inflammatory macrophages. By expressing
arginase, pro-resolving macrophages generate ornithine, which
promotes proliferation and repair, whereas pro-inflammatory
(M1) macrophages express inducible nitric oxide synthase and
generate nitric oxide (NO), an important molecule in host defense
against invading pathogens, which also inhibits cell proliferation [reviewed in Ref. (36)]. Intermediates in the two enzymatic
pathways act antagonistically, inhibiting the other when they are
activated (37). Consistent with a role in promotion of inflammation, M1 macrophages are also often associated with an increased
production of pro-inflammatory cytokines such as TNFα and IL1β. In contrast, M2 macrophages, which are pro-resolving, are
often associated with increased production of anti-inflammatory
cytokine IL-10.
Although the idea of macrophages having separate phenotypes is helpful in defining their action, it is also somewhat
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Complement anaphylatoxins, C3a and C5a, are pro-inflammatory
and trigger monocyte and macrophage activation through various signaling mechanisms. For example, upon LPS stimulation
in human monocytes, C3a induces NLRP3 inflammasome activation through an increase in ATP release mediated by extracellular
signal-regulated kinase 1/2 (ERK1/2) (40). Samstad et al. found
that C5a produced during complement activation by cholesterol
crystals, influenced inflammation through NLRP3 inflammasome
activation and IL-1β and TNFα release, and increased the production of reactive oxygen species (ROS) (41). C5a was also correlated
with IL-6 induction and development of inflammatory T-helper
17 cells (42), as well as affecting IL-17 and IL-23 production
(43). Interestingly, sublethal MAC (C5b-9) deposition rather than
C3a or C5a was demonstrated to trigger inflammasome activation in murine dendritic cells following LPS stimulation (44). In
the CNS, C5a provides a chemotactic and activation signal for
microglia and astrocytes [reviewed in Ref. (45)]. It can also synergize with damage signals such as amyloid beta (Aβ) to trigger
enhanced inflammatory cytokine production (46). Thus, activation of complement by extracellular Aβ plaques can exacerbate
inflammation and may play a substantial role in the pathogenesis of AD. Indeed, treatment with a C5a receptor antagonist was
shown to decrease deposition of fibrillar Aβ and inflammatory glia
and improve cognitive performance in mice models of AD (47).
Many other diseases are associated with anaphylatoxin signaling,
including allergic, infectious, autoimmune diseases, and cancer
(48). Clearly, the pro-inflammatory signaling provided by the anaphylatoxins contributes to both beneficial (pathogen clearing) and
detrimental (inflammatory disease-related) inflammation.

COMPLEMENT OPSONINS REGULATE MACROPHAGE
ACTIVATION
C3b-MEDIATED OPSONIZATION

One of the major effector functions of the complement system is the tagging, or opsonization, of pathogens and/or cellular
debris for clearance by phagocytes. Complement component C3
is the most abundant complement component in blood at about
1.2 mg/ml, and as such, permissible surfaces become readily coated
in C3b and iC3b following cleavage of C3. CR3 is a major phagocytic receptor expressed on macrophages that is involved in clearance of iC3b opsonized particles (Figure 1). In contrast to other
phagocytic receptors such as Fcγ receptors, engulfment of iC3b
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Complement-dependent macrophage polarization

coated particles via CR3 has traditionally been considered antiinflammatory. For example, CR3-dependent engulfment does not
activate the arachidonic acid cascade (49) or the release of toxic
oxygen products (50). In early experiments assessing macrophage
heterogeneity, Stein and colleagues demonstrated increased secretion of TNFα from macrophages elicited into the peritoneal cavity
by inflammatory mediators (e.g., thioglycollate) when compared
to resident peritoneal macrophages. However, independent of the
macrophage phenotype, all macrophages failed to secrete significant levels of TNFα following ligation of CR3 whereas ligation of
Fcγ receptors led to TNFα release by all macrophage subsets (51).
In studies with the macrophage intracellular pathogen Mycobacterium avium, C3-independent phagocytosis of M. avium resulted
in enhanced TNFα production (52, 53). More recent reports
assessing pathogenesis of Francisella tularensis support a role
for CR3 in inducing immune suppression and facilitating infection with this intracellular pathogen (54). While CR3-dependent
immune suppression is detrimental in the course of F. tularensis
infection or other infectious disease processes, it is beneficial in
the context of clearance of apoptotic cells and/or cellular debris.
COMPLEMENT-DEPENDENT ENGULFMENT OF APOPTOTIC CELLS

Complement components readily coat the surface of apoptotic
cells and facilitate ingestion by macrophages (55, 56). Ingestion
of apoptotic cells is a silent process accompanied by the production of anti-inflammatory cytokines TGFβ and IL-10 (57).
Complement mediated opsonization of apoptotic cells is largely
dependent on the classical complement pathway, and deficiencies in early components of the classical pathway (C1q, C4, and
C2) result in inefficient disposal of apoptotic cells and subsequent autoimmunity (58–60). While C1q deficiency results in
development of lupus in virtually all cases, absence of C3, C2,
and C4 results in lupus at lower frequency indicating a role for
C1q beyond classical complement pathway activation in regulation of the immune response (61). Recent reports have indicated
that C1q regulates the monocyte/macrophage/dendritic cell phenotype leading to development of a phagocyte that is specialized
to resolve inflammation. Specifically, the C1q-stimulated phagocyte is pro-efferocytic and anti-inflammatory (Figure 2). As such,
complement opsonins are mediating activity beyond the immediate stimulation of enhanced phagocytosis; they are inducing
a macrophage phenotype, or polarizing macrophages toward a
pro-resolving phenotype. This would be consistent with observation that C1q deficiency results in autoimmunity and chronic
inflammation.
Korb and Ahearn were the first to describe a role for C1q in the
clearance of apoptotic cells, and suggested that C1q-dependent
engulfment of apoptotic cells was important in prevention of
autoimmunity in lupus (62). There has been wide support for
this hypothesis, and Bhatia et al. demonstrated that removal of
apoptotic cells by C1q was also important in prevention of the
inflammatory disease atherosclerosis (63). However, the receptors/signal transduction pathway leading to C1q-dependent efferocytosis has not been clearly delineated [reviewed in Ref. (56)].
C1q binds to apoptotic cells and serves as a bridging molecule linking the apoptotic cell to the phagocyte via calreticulin (cC1qR)
and its binding partner, the phagocytic receptor LRP (CD91)

Frontiers in Immunology | Inflammation

FIGURE 2 | Complement regulates macrophage polarization. In culture,
M1 macrophages (inflammatory, pathogen clearing) are induced by IFNγ or
PAMPs and are characterized by production of iNOS and pro-inflammatory
cytokines such as TNFα and IL-1β. M2 macrophages (resolving, apoptotic
cell and damaged molecule clearing) are induced by IL-4 and characterized
by arginase and anti-inflammatory cytokine IL-10 production. Complement
components C3a, C5a, and C5b-9 modulate cytokine production in
macrophages toward an inflammatory (M1-like) phenotype. Apoptotic cells
and targets opsonized with complement components C1q or C3b increase
clearance and modulate cytokine production in macrophages toward an
anti-inflammatory, resolving (M2-like) phenotype and can block
PAMP-mediated pro-inflammatory signaling. Thus, complement plays a dual
role in macrophage activation and polarization depending on the target.

(6). However, macrophages deficient in LRP still respond to C1q
with enhanced phagocytosis and efferocytosis indicating that there
are multiple mechanisms of C1q-dependent engulfment (64). We
demonstrated that mouse bone marrow derived macrophages and
peritoneal macrophages stimulated with C1q upregulated expression of engulfment machinery including Mer tyrosine kinase and
the MerTK ligand Gas6, leading to development of a macrophage
that is primed for efferocytosis (65). More recently, we showed that
this pathway is shared with a C1q homolog, adiponectin (66) and
not with MBL, a C1q-related collectin (65).
C1q AND ADIPONECTIN MEDIATE EFFEROCYTOSIS VIA A SHARED
PATHWAY

Adiponectin is referred to as an adipokine; it is produced by
adipocytes and is secreted into circulation where it modulates
biological responses via several receptors including adiponectin
receptor 1 (AdipoR1), adiponectin receptor 2 (AdipoR2), or
T cadherin (T-cad) (67, 68). More recent studies have suggested there is an additional receptor on macrophages that mediates adiponectin signaling; however, this receptor has not been
identified (69). Adiponectin signaling leads to activation of 50
adenosine monophosphate-activated protein kinase (AMPK) and
peroxisome proliferator-activated receptor alpha (PPARα), and
adiponectin-dependent regulation of metabolism is mediated via

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these key signaling nodes (70). Much information has been gathered regarding the role of AMPK in the regulation of metabolic
activity; however, recent studies suggest that AMPK also influences
the immune response, including macrophage cytokine expression and phagocytosis. For example, silencing of AMPK inhibits
LPS- and fatty acid-mediated inflammation in macrophages
(71). In addition, AMPK activation is associated with enhanced
phagocytosis and efferocytosis, as well as macrophage polarization (72, 73). Similarly, C1q stimulates enhanced phagocytosis and diminution of pro-inflammatory cytokine production
from myeloid cells, and we recently demonstrated that C1q and
adiponectin-dependent Mer expression and efferocytosis require
activation of AMPK (66). These studies have begun to define
the mechanism by which C1q and related opsonins modulate
macrophage activation.
C1q AND MBL INHIBIT PRO-INFLAMMATORY AND PROMOTE
ANTI-INFLAMMATORY CYTOKINE PRODUCTION

In line with these observations, Fraser and colleagues demonstrated that both C1q-stimulated human monocytes and C1qstimulated mouse microglia produce increased anti-inflammatory
IL-10 and decreased pro-inflammatory cytokines following stimulation with the TLR4 ligand LPS (21, 22). Similar activity was
shown for MBL indicating that this is a distinct mechanism
for macrophage activation, independent of AMPK-mediated Mer
expression since MBL failed to upregulate Mer-dependent efferocytosis. C1q/MBL-dependent activation of NFκB p50p50 homodimers were suggested to contribute to the anti-inflammatory phenotype via competitive inhibition of pro-inflammatory NFκB
p50p65 heterodimer activation and/or via transcriptional activation of IL-10 (74). The same group demonstrated that C1q
promoted M2 polarization and limited inflammasome activation in human monocyte derived macrophages (75). Interferonalpha (IFN-α) is a pro-inflammatory cytokine that contributes
to dendritic cell activation, a breakdown in peripheral tolerance and autoimmunity (76). C1q modulates IFN-α production from human phagocytes in response to a variety of stimuli (75, 77, 78). Santer and colleagues demonstrated that C1q
deficiency in human lupus patients resulted in elevated IFNα levels in serum and cerebrospinal fluid, and that C1q suppressed immune-complex stimulated IFN-α production from
human monocytes (78–80). The anti-inflammatory effects of
C1q are not limited to apoptotic cells. C1q has also been
shown to enhance the uptake of atherogenic forms of lipoproteins such as oxidized or acetylated LDL (oxLDL, AcLDL)
(81). During clearance of oxLDL by macrophages, C1q also
modulates cytokine production toward an anti-inflammatory,
resolving phenotype and dampens transcriptional activity by
p50/p65 NFκB heterodimers, which may be important in limiting inflammation in the early atherosclerotic lesion (82).
Combined, these data further support the hypothesis that
C1q programs macrophages toward an anti-inflammatory, proefferocytic/phagocytic phenotype. Future studies should delineate the relative activity of C1q-dependent constitutive efferocytosis/phagocytosis versus programed polarization in the
contribution toward protection from autoimmune and inflammatory disease.

www.frontiersin.org

Complement-dependent macrophage polarization

CONCLUDING REMARKS
The complement system has traditionally been considered an arm
of the innate immune response required for promotion of inflammation and pathogen clearance. While these functions of complement are essential to host defense, more recent advances demonstrate a novel role for components of the complement system
in resolution of inflammation and protection from autoimmune
and inflammatory diseases including SLE, neurodegenerative disease, and atherosclerosis. In particular, complement component
C1q directs macrophage polarization leading to generation of
pro-resolving macrophages that promote clearance of apoptotic
cells with diminished pro-inflammatory cytokine production and
increased anti-inflammatory cytokine production. C1q is synthesized by macrophages in response to tissue injury and is likely to
be an important signal in resolution of inflammation independent
of other complement components. Moreover, C1q-related molecules such as the complement component MBL, and the adipokine
adiponectin, also downregulate macrophage-mediated inflammatory responses and upregulate efferocytosis. Identification of
the molecular mechanisms by which these molecules govern
macrophage activation, as well as their relative contribution to disease resolution, should reveal pathways to target for development
of novel therapeutics in autoimmune and inflammatory disease.

AUTHOR CONTRIBUTIONS
Suzanne Slater Bohlson outlined the review, invited co-authors,
wrote the section on complement opsonins, extensively edited all
sections, and prepared the manuscript for publication. Sean David
O’Conner drafted the section on macrophage polarization. Holly
Jo Hulsebus drafted the section on complement anaphylatoxins.
Minh-Minh Ho designed and created the figures and figure legends. Deborah Ann Fraser wrote the section on complement and
extensively edited the entire manuscript. All authors reviewed the
final document prior to submission.

ACKNOWLEDGMENTS
This work was supported by NIH NIAID R56AI099010-02 to
Suzanne Slater Bohlson.

REFERENCES
1. Fraser DA, Tenner AJ. Directing an appropriate immune response: the role of
defense collagens and other soluble pattern recognition molecules. Curr Drug
Targets (2008) 9(2):113–22. doi:10.2174/138945008783502476
2. Biro A, Thielens NM, Cervenak L, Prohaszka Z, Fust G, Arlaud GJ. Modified low
density lipoproteins differentially bind and activate the C1 complex of complement. Mol Immunol (2007) 44(6):1169–77. doi:10.1016/j.molimm.2006.06.013
3. Jiang H, Burdick D, Glabe CG, Cotman CW, Tenner AJ. Beta-amyloid activates
complement by binding to a specific region of the collagen-like domain of the
C1q A chain. J Immunol (1994) 152(10):5050–9.
4. Paidassi H, Tacnet-Delorme P,Verneret M, Gaboriaud C, Houen G, Duus K, et al.
Investigations on the C1q-calreticulin-phosphatidylserine interactions yield
new insights into apoptotic cell recognition. J Mol Biol (2011) 408(2):277–90.
doi:10.1016/j.jmb.2011.02.029
5. Fraser DA, Pisalyaput K, Tenner AJ. C1q enhances microglial clearance of apoptotic neurons and neuronal blebs, and modulates subsequent inflammatory
cytokine production. J Neurochem (2010) 112(3):733–43. doi:10.1111/j.14714159.2009.06494.x
6. Ogden CA, deCathelineau A, Hoffmann PR, Bratton D, Ghebrehiwet B, Fadok
VA, et al. C1q and mannose binding lectin engagement of cell surface calreticulin and CD91 initiates macropinocytosis and uptake of apoptotic cells. J Exp
Med (2001) 194(6):781–95. doi:10.1084/jem.194.6.781

August 2014 | Volume 5 | Article 402 | 259

Bohlson et al.

7. Honore C, Hummelshoj T, Hansen BE, Madsen HO, Eggleton P, Garred P. The
innate immune component ficolin 3 (Hakata antigen) mediates the clearance
of late apoptotic cells. Arthritis Rheum (2007) 56(5):1598–607. doi:10.1002/art.
22564
8. Matsushita M. Ficolins in complement activation. Mol Immunol (2013)
55(1):22–6. doi:10.1016/j.molimm.2012.08.017
9. Skattum L, van Deuren M, van der Poll T, Truedsson L. Complement deficiency states and associated infections. Mol Immunol (2011) 48(14):1643–55.
doi:10.1016/j.molimm.2011.05.001
10. Leffler J, Bengtsson AA, Blom AM. The complement system in systemic lupus
erythematosus: an update. Ann Rheum Dis (2014) 73(9):1601–6. doi:10.1136/
annrheumdis-2014-205287
11. Veerhuis R, Nielsen HM, Tenner AJ. Complement in the brain. Mol Immunol
(2011) 48(14):1592–603. doi:10.1016/j.molimm.2011.04.003
12. Hertle E, Stehouwer CD, van Greevenbroek MM. The complement system in
human cardiometabolic disease. Mol Immunol (2014). doi:10.1016/j.molimm.
2014.06.031
13. Pagowska-Klimek I, Cedzynski M. Mannan-binding lectin in cardiovascular disease. Biomed Res Int (2014) 2014:616817. doi:10.1155/2014/616817
14. Klickstein LB, Barbashov SF, Liu T, Jack RM, Nicholson-Weller A. Complement receptor type 1 (CR1, CD35) is a receptor for C1q. Immunity (1997)
7(3):345–55. doi:10.1016/S1074-7613(00)80356-8
15. Krych-Goldberg M, Atkinson JP. Structure-function relationships of complement receptor type 1. Immunol Rev (2001) 180:112–22. doi:10.1034/j.1600065X.2001.1800110.x
16. Fagerholm SC, MacPherson M, James MJ, Sevier-Guy C, Lau CS. The
CD11b-integrin (ITGAM) and systemic lupus erythematosus. Lupus (2013)
22(7):657–63. doi:10.1177/0961203313491851
17. Walker MG. Z39Ig is co-expressed with activated macrophage genes. Biochim
Biophys Acta (2002) 1574(3):387–90. doi:10.1016/S0167-4781(01)00358-X
18. Helmy KY, Katschke KJ Jr, Gorgani NN, Kljavin NM, Elliott JM, Diehl L,
et al. CRIg: a macrophage complement receptor required for phagocytosis
of circulating pathogens. Cell (2006) 124(5):915–27. doi:10.1016/j.cell.2005.12.
039
19. Ghebrehiwet B, Lim BL, Peerschke EI, Willis AC, Reid KB. Isolation, cDNA
cloning, and overexpression of a 33-kD cell surface glycoprotein that binds to
the globular “heads” of C1q. J Exp Med (1994) 179(6):1809–21. doi:10.1084/
jem.179.6.1809
20. Bobak DA, Gaither TA, Frank MM, Tenner AJ. Modulation of FcR function by
complement: subcomponent C1q enhances the phagocytosis of IgG-opsonized
targets by human monocytes and culture-derived macrophages. J Immunol
(1987) 138(4):1150–6.
21. Fraser DA, Laust AK, Nelson EL, Tenner AJ. C1q differentially modulates
phagocytosis and cytokine responses during ingestion of apoptotic cells by
human monocytes, macrophages, and dendritic cells. J Immunol (2009)
183(10):6175–85. doi:10.4049/jimmunol.0902232
22. Fraser DA, Bohlson SS, Jasinskiene N, Rawal N, Palmarini G, Ruiz S, et al.
C1q and MBL, components of the innate immune system, influence monocyte
cytokine expression. J Leukoc Biol (2006) 80(1):107–16. doi:10.1189/jlb.1105683
23. Ghai R, Waters P, Roumenina LT, Gadjeva M, Kojouharova MS, Reid KB,
et al. C1q and its growing family. Immunobiology (2007) 212(4–5):253–66.
doi:10.1016/j.imbio.2006.11.001
24. Schaffler A, Buechler C. CTRP family: linking immunity to metabolism. Trends
Endocrinol Metab (2012) 23(4):194–204. doi:10.1016/j.tem.2011.12.003
25. Yokota T, Oritani K, Takahashi I, Ishikawa J, Matsuyama A, Ouchi N, et al.
Adiponectin, a new member of the family of soluble defense collagens, negatively regulates the growth of myelomonocytic progenitors and the functions of
macrophages. Blood (2000) 96(5):1723–32.
26. Yamauchi T, Nio Y, Maki T, Kobayashi M, Takazawa T, Iwabu M, et al. Targeted
disruption of AdipoR1 and AdipoR2 causes abrogation of adiponectin binding
and metabolic actions. Nat Med (2007) 13(3):332–9. doi:10.1038/nm1557
27. Ames RS, Li Y, Sarau HM, Nuthulaganti P, Foley JJ, Ellis C, et al. Molecular
cloning and characterization of the human anaphylatoxin C3a receptor. J Biol
Chem (1996) 271(34):20231–4. doi:10.1074/jbc.271.34.20231
28. Gerard NP, Gerard C. The chemotactic receptor for human C5a anaphylatoxin.
Nature (1991) 349(6310):614–7. doi:10.1038/349614a0
29. Cain SA, Monk PN. The orphan receptor C5L2 has high affinity binding sites
for complement fragments C5a and C5a des-Arg(74). J Biol Chem (2002)
277(9):7165–9. doi:10.1074/jbc.C100714200

Frontiers in Immunology | Inflammation

Complement-dependent macrophage polarization

30. Okinaga S, Slattery D, Humbles A, Zsengeller Z, Morteau O, Kinrade
MB, et al. C5L2, a nonsignaling C5A binding protein. Biochemistry (2003)
42(31):9406–15. doi:10.1021/bi034489v
31. Kalant D, Cain SA, Maslowska M, Sniderman AD, Cianflone K, Monk PN. The
chemoattractant receptor-like protein C5L2 binds the C3a des-Arg77/acylationstimulating protein. J Biol Chem (2003) 278(13):11123–9. doi:10.1074/jbc.
M206169200
32. Monk PN, Scola AM, Madala P, Fairlie DP. Function, structure and therapeutic
potential of complement C5a receptors. Br J Pharmacol (2007) 152(4):429–48.
doi:10.1038/sj.bjp.0707332
33. Zwirner J, Gotze O, Begemann G, Kapp A, Kirchhoff K, Werfel T. Evaluation
of C3a receptor expression on human leucocytes by the use of novel monoclonal antibodies. Immunology (1999) 97(1):166–72. doi:10.1046/j.1365-2567.
1999.00764.x
34. Hunt JR, Martin CB, Martin BK. Transcriptional regulation of the murine
C5a receptor gene: NF-Y is required for basal and LPS induced expression
in macrophages and endothelial cells. Mol Immunol (2005) 42(11):1405–15.
doi:10.1016/j.molimm.2005.01.002
35. Soruri A, Kiafard Z, Dettmer C, Riggert J, Kohl J, Zwirner J. IL-4 downregulates anaphylatoxin receptors in monocytes and dendritic cells and impairs
anaphylatoxin-induced migration in vivo. J Immunol (2003) 170(6):3306–14.
doi:10.4049/jimmunol.170.6.3306
36. Mills CD. M1 and M2 macrophages: oracles of health and disease. Crit Rev
Immunol (2012) 32(6):463–88. doi:10.1615/CritRevImmunol.v32.i6.10
37. Morris SM Jr. Recent advances in arginine metabolism: roles and regulation of
the arginases. Br J Pharmacol (2009) 157(6):922–30. doi:10.1111/j.1476-5381.
2009.00278.x
38. Martinez FO, Gordon S. The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000Prime Rep (2014) 6:13. doi:10.12703/P6-13
39. Mills CD. Macrophage arginine metabolism to ornithine/urea or nitric
oxide/citrulline: a life or death issue. Crit Rev Immunol (2001) 21(5):399–425.
doi:10.1615/CritRevImmunol.v21.i5.10
40. Asgari E, Le Friec G, Yamamoto H, Perucha E, Sacks SS, Kohl J, et al. C3a modulates IL-1beta secretion in human monocytes by regulating ATP efflux and
subsequent NLRP3 inflammasome activation. Blood (2013) 122(20):3473–81.
doi:10.1182/blood-2013-05-502229
41. Samstad EO, Niyonzima N, Nymo S, Aune MH, Ryan L, Bakke SS, et al. Cholesterol crystals induce complement-dependent inflammasome activation and
cytokine release. J Immunol (2014) 192(6):2837–45. doi:10.4049/jimmunol.
1302484
42. Fang C, Zhang X, Miwa T, Song WC. Complement promotes the development
of inflammatory T-helper 17 cells through synergistic interaction with toll-like
receptor signaling and interleukin-6 production. Blood (2009) 114(5):1005–15.
doi:10.1182/blood-2009-01-198283
43. Grailer JJ, Bosmann M, Ward PA. Regulatory effects of C5a on IL-17A, IL-17F,
and IL-23. Front Immunol (2013) 3:387. doi:10.3389/fimmu.2012.00387
44. Laudisi F, Spreafico R, Evrard M, Hughes TR, Mandriani B, Kandasamy M, et al.
Cutting edge: the NLRP3 inflammasome links complement-mediated inflammation and IL-1beta release. J Immunol (2013) 191(3):1006–10. doi:10.4049/
jimmunol.1300489
45. Yao J, Harvath L, Gilbert DL, Colton CA. Chemotaxis by a CNS macrophage,
the microglia. J Neurosci Res (1990) 27(1):36–42. doi:10.1002/jnr.490270106
46. O’Barr S, Cooper NR. The C5a complement activation peptide increases IL1beta and IL-6 release from amyloid-beta primed human monocytes: implications for Alzheimer’s disease. J Neuroimmunol (2000) 109(2):87–94. doi:10.
1016/S0165-5728(00)00291-5
47. Fonseca MI, Ager RR, Chu SH, Yazan O, Sanderson SD, LaFerla FM, et al.
Treatment with a C5aR antagonist decreases pathology and enhances behavioral performance in murine models of Alzheimer’s disease. J Immunol (2009)
183(2):1375–83. doi:10.4049/jimmunol.0901005
48. Klos A, Tenner AJ, Johswich KO, Ager RR, Reis ES, Kohl J. The role of the
anaphylatoxins in health and disease. Mol Immunol (2009) 46(14):2753–66.
doi:10.1016/j.molimm.2009.04.027
49. Aderem AA, Wright SD, Silverstein SC, Cohn ZA. Ligated complement receptors
do not activate the arachidonic acid cascade in resident peritoneal macrophages.
J Exp Med (1985) 161(3):617–22. doi:10.1084/jem.161.3.617
50. Wright SD, Silverstein SC. Receptors for C3b and C3bi promote phagocytosis
but not the release of toxic oxygen from human phagocytes. J Exp Med (1983)
158(6):2016–23. doi:10.1084/jem.158.6.2016

August 2014 | Volume 5 | Article 402 | 260

Bohlson et al.

51. Stein M, Gordon S. Regulation of tumor necrosis factor (TNF) release by
murine peritoneal macrophages: role of cell stimulation and specific phagocytic
plasma membrane receptors. Eur J Immunol (1991) 21(2):431–7. doi:10.1002/
eji.1830210227
52. Bohlson SS, Strasser JA, Bower JJ, Schorey JS. Role of complement in Mycobacterium avium pathogenesis: in vivo and in vitro analyses of the host response to
infection in the absence of complement component C3. Infect Immun (2001)
69(12):7729–35. doi:10.1128/IAI.69.12.7729-7735.2001
53. Irani VR, Maslow JN. Induction of murine macrophage TNF-alpha synthesis
by Mycobacterium avium is modulated through complement-dependent interaction via complement receptors 3 and 4 in relation to M. avium glycopeptidolipid. FEMS Microbiol Lett (2005) 246(2):221–8. doi:10.1016/j.femsle.2005.
04.008
54. Dai S, Rajaram MV, Curry HM, Leander R, Schlesinger LS. Fine tuning inflammation at the front door: macrophage complement receptor 3-mediates phagocytosis and immune suppression for Francisella tularensis. PLoS Pathog (2013)
9(1):e1003114. doi:10.1371/journal.ppat.1003114
55. Mevorach D, Mascarenhas JO, Gershov D, Elkon KB. Complement-dependent
clearance of apoptotic cells by human macrophages. J Exp Med (1998)
188(12):2313–20. doi:10.1084/jem.188.12.2313
56. Galvan MD, Greenlee-Wacker MC, Bohlson SS. C1q and phagocytosis: the
perfect complement to a good meal. J Leukoc Biol (2012) 92(3):489–97.
doi:10.1189/jlb.0212099
57. Fadok VA, Bratton DL, Konowal A, Freed PW, Westcott JY, Henson
PM. Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production through autocrine/paracrine mechanisms involving TGF-beta, PGE2, and PAF. J Clin Invest (1998) 101(4):890–8. doi:10.1172/
JCI1112
58. Gullstrand B, Martensson U, Sturfelt G, Bengtsson AA, Truedsson L. Complement classical pathway components are all important in clearance of apoptotic and secondary necrotic cells. Clin Exp Immunol (2009) 156(2):303–11.
doi:10.1111/j.1365-2249.2009.03896.x
59. Botto M,Walport MJ. C1q, autoimmunity and apoptosis. Immunobiology (2002)
205(4–5):395–406. doi:10.1078/0171-2985-00141
60. Taylor PR, Carugati A, Fadok VA, Cook HT,Andrews M, Carroll MC, et al. A hierarchical role for classical pathway complement proteins in the clearance of apoptotic cells in vivo. J Exp Med (2000) 192(3):359–66. doi:10.1084/jem.192.3.359
61. Walport MJ, Davies KA, Botto M. C1q and systemic lupus erythematosus.
Immunobiology (1998) 199(2):265–85. doi:10.1016/S0171-2985(98)80032-6
62. Korb LC, Ahearn JM. C1q binds directly and specifically to surface blebs of
apoptotic human keratinocytes: complement deficiency and systemic lupus erythematosus revisited. J Immunol (1997) 158(10):4525–8.
63. Bhatia VK, Yun S, Leung V, Grimsditch DC, Benson GM, Botto MB, et al. Complement C1q reduces early atherosclerosis in low-density lipoprotein receptordeficient mice. Am J Pathol (2007) 170(1):416–26. doi:10.2353/ajpath.2007.
060406
64. Lillis AP, Greenlee MC, Mikhailenko I, Pizzo SV, Tenner AJ, Strickland DK, et al.
Murine low-density lipoprotein receptor-related protein 1 (LRP) is required
for phagocytosis of targets bearing LRP ligands but is not required for C1qtriggered enhancement of phagocytosis. J Immunol (2008) 181(1):364–73.
doi:10.4049/jimmunol.181.1.364
65. Galvan MD, Foreman DB, Zeng E, Tan JC, Bohlson SS. Complement component C1q regulates macrophage expression of Mer tyrosine kinase to promote
clearance of apoptotic cells. J Immunol (2012) 188(8):3716–23. doi:10.4049/
jimmunol.1102920
66. Galvan MD, Hulsebus H, Heitker T, Zeng E, Bohlson S. Complement protein C1q and adiponectin stimulate Mer tyrosine kinase dependent engulfment of apoptotic cells through a shared pathway. J Innate Immun (2014).
doi:10.1159/000363295
67. Kadowaki T, Yamauchi T. Adiponectin and adiponectin receptors. Endocr Rev
(2005) 26(3):439–51. doi:10.1210/er.2005-0005
68. Yamauchi T, Kamon J, Ito Y, Tsuchida A, Yokomizo T, Kita S, et al. Cloning of
adiponectin receptors that mediate antidiabetic metabolic effects. Nature (2003)
423(6941):762–9. doi:10.1038/nature01705
69. Awazawa M, Ueki K, Inabe K, Yamauchi T, Kubota N, Kaneko K, et al.
Adiponectin enhances insulin sensitivity by increasing hepatic IRS-2 expression via a macrophage-derived IL-6-dependent pathway. Cell Metab (2011)
13(4):401–12. doi:10.1016/j.cmet.2011.02.010

www.frontiersin.org

Complement-dependent macrophage polarization

70. Heiker JT, Kosel D, Beck-Sickinger AG. Molecular mechanisms of signal
transduction via adiponectin and adiponectin receptors. Biol Chem (2010)
391(9):1005–18. doi:10.1515/bc.2010.104
71. Sag D, Carling D, Stout RD, Suttles J. Adenosine 5’-monophosphate-activated
protein kinase promotes macrophage polarization to an anti-inflammatory
functional phenotype. J Immunol (2008) 181(12):8633–41. doi:10.4049/
jimmunol.181.12.8633
72. Mounier R, Theret M, Arnold L, Cuvellier S, Bultot L, Goransson O, et al. AMPKalpha1 regulates macrophage skewing at the time of resolution of inflammation during skeletal muscle regeneration. Cell Metab (2013) 18(2):251–64.
doi:10.1016/j.cmet.2013.06.017
73. Jiang S, Park DW, Stigler WS, Creighton J, Ravi S, Darley-Usmar V, et al. Mitochondria and AMP-activated protein kinase-dependent mechanism of efferocytosis. J Biol Chem (2013) 288(36):26013–26. doi:10.1074/jbc.M113.489468
74. Fraser DA, Arora M, Bohlson SS, Lozano E, Tenner AJ. Generation of inhibitory
NFkappaB complexes and phosphorylated cAMP response element-binding
protein correlates with the anti-inflammatory activity of complement protein
C1q in human monocytes. J Biol Chem (2007) 282(10):7360–7. doi:10.1074/jbc.
M605741200
75. Benoit ME, Clarke EV, Morgado P, Fraser DA, Tenner AJ. Complement protein C1q directs macrophage polarization and limits inflammasome activity during the uptake of apoptotic cells. J Immunol (2012) 188(11):5682–93.
doi:10.4049/jimmunol.1103760
76. Blanco P, Palucka AK, Pascual V, Banchereau J. Dendritic cells and cytokines
in human inflammatory and autoimmune diseases. Cytokine Growth Factor Rev
(2008) 19(1):41–52. doi:10.1016/j.cytogfr.2007.10.004
77. Lood C, Gullstrand B, Truedsson L, Olin AI, Alm GV, Ronnblom L, et al.
C1q inhibits immune complex-induced interferon-alpha production in plasmacytoid dendritic cells: a novel link between C1q deficiency and systemic
lupus erythematosus pathogenesis. Arthritis Rheum (2009) 60(10):3081–90.
doi:10.1002/art.24852
78. Santer DM, Hall BE, George TC, Tangsombatvisit S, Liu CL, Arkwright
PD, et al. C1q deficiency leads to the defective suppression of IFN-alpha in
response to nucleoprotein containing immune complexes. J Immunol (2010)
185(8):4738–49. doi:10.4049/jimmunol.1001731
79. Santer DM, Yoshio T, Minota S, Moller T, Elkon KB. Potent induction of
IFN-alpha and chemokines by autoantibodies in the cerebrospinal fluid of
patients with neuropsychiatric lupus. J Immunol (2009) 182(2):1192–201.
doi:10.4049/jimmunol.182.2.1192
80. Santer DM, Wiedeman AE, Teal TH, Ghosh P, Elkon KB. Plasmacytoid dendritic cells and C1q differentially regulate inflammatory gene induction by lupus
immune complexes. J Immunol (2012) 188(2):902–15. doi:10.4049/jimmunol.
1102797
81. Fraser DA, Tenner AJ. Innate immune proteins C1q and mannan-binding
lectin enhance clearance of atherogenic lipoproteins by human monocytes
and macrophages. J Immunol (2010) 185(7):3932–9. doi:10.4049/jimmunol.
1002080
82. Spivia WR, Magno P, Le P, Fraser DA. Complement protein C1q promotes
macrophage anti-inflammatory M2-like polarization during the clearance of
atherogenic lipoproteins. Inflamm Res (2014). doi:10.1007/s00011-014-0762-0

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 26 June 2014; accepted: 07 August 2014; published online: 21 August 2014.
Citation: Bohlson SS, O’Conner SD, Hulsebus HJ, Ho M-M and Fraser DA (2014)
Complement, C1q, and C1q-related molecules regulate macrophage polarization. Front.
Immunol. 5:402. doi: 10.3389/fimmu.2014.00402
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Bohlson, O’Conner, Hulsebus, Ho and Fraser. This is an open-access
article distributed under the terms of the Creative Commons Attribution License (CC
BY). The use, distribution or reproduction in other forums is permitted, provided the
original author(s) or licensor are credited and that the original publication in this
journal is cited, in accordance with accepted academic practice. No use, distribution or
reproduction is permitted which does not comply with these terms.

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REVIEW ARTICLE
published: 27 November 2014
doi: 10.3389/fimmu.2014.00580

Purinergic and calcium signaling in macrophage function
and plasticity
Bimal N. Desai * and Norbert Leitinger *
Department of Pharmacology, University of Virginia, Charlottesville, VA, USA

Edited by:
Klaus Ley, La Jolla Institute for Allergy
and Immunology, USA
Reviewed by:
Geraldine Olivia Canny, Ecole
Polytechnique Fédérale de Lausanne,
Switzerland
Caglar Cekic, Bilkent University,
Turkey
*Correspondence:
Bimal N. Desai and Norbert Leitinger ,
Department of Pharmacology,
University of Virginia, Charlottesville,
VA, USA
e-mail: bdesai@virginia.edu;
nl2q@virginia.edu

In addition to a fundamental role in cellular bioenergetics, the purine nucleotide adenosine
triphosphate (ATP) plays a crucial role in the extracellular space as a signaling molecule. ATP
and its metabolites serve as ligands for a family of receptors that are collectively referred
to as purinergic receptors. These receptors were first described and characterized in the
nervous system but it soon became evident that they are expressed ubiquitously. In the
immune system, purinergic signals regulate the migration and activation of immune cells
and they may also orchestrate the resolution of inflammation (1, 2). The intracellular signal
transduction initiated by purinergic receptors is strongly coupled to Ca2+ -signaling, and
co-ordination of these pathways plays a critical role in innate immunity. In this review, we
provide an overview of purinergic and Ca2+ -signaling in the context of macrophage phenotypic polarization and discuss the implications on macrophage function in physiological
and pathological conditions.
Keywords: macrophages, calcium, purinergic receptors, inflammation, inflammasome activation

PURINERGIC RECEPTORS IN MACROPHAGES
Purinergic receptors are divided into P1 and P2 receptors. The
adenosine receptors are referred to as P1 receptors. P2 receptors
are the receptors for adenosine triphosphate (ATP) and can be
further subdivided into metabotropic P2Y receptors, which are
G-protein-coupled receptors and ionotropic P2X receptors, which
are cation-selective ion channels. Macrophages express a wide variety of P2X and P2Y receptors; analysis of mouse macrophages
using a variety of techniques indicates the presence of P2X4 , P2X7 ,
P2Y1 , P2Y2 , P2Y4 , and P2Y6 receptors (3). Additionally, the expression of P2Y12 receptor has been reported in microglia (4) and
macrophages (5). Historically, macrophages were also thought
to express an additional ATP-sensitive large conductance channel
termed P2Z receptor (6). However, this receptor was later shown
to be P2X7 by Surprenant et al. (7).
RELEASE OF ATP AND OTHER NUCLEOTIDES INTO THE EXTRACELLULAR
SPACE

Cytosolic ATP can be secreted through the release of ATP-loaded
vesicles or through the activation of large conductance channels (8). A variety of inflammation-related biological processes
result in ATP release from cells, and macrophages respond to this
extracellular ATP rapidly. Elegant studies by Ravichandran and
colleagues have shown that cells undergoing apoptosis release ATP
as a find-me signal that attracts phagocytes (9). In the case of dying
cells, the release of ATP and other nucleotides is accomplished
through Pannexin 1, a hemi-channel that is activated through
caspase-dependent cleavage (10). Interestingly, monocytes stimulated with pathogen-associated ligands or danger molecules, such
as uric acid can secrete ATP, which may execute an autocrine signal that results in the activation of inflammasomes and secretion
of IL-1β and IL-18 (11). Moreover, the activation of the complement cascade has also been shown to elicit ATP efflux from

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macrophages and subsequent autocrine activation of the NLRP3
inflammasome (12).
THE FUNCTION OF P2Y RECEPTORS IN MACROPHAGES

In 1989, Dubyak and colleagues showed that treatment of
macrophages with extracellular ATP elicits elevations in intracellular Ca2+ in a wide variety of myeloid cells but not in lymphocytes. These Ca2+ -elevations correlated with the hydrolysis
of inositol phospholipids suggesting that the ATP receptors were
Gq-coupled (13). Over the course of the last two decades, it has
become clear that the P2Y receptors on macrophages are Gq and
Gi/o-coupled and that they perform a critical function in ATPresponsive chemotaxis. For instance, the chemotaxis of cultured
microglia in response to extracellular ATP was shown to be dependent on Gi/o-coupled P2Y receptors by Honda et al. (14). This
receptor was later identified as P2Y12 . Microglia deficient in P2Y12
fail to polarize and migrate toward an ATP source in vitro and is
unable to extend their processes toward sites of brain damage in
mice (4). On a related note, microglial phagocytosis is triggered
by UDP that is released by damaged neurons and is dependent on
P2Y6 receptors (15). In monocytes and macrophages, P2Y2 plays
a crucial chemotactic role in locating apoptotic cells releasing ATP
(9). This study used a murine air-pouch model to demonstrate
that cell supernatants from apoptotic cells were able to recruit
monocytes and macrophages in vivo and that this recruitment
was diminished in mice lacking the P2Y2 receptor. In models of
lung inflammation, P2Y2 plays a prominent role in the chemotaxis of dendritic cells and eosinophils. Moreover, mice deficient
in the P2Y2 receptor show reduced airway inflammation in lung
inflammation models where ATP has been shown to accumulate
in the airways (16). Macrophages navigating in a gradient of C5a
secrete ATP and use a purinergic feedback loop that involves P2Y2 ,
P2Y12 , and P1 receptors to migrate (5). However, ATP-triggered

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ATP and macrophage function

Table 1 | Purinergic receptors and TRP channels in macrophages.
Activation

Downstream signaling

A1

Adenosine

Gi-coupled

A2A

Adenosine

Gs-coupled

Cellular function

Disease model phenotype after targeting

Augment M2 polarization (63)

Extensive tissue damage and prolonged inflammation (67)
Agonists induce alleviation of neural inflammation (70)

A2B

Adenosine

Gs/Gq-coupled (?)

Augment M2 polarization (63)

A3

Adenosine

Gi-coupled

Downregulation of inflammatory

Gene deletion exacerbates lung inflammation (71)
Increased mortality in a sepsis model (72)
Reduced arthritis using agonists (67)

cytokines (65, 66)
P2X4

?

?

?

?

P2X7

1 mM ATP

Non-selective cation

Activation of NLRP3 and

Resistance to contact allergen sensitivity (23)

flux

caspase-1
Lysosomal secretion of

Reduced GVHD (24)

cathepsins (27)

Resistant to pulmonary inflammation (25)

P2Y2

ATP (9)

?

Dendritic cell chemotaxis (9)

Reduced airway inflammation (16)

P2Y4

?

?

?

?

P2Y6

UDP (15)

Gq-coupled?

Microglial phagocytosis (15)

P2Y12

ATP (4)

Gi/o-coupled (14)

Microglial chemotaxis (4)

TRPC1

?

Ca2+ -influx (42)

Unconventional secretion (42)

TRPV2

?

Ca2+ -influx

Initiation of phagocytosis (39)

TRPM2

ROS

Ca2+ -influx (38)

TRPML1

?

Lysosomal
(42)

(39)

Ca2+ -release

Chemokine secretion (38)

Reduced neutrophil infiltration and intestinal inflammation (38)

Focal exocytosis during

Decreased bacterial clearance

phagocytosis (42)

?, Unknown.

chemotactic differences in M1 and M2 macrophages have not
been explored and the functional contribution of various P2Y
receptors in macrophage phenotypes remains uncharacterized
(Table 1).
THE FUNCTION OF P2X RECEPTORS IN MACROPHAGES

In comparison to P2Y receptors, the P2X receptors have a significantly lower affinity for ATP but their ability to respond to ATP
is influenced by the ionic conditions. In the case of P2X7 , replacement of Na+ with K+ greatly increases the responsiveness to ATP
suggesting a physiological role in damaged tissues with altered
ionic conditions (17). Activation of P2X7 by high concentrations
of ATP mediates caspase-1-dependent cell death accompanied by
the release of proinflammatory cytokines, such as IL-1β and IL-18.
This process is greatly potentiated in macrophages activated by LPS
(18). The processing of IL-1β and IL-18 by caspase-1 is followed
by their unconventional secretion with or without accompanying
pyroptosis, a caspase-1 mediated pathway of inflammatory cell
death. These mechanisms appear to be greatly potentiated by the
influx of extracellular Ca2+ through the P2X7 channels (19). Precisely how Ca2+ modulates the machinery mediating the secretion
of IL-1β is not clear. Activation of P2X7 also induces membrane
blebbing and activation of Rho-effector kinases but whether the
influx of Ca2+ is essential for these processes is also not clear. Concomitantly, P2X7 is thought to regulate inflammasome-dependent
activation of caspase-1 by mediating K+ efflux (20) and through

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the potentiation of an NFκB-driven transcriptional program (21).
In a related process, P2X7 has been shown to control the secretion
of MHC class II-containing exosomes in NLRP3-dependent but
caspase-1-independent manner (22).
Due to the crucial role played by P2X7 in the regulation of the
NLRP3 inflammasome, it has been implicated as a molecular target
in a large variety of inflammatory diseases (Table 1). Mice deficient
in P2X7 are not sensitized to contact allergens and fail to release
IL-1β in response to LPS and ATP (23). This study suggests that the
ligation of P2X7 by ATP is crucial for initiating skin inflammation.
Similarly, P2X7 expression on antigen presenting cells appears to
have a major impact on graft-versus-host disease (GVHD) (24).
How P2X7 affects antigen processing and presentation is not clear
yet. The P2X7 -deficient mice have also been shown to be highly
resistant to pulmonary inflammation induced by exposure to cigarette smoke (25). In the case of intestinal inflammation, mast cells
expressing P2X7 have been shown to play a central role in initiating the inflammatory cascade (26). In this case, it seems likely that
the influx of Ca2+ through P2X7 potentiates the degranulation of
mast cells. A similar role for P2X7 -mediated potentiation of lysosomal secretion of cathepsins has been reported in a mouse model
of arthritis (27). Recently, it was discovered that monocytes from
patients afflicted with Behcets disease, a severe auto-inflammatory
disorder, have increased expression of P2X7 (28). Recent studies have also implicated P2X7 in the modulation of adaptive
immunity through the control of antigen presentation on MHC

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Desai and Leitinger

class I molecules (29). In addition to P2X7 , macrophages also
express P2X1 channels but the functional significance is not yet
clear (30). Although M1 macrophages are more efficient at ATPinduced secretion of IL-1β (31), no significant differences in the
expression levels of P2X7 have been noted. It has been suggested
that P2X7 activation is decoupled from IL-1β regulation in M2
macrophages (31).

CALCIUM CHANNELS IN MACROPHAGES
As non-excitable cells, macrophages rely on Ca2+ -permeable channels that are not gated by voltage. In addition to P2X channels,
macrophages express the store-operated Orai channels and some
members of the transient receptor potential (TRP) channel superfamily. The regulation and function of these ion channels remains
a mystery and is an emerging topic of significance to inflammation
(Table 1).
ORAI CHANNELS IN MACROPHAGES

Historically, the elevations of intracellular Ca2+ in macrophages
were first observed in response to platelet-activating factor (PAF)
(32). In accord with the classic store-operated Ca2+ -entry, PAF
first elicited the mobilization of intracellular Ca2+ stores through
a Gq-coupled pathway. The emptying of the ER stores was then
followed by the opening of the Ca2+ -permeable channels in the
plasma membrane, the so-called CRAC channels (33). For almost
two decades, the identity of CRAC channels remained a mystery
but we now know their molecular identities as Orai channels (34).
Recent discoveries have unraveled the regulatory mechanisms of
Orai channels but their functional role in macrophage biology
remains undefined. Some observations have linked store-operated
Ca2+ response to the production of reactive oxygen species (ROS)
in macrophages but definitive work and mechanistic insights have
not been forthcoming (35). The Ca2+ -influx necessary for the
engulfment of apoptotic cells by macrophages is thought to be
mediated by Orai channels and genetic studies in Caenorhabditis elegans support this notion (36) but how these channels are
activated when macrophages encounter apoptotic cells is not clear
and the precise role of Ca2+ in the engulfment process has not
been clarified. Since P2Y receptors can be Gq-linked, the subsequent depletion of Ca2+ stores through IP3 receptors should
result in activation of Orai channels. Whether this actually occurs
and whether Ca2+ -influx through Orai channels is critical for the
cellular outputs of P2Y receptor stimulation is not yet clear.
TRP CHANNELS IN MACROPHAGES

The 28 members of TRP channel superfamily are subdivided
into TRPC (seven members), TRPV (six members), TRPM (eight
members), TRPML (three members), TRPP (three members),
and TRPA (one member) families (37). TRP channels are cationselective channels that are weakly voltage-sensitive and diversely
gated by temperature, mechanical force, electrophiles, ligands, and
internal cues, such as membrane composition and pH. Recent
reports have highlighted the potent functional impact of these
channels in macrophages. In monocytes lacking TRPM2, the
Ca2+ -influx in response to ROS is diminished and the cells are
unable to produce chemokines necessary for the recruitment of
other cells (38). In a mouse model of intestinal inflammation,

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ATP and macrophage function

ulceration and neutrophil infiltration were significantly attenuated
in mice lacking TRPM2 (38).
Transient receptor potential channels have also been shown to
play a major role in phagocytosis. Macrophages lacking TRPV2 are
deficient in the triggering of phagocytosis when they encounter
zymosan and IgG opsonized particles (39). Whether this function of TRPV2 is coupled to the influx of Ca2+ or other cations is
not entirely clear but abnormalities in cytoskeletal rearrangements
during phagocytosis were observed and the cells were also found
to be deficient in chemotaxis. Mice lacking TRPV2 respond poorly
when challenged with Listeria monocytogenes. They show increased
mortality and greater bacterial load in their organs (39). During
phagocytosis, macrophages replenish their membranes through a
process termed focal exocytosis. This process was thought to be
independent of Ca2+ (40), but for the phagocytosis of large particles, the process requires the activity of TRPML1 (41). Xu and
colleagues have shown that TRPML1 is a lysosomal channel that is
essential for the phagocytosis of large particles. Through the combined use of electrophysiology and live-cell imaging, authors show
convincingly that TRPML1 mediates the release of lysosomal Ca2+
at the site of membrane uptake during large particle phagocytosis.
Recently, TRPC1 has been shown to play a role in restraining
the unconventional secretion of IL-1β (42). Secretion of IL-1β is
greatly potentiated after the degradation of TRPC1 by caspase-11
and macrophages lacking TRPC1 show increased secretion of IL1β in response to inflammatory stimuli. The precise mechanism
through which TRPC1 regulates this unconventional secretion
machinery is not yet clear.

COUPLING OF PURINERGIC AND CALCIUM SIGNALING IN
MACROPHAGES
Extracellular ATP induces Ca2+ elevations in myeloid cells through
the activation of Gq -coupled P2Y receptors and Ca2+ -permeable
P2X channels. P2Y receptors have a higher affinity for ATP and can
elicit Ca2+ mobilization from the intracellular stores at low micromolar concentrations of extracellular ATP (13). In contrast, the
P2X channels open at millimolar concentrations of ATP and mediate the influx of extracellular Ca2+ and other cations (43). Notably,
the mobilization of Ca2+ stored in the endoplasmic reticulum is
not sufficient for the activation of caspase-1 and secretion of IL1β. When cells are stimulated with ATP in extracellular medium
that is depleted of Ca2+ , IL-1β secretion is nearly abolished (19).
These observations indicate that activation of caspase-1 requires
a sustained and more intense rise in intracellular Ca2+ , which
can be mediated by the activation of P2X7 channels but not P2Y
receptors.
An alternative explanation for differential requirements of
Ca2+ stores and Ca2+ -entry in the activation of caspase-1 involves
the efflux of K+ through the activated P2X7 channels. In this
model, a concomitant efflux of K+ is necessary for the activation of caspase-1 and the rise in intracellular Ca2+ without K+
efflux is insufficient (44). In any case, although the activation of
P2Y receptors by low concentrations of ATP is insufficient to activate caspase-1, the resulting Ca2+ oscillations have been shown
to promote the transcription of proinflammatory cytokines such
as IL-6 (45). The relative contributions of P2X and P2Y receptors
in nucleotide-induced Ca2+ -signaling have not been adequately

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Desai and Leitinger

defined but the use of knockout mice has provided useful insights
into this complex aspect of inflammation (46). In myeloid cells,
Ca2+ -dependent activation of PKC plays a pivotal role in the NFκB
pathway (47) and the cellular outputs at the site of inflammation
are thus likely to be shaped by the purinergic microenvironment.
Even in the absence of purinergic signals, Ca2+ stores can be
mobilized by Toll-like receptors through the activation of tyrosine kinases and phospholipase C (48), but the presence of ATP
in the microenvironment likely functions as a potent amplifying
mechanism for inflammatory processes.
In addition to the regulation of proinflammatory gene expression and cytokine secretion, Ca2+ -signaling plays a major role in
phagosome maturation. This link is exploited by the internalized
mycobacterium for immunoevasive block of phagosome maturation (49). Although the role of Ca2+ in phagosome maturation
is incompletely defined, it is clear that the lysosomal synaptotagmin VII, a Ca2+ -sensitive protein, is essential for the fusion of
lysosomes with phagosomes (50, 51). Ca2+ -influx is also essential
for the engulfment of apoptotic cells and a subsequent antiinflammatory response (36). Macrophage phenotypic polarization
results in significant differences in the execution of phagocytosis
and phagosome maturation but the associated differences in the
role of purinergic and Ca2+ -signals between differentially polarized macrophages remain undefined (52). Ca2+ -influx has also
been shown to be essential to maintain the leading-edge structure
in migrating macrophages. In this context, Ca2+ -influx may be
necessary for the activity of PKCα, which is preferentially localized
at the leading edge (53).

CHEMOTACTIC AND PHAGOCYTIC RESPONSES OF
MACROPHAGES TO EXTRACELLULAR ATP
Macrophages and other cells of myeloid lineage respond at three
basic levels to extracellular ATP gradients. First, they migrate
toward increasing ATP concentrations; second, they use the ATP
gradients emanating from dying cells as a “find-me” signal to
locate and phagocytose the cell corpse; and third, at high concentrations of ATP, macrophages respond by robust secretion of
proinflammatory cytokines.
Chemotactic responses to ATP were first convincingly demonstrated using cultured microglial cells. Extracellular ATP at micromolar concentrations induced pronounced membrane ruffling,
chemokinesis, and chemotaxis (14). This aspect of purinergic
response was not confined to ATP gradients emanating from a
distant site, indicating ATP acted on cells in an autocrine manner.
Indeed, migrating human neutrophils release ATP from their leading edges to amplify and steer their migration using an autocrine
feedback loop that involves multiple types of purinergic receptors
(54). In macrophages, the chemotactic response to C5a also utilizes an “autocrine purinergic loop” that involves the release of ATP
at the leading edge and activation of multiple purinergic receptors
(5). In asthmatic airway inflammation, ATP-induced chemotaxis
appears to play a critical role in eosinophil and dendritic cell
infiltration (55).
The analysis of dendritic cell responses to ATP clearly demonstrates that the chemotactic response to ATP is mechanistically
dissociated from other cellular effects of ATP such as secretion
of proinflammatory cytokines (16). However, the ATP-induced

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ATP and macrophage function

chemotactic response is intricately connected to the role of
purinergic signaling in the location and phagocytosis of apoptotic
cells. In the central nervous system, damaged neurons release UDP,
which triggers the phagocytic response in neighboring microglial
cells (15) and similar mechanisms are likely at work in other
tissue-resident macrophages. A definitive role for extracellular ATP
as a find-me signal for phagocytes was demonstrated by Elliott et al.
(9). The release of ATP and UTP is dependent on the activation
of caspases and Pannexin 1 during the early stages of apoptosis
and the concentration gradient generated by such release is highly
efficient at recruiting monocytes and macrophages. Whether different macrophage phenotypes migrate differently in response to
ATP gradients has not been explored. Phagocytosis of dying cells
that release ATP serves an anti-inflammatory role, and phagocytotic capacity is drastically inhibited in Mox macrophages that
accumulate at the sites of oxidative tissue damage (56). In this context, it would be interesting to know whether alternatively activated
M2 or M2-like macrophages show any significant specialization in
locating dying cells using the ATP gradients.

EXTRACELLULAR ATP AS A TRIGGER FOR
PROINFLAMMATORY CYTOKINE SECRETION AND
PYROPTOSIS
Sustained exposure to relatively high concentrations of ATP has
been shown to be a critical signal for the secretion of proinflammatory cytokines, such as IL-1β and IL-18. The unconventional
secretion of these cytokines is accomplished through proteolytic
processing by caspase-1. The activation of caspase-1 is regulated
by large multimeric complexes called inflammasomes and the activation of one such inflammasome, the NLRP3 inflammasome, is
highly sensitive to the presence of extracellular ATP. Early evidence
for ATP-induced maturation of IL-1β came from studies involving
apoptosis of peritoneal exudate cells when exposed to high concentrations of extracellular ATP. This form of apoptosis, which we
now refer to as pyroptosis, was accompanied by proteolytic processing and release of IL-1β (57), and was especially pronounced
in LPS-stimulated mouse peritoneal macrophages where exposure
to millimolar concentrations of ATP resulted in rapid processing
and release of IL-1β. It was further demonstrated that exposure to
high concentrations of extracellular K+ prevented the processing
and release of IL-1β (58), suggesting that depletion of intracellular
K+ was essential for ATP-induced IL-1β processing. Subsequent
studies of this phenomenon were greatly facilitated by the isolation of the human monocytic cell line THP-1, which was shown to
be highly sensitive to purinergic stimulation of IL-1β processing.
At least in human monocytes, the purinergic activation of IL-1β
processing and secretion is also accompanied by release of proteolytically activated caspase-1 and a commitment to cell death (59).
A study conducted by Dixit and colleagues tested the role
of channel-mediated ATP release by characterizing Pannexin 1deficient mice in the context of inflammasome activation (60).
Authors show that the activation of caspase-1 and secretion of
IL-1β in response to a wide variety of stimuli including ATP is
normal in Pannexin-1-deficient macrophages. In contrast, Pannexin 1-deficient thymocytes failed to recruit macrophages after
undergoing apoptosis. Overall, these studies indicate that ATP
released through Pannexin 1 is sufficient to reach extracellular

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Desai and Leitinger

concentrations that are of functional relevance to chemotaxis.
However, the activation of caspase-1 typically requires high concentrations of ATP that are unlikely to be reached when cytosolic
ATP is released through Pannexin 1. The role of vesicular release
of ATP in the autocrine activation of inflammasome has not yet
been ruled out and that may hold the key to reconcile these studies. A key component of vesicular ATP release is the vesicular
nucleotide transporter VNUT (also known as SLC17A9), which
is responsible for the accumulation of ATP into secretory vesicles (61). The human monocytic cell line THP-1 has been shown
to express VNUT, which mediates the rapid secretion of ATP in
response to LPS treatment (62). The function of vesicular ATP
secretion in mouse macrophages and its physiological significance
have not been reported yet. It is also not clear whether M1 and
M2 macrophages exhibit mechanistic and functional differences
in ATP release mechanisms.
The mechanisms through which activated caspase-1 and IL1β are secreted have remained unclear but there is evidence
for the involvement of Ca2+ -influx elicited by ATP (19). Interestingly, although high concentrations of ATP are required for
caspase-1 activation, low concentrations of ATP (10 µM) or UTP
(10 µM) are sufficient to induce oscillations in intracellular Ca2+
and increased transcription of the proinflammatory cytokine IL-6
(45). An autocrine role for ATP-induced activation of inflammasome has also been suggested in the case of primary human
monocytes that are stimulated by danger-associated uric acid.
According to this model, the activation of inflammasome is dependent upon the initial release of ATP, which then acts on purinergic
receptors in an autocrine manner. Whether such secretion results
in local ATP concentrations that are high enough to activate
the inflammasomes is not clear (11). In addition to the regulation of IL-1β and IL-18 secretion, ATP has also been shown to
regulate the secretion of lysosomal cathepsins (27). These proteases are involved in the degradation of extracellular matrix and
can result in auto-inflammatory tissue damage. Recently, ATP
was shown to potentiate the release of IFNβ in LPS-stimulated
macrophages (30).
With respect to macrophage phenotypes, a recent study provided evidence that inflammatory M1 macrophages are more
sensitive to ATP and more efficient at ATP-induced IL-1β release
when compared to M2 macrophages (31). While the physiological
significance of these findings remains to be elucidated, this study
provides the initial exploratory foray into these outstanding questions pertaining to how macrophage polarization fine-tunes the
sensitivity to the purinergic microenvironment.

THE ATP METABOLITE ADENOSINE REGULATES THE
RESOLUTION OF INFLAMMATION
Many cells express membrane-bound ectonucleotidases that convert the extracellular ATP and ADP to adenosine. A common
pathway involves the conversion of ATP and ADP to AMP by
CD39 and subsequent conversion of AMP to adenosine by CD73.
These enzymatic biochemical conversions have potent implications for the termination of inflammatory response due to the
reduction in ATP levels. More significantly, adenosine serves as
a ligand for G-protein coupled adenosine receptors or P1 receptors on myeloid cells (Table 1). The four P1 adenosine receptors

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ATP and macrophage function

(A1 , A2A , A2B , and A3 ) transmit a “calm down” signal that may
orchestrate the resolution of inflammation, a process conceptually
different from anti-inflammatory signals that restrain the initiation of inflammatory process by preventing the recruitment and
activation of immunocytes. Pertinently, extracellular adenosine
has been shown to augment the polarization of macrophages
toward the M2 phenotype (63).
Early studies showed that adenosine inhibited the secretion of
TNFα, IL-6, and IL-8 by LPS-activated human monocytes (64).
Subsequently, it was shown that the adenosine receptor A3 plays
a major role in downregulating the synthesis of proinflammatory cytokines in monocytes and macrophages in response to
extracellular adenosine (65, 66). Synthetic agonists of A3 adenosine receptor have been shown to have potent therapeutic effects
in various mouse models of rheumatoid arthritis (67). A similar role for the A2A adenosine receptor has also been reported.
Mice deficient in the A2A receptor show extensive tissue damage
and prolonged inflammation to sub-threshold doses of inflammatory stimuli in three different disease models (68). In the context
of infectious diseases and polymicrobial sepsis, A2A receptors
are required for the control of IL-10 production by alternatively
activated macrophages (69). The A2A receptors also play an antiinflammatory role in neuroinflammation. The ligation of A2A
receptors on the activated microglia has been shown to retract
their processes and initiate the resolution of inflammation in the
brain (70).
In the case of endotoxin-induced lung injury, the pharmacological inhibition or genetic deletion of A2B receptors greatly
exacerbates lung injury. In contrast, the A2B receptor agonist attenuates endotoxin-induced lung inflammation (71). The role of A2B
receptors in dampening endotoxin-induced inflammation is evident in mouse models of polymicrobial sepsis. Deletion of A2B
receptors greatly increases the mortality of mice from cecal ligation
and puncture-induced sepsis (72). The A2B -deficient mice showed
increased levels of proinflammatory cytokines and chemokines in
the serum, coincident with augmented activation of NFκB and
p38 in the spleen.
In summary, the adenosine receptors provide crucial information to monocytes and macrophages and calibrate their response
to the complex mix of purinergic stimuli in the inflammatory
microenvironment (73). In this context, the ratio of ATP and
adenosine may provide the crucial cues necessary for the polarization of macrophages toward the M2 phenotype induced by IL-4.
The role of A2A and A2B receptors to augment this process of M2
polarization has been illuminated (63) but surprisingly, a recent
study shows that adenosine may control this process independent
of IL-4 (74). Similarly, a recent study provides evidence suggesting
a role for adenosine in IL-10-induced STAT3 activation in alternatively activated phenotype termed M2c (75). Manipulation of
ATP and adenosine levels in the tissue microenvironment is thus
likely to emerge as a potent mechanism to guide the plasticity of
macrophages and holds clinical potential for therapeutic intervention. This concept may find traction in a large variety of diseases
where inflammation plays a major pathological role. A recent preclinical study exemplifies the application of this strategy in the
treatment of osteolysis; the authors show that activation of A2A
receptors prevents wear-induced osteolysis (76).

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EXTRACELLULAR ATP PLAYS A PROMINENT ROLE IN
INFLAMMATORY DISEASES
Some of the earliest evidence indicating a role for extracellular ATP in monocyte and macrophage function came from the
findings that the cells of the myeloid lineage exhibit rapid elevations in intracellular Ca2+ when treated with micromolar amounts
of ATP (13). The in vivo significance of this finding was further
highlighted by the studies of Bertics and colleagues who showed
that LPS-induced activation of macrophages was greatly enhanced
by extracellular ATP (77) and that mice treated with the adenine
nucleotide analog 2-methylthio-ATP were protected from endotoxic shock (78). Subsequent studies indicated a potent role for
purinergic signals in controlling the inflammatory gene expression
in response to LPS stimulation (79). The extensive contribution
of purinergic signaling in inflammatory processes has now been
established in a wide variety of pathologies (2), some of which are
outlined below.
The tissue-resident macrophages of the central nervous system, the microglia, have been shown to be especially sensitive to
extracellular ATP. In vitro studies with cultured microglia revealed
that extracellular ATP and ADP stimulate chemotaxis and morphological changes (14). Local trauma in the brain, which results
in cell death, is thought to increase the extracellular levels of
ATP significantly and elegant studies using multiphoton imaging
have demonstrated that microglia respond rapidly to local injury
through dynamic changes in their morphology. The convergence
of microglial processes at the injury site could also be stimulated
by local injection of ATP and this response was demonstrated to
be highly sensitive to the presence of ATP-hydrolyzing enzymes
and blockers of purinergic receptors (80).
In the lungs of asthmatic patients, allergic challenges cause
rapid accumulation of ATP and this has been modeled successfully in mice using experimentally induced asthma (55). Interestingly, hydrolysis of ATP in the airways through the application
of apyrase greatly reduces eosinophil infiltration, production of
Th2 cytokines, and bronchial hyper-reactivity. Corollary experiments show that exogenous ATP potentiates airway inflammation
(55) and similar findings have demonstrated a role for purinergic
signals in cigarette smoke-induced inflammation and emphysema
(25). The significance of purinergic signals in allergic reactions
is also evident in the skin. The accumulation of ATP in response
to contact allergans is critical for the production of inflammatory
cytokines by the myeloid cells and the subsequent sensitization
process (23). Accumulation of ATP has also been observed in
the ascites of patients and mice undergoing GVHD. In mice, the
severity of GVHD is greatly reduced by neutralizing the ATP
or by blocking purinergic signaling (24). Furthermore, recent
studies have also implicated extracellular ATP in the development of intestinal inflammation in patients with Crohn’s disease.
In corresponding mouse models, blocking purinergic signaling
greatly reduces the activation of intestinal mast cells and thereby
blocks the subsequent rise in proinflammatory cytokines and
leukotrienes (26).
In summary, studies in human beings and mice provide conclusive evidence that purinergic signals play a major role in inflammation and tissue injury. Purinergic receptors are expressed ubiquitously and a comprehensive understanding of how purinergic

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ATP and macrophage function

signals influence the physiology and pathology is still in rudimentary stages. Purinergic control of macrophage function promises
to play a central role in these processes and understanding the
effects of purinergic signals on macrophage function provides an
immediate window toward therapeutic intervention.

CONCLUDING REMARKS
Our understanding of the fundamental role played by purinergic
and Ca2+ signaling in macrophage activity is increasing rapidly but
the signaling mechanisms that drive specific cellular outputs still
remain largely enigmatic. The close coupling of purinergic stimulation and Ca2+ influx suggests that the purinergic receptors, Orai
channels, and TRP channels function in a co-ordinated network
that responds rapidly to the changes in the inflammatory microenvironment. By virtue of being excellent drug targets, purinergic
G-protein coupled receptors and ion channels offer an enticing pharmacological path to shape the plasticity of macrophage
function in various diseases. To make this a reality, we will
need to develop experimental models where the influence of the
purinergic microenvironment and the resulting Ca2+ -dynamics
in macrophages can be interrogated in situ. All indications are that
we have only just scratched the surface in this exciting area of
innate immunity.

ACKNOWLEDGMENTS
This work was supported by NIH-P01HL120840.

REFERENCES
1. Junger WG. Immune cell regulation by autocrine purinergic signalling. Nat Rev
Immunol (2011) 11(3):201–12. doi:10.1038/nri2938
2. Idzko M, Ferrari D, Eltzschig HK. Nucleotide signalling during inflammation.
Nature (2014) 509(7500):310–7. doi:10.1038/nature13085
3. Coutinho-Silva R, Ojcius DM, Gorecki DC, Persechini PM, Bisaggio RC,
Mendes AN, et al. Multiple P2X and P2Y receptor subtypes in mouse J774,
spleen and peritoneal macrophages. Biochem Pharmacol (2005) 69(4):641–55.
doi:10.1016/j.bcp.2004.11.012
4. Haynes SE, Hollopeter G, Yang G, Kurpius D, Dailey ME, Gan WB, et al. The
P2Y12 receptor regulates microglial activation by extracellular nucleotides. Nat
Neurosci (2006) 9(12):1512–9. doi:10.1038/nn1805
5. Kronlage M, Song J, Sorokin L, Isfort K, Schwerdtle T, Leipziger J, et al. Autocrine
purinergic receptor signaling is essential for macrophage chemotaxis. Sci Signal
(2010) 3(132):ra55. doi:10.1126/scisignal.2000588
6. Falzoni S, Munerati M, Ferrari D, Spisani S, Moretti S, Di Virgilio F. The purinergic P2Z receptor of human macrophage cells. Characterization and possible
physiological role. J Clin Invest (1995) 95(3):1207–16. doi:10.1172/JCI117770
7. Surprenant A, Rassendren F, Kawashima E, North RA, Buell G. The cytolytic
P2Z receptor for extracellular ATP identified as a P2X receptor (P2X7). Science
(1996) 272(5262):735–8. doi:10.1126/science.272.5262.735
8. Lazarowski ER. Vesicular and conductive mechanisms of nucleotide release.
Purinergic Signal (2012) 8(3):359–73. doi:10.1007/s11302-012-9304-9
9. Elliott MR, Chekeni FB, Trampont PC, Lazarowski ER, Kadl A, Walk SF, et al.
Nucleotides released by apoptotic cells act as a find-me signal to promote phagocytic clearance. Nature (2009) 461(7261):282–6. doi:10.1038/nature08296
10. Chekeni FB, Elliott MR, Sandilos JK, Walk SF, Kinchen JM, Lazarowski ER, et al.
Pannexin 1 channels mediate ‘find-me’ signal release and membrane permeability during apoptosis. Nature (2010) 467(7317):863–7. doi:10.1038/nature09413
11. Piccini A, Carta S, Tassi S, Lasiglie D, Fossati G, Rubartelli A. ATP is released by
monocytes stimulated with pathogen-sensing receptor ligands and induces IL1beta and IL-18 secretion in an autocrine way. Proc Natl Acad Sci U S A (2008)
105(23):8067–72. doi:10.1073/pnas.0709684105
12. Asgari E, Le Friec G, Yamamoto H, Perucha E, Sacks SS, Kohl J, et al. C3a modulates IL-1beta secretion in human monocytes by regulating ATP efflux and
subsequent NLRP3 inflammasome activation. Blood (2013) 122(20):3473–81.
doi:10.1182/blood-2013-05-502229

November 2014 | Volume 5 | Article 580 | 267

Desai and Leitinger

13. Cowen DS, Lazarus HM, Shurin SB, Stoll SE, Dubyak GR. Extracellular
adenosine triphosphate activates calcium mobilization in human phagocytic
leukocytes and neutrophil/monocyte progenitor cells. J Clin Invest (1989)
83(5):1651–60. doi:10.1172/JCI114064
14. Honda S, Sasaki Y, Ohsawa K, Imai Y, Nakamura Y, Inoue K, et al. Extracellular
ATP or ADP induce chemotaxis of cultured microglia through Gi/o-coupled
P2Y receptors. J Neurosci (2001) 21(6):1975–82.
15. Koizumi S, Shigemoto-Mogami Y, Nasu-Tada K, Shinozaki Y, Ohsawa K, Tsuda
M, et al. UDP acting at P2Y6 receptors is a mediator of microglial phagocytosis.
Nature (2007) 446(7139):1091–5. doi:10.1038/nature05704
16. Muller T, Robaye B, Vieira RP, Ferrari D, Grimm M, Jakob T, et al. The
purinergic receptor P2Y2 receptor mediates chemotaxis of dendritic cells and
eosinophils in allergic lung inflammation. Allergy (2010) 65(12):1545–53.
doi:10.1111/j.1398-9995.2010.02426.x
17. Gudipaty L, Humphreys BD, Buell G, Dubyak GR. Regulation of P2X(7)
nucleotide receptor function in human monocytes by extracellular ions and
receptor density. Am J Physiol Cell Physiol (2001) 280(4):C943–53.
18. Le Feuvre RA, Brough D, Iwakura Y, Takeda K, Rothwell NJ. Priming of
macrophages with lipopolysaccharide potentiates P2X7-mediated cell death via
a caspase-1-dependent mechanism, independently of cytokine production. J
Biol Chem (2002) 277(5):3210–8. doi:10.1074/jbc.M104388200
19. Gudipaty L, Munetz J, Verhoef PA, Dubyak GR. Essential role for Ca2+
in regulation of IL-1beta secretion by P2X7 nucleotide receptor in monocytes, macrophages, and HEK-293 cells. Am J Physiol Cell Physiol (2003)
285(2):C286–99. doi:10.1152/ajpcell.00070.2003
20. Kahlenberg JM, Dubyak GR. Mechanisms of caspase-1 activation by P2X7
receptor-mediated K+ release. Am J Physiol Cell Physiol (2004) 286(5):C1100–8.
doi:10.1152/ajpcell.00494.2003
21. Kahlenberg JM, Lundberg KC, Kertesy SB, Qu Y, Dubyak GR. Potentiation
of caspase-1 activation by the P2X7 receptor is dependent on TLR signals and requires NF-kappaB-driven protein synthesis. J Immunol (2005)
175(11):7611–22. doi:10.4049/jimmunol.175.11.7611
22. Qu Y, Ramachandra L, Mohr S, Franchi L, Harding CV, Nunez G, et al. P2X7
receptor-stimulated secretion of MHC class II-containing exosomes requires the
ASC/NLRP3 inflammasome but is independent of caspase-1. J Immunol (2009)
182(8):5052–62. doi:10.4049/jimmunol.0802968
23. Weber FC, Esser PR, Muller T, Ganesan J, Pellegatti P, Simon MM, et al. Lack of
the purinergic receptor P2X(7) results in resistance to contact hypersensitivity.
J Exp Med (2010) 207(12):2609–19. doi:10.1084/jem.20092489
24. Wilhelm K, Ganesan J, Muller T, Durr C, Grimm M, Beilhack A, et al. Graftversus-host disease is enhanced by extracellular ATP activating P2X7R. Nat Med
(2010) 16(12):1434–8. doi:10.1038/nm.2242
25. Lucattelli M, Cicko S, Muller T, Lommatzsch M, De Cunto G, Cardini S,
et al. P2X7 receptor signaling in the pathogenesis of smoke-induced lung
inflammation and emphysema. Am J Respir Cell Mol Biol (2011) 44(3):423–9.
doi:10.1165/rcmb.2010-0038OC
26. Kurashima Y, Amiya T, Nochi T, Fujisawa K, Haraguchi T, Iba H, et al. Extracellular ATP mediates mast cell-dependent intestinal inflammation through P2X7
purinoceptors. Nat Commun (2012) 3:1034. doi:10.1038/ncomms2023
27. Lopez-Castejon G, Theaker J, Pelegrin P, Clifton AD, Braddock M, Surprenant A.
P2X(7) receptor-mediated release of cathepsins from macrophages is a cytokineindependent mechanism potentially involved in joint diseases. J Immunol (2010)
185(4):2611–9. doi:10.4049/jimmunol.1000436
28. Castrichini M, Lazzerini PE, Gamberucci A, Capecchi PL, Franceschini R, Natale
M, et al. The purinergic P2x7 receptor is expressed on monocytes in Behcet’s
disease and is modulated by TNF-alpha. Eur J Immunol (2014) 44(1):227–38.
doi:10.1002/eji.201343353
29. Baroja-Mazo A, Barbera-Cremades M, Pelegrin P. P2X7 receptor activation
impairs exogenous MHC class I oligopeptides presentation in antigen presenting
cells. PLoS One (2013) 8(8):e70577. doi:10.1371/journal.pone.0070577
30. Gavala ML, Liu YP, Lenertz LY, Zeng L, Blanchette JB, Guadarrama AG, et al.
Nucleotide receptor P2RX7 stimulation enhances LPS-induced interferonbeta production in murine macrophages. J Leukoc Biol (2013) 94(4):759–68.
doi:10.1189/jlb.0712351
31. Pelegrin P, Surprenant A. Dynamics of macrophage polarization reveal new
mechanism to inhibit IL-1beta release through pyrophosphates. EMBO J (2009)
28(14):2114–27. doi:10.1038/emboj.2009.163

Frontiers in Immunology | Molecular Innate Immunity

ATP and macrophage function

32. Randriamampita C, Trautmann A. Biphasic increase in intracellular calcium induced by platelet-activating factor in macrophages. FEBS Lett (1989)
249(2):199–206. doi:10.1016/0014-5793(89)80624-6
33. Randriamampita C, Bismuth G, Trautmann A. Ca(2+)-induced Ca2+ release
amplifies the Ca2+ response elicited by inositol trisphosphate in macrophages.
Cell Regul (1991) 2(7):513–22.
34. Hogan PG, Lewis RS, Rao A. Molecular basis of calcium signaling in lymphocytes: STIM and ORAI. Annu Rev Immunol (2010) 28:491–533. doi:10.1146/
annurev.immunol.021908.132550
35. Jin SW, Zhang L, Lian QQ, Yao SL, Wu P, Zhou XY, et al. Close functional coupling between Ca2+ release-activated Ca2+ channels and reactive oxygen species
production in murine macrophages. Mediators Inflamm (2006) 2006(6):36192.
doi:10.1155/MI/2006/36192
36. Gronski MA, Kinchen JM, Juncadella IJ, Franc NC, Ravichandran KS. An
essential role for calcium flux in phagocytes for apoptotic cell engulfment
and the anti-inflammatory response. Cell Death Differ (2009) 16(10):1323–31.
doi:10.1038/cdd.2009.55
37. Nilius B, Szallasi A. Transient receptor potential channels as drug targets: from
the science of basic research to the art of medicine. Pharmacol Rev (2014)
66(3):676–814. doi:10.1124/pr.113.008268
38. Yamamoto S, Shimizu S, Kiyonaka S, Takahashi N, Wajima T, Hara Y,
et al. TRPM2-mediated Ca2+influx induces chemokine production in monocytes that aggravates inflammatory neutrophil infiltration. Nat Med (2008)
14(7):738–47. doi:10.1038/nm1758
39. Link TM, Park U, Vonakis BM, Raben DM, Soloski MJ, Caterina MJ. TRPV2 has
a pivotal role in macrophage particle binding and phagocytosis. Nat Immunol
(2010) 11(3):232–9. doi:10.1038/ni.1842
40. Di A, Nelson DJ, Bindokas V, Brown ME, Libunao F, Palfrey HC. Dynamin
regulates focal exocytosis in phagocytosing macrophages. Mol Biol Cell (2003)
14(5):2016–28. doi:10.1091/mbc.E02-09-0626
41. Samie M, Wang X, Zhang X, Goschka A, Li X, Cheng X, et al. A TRP channel in
the lysosome regulates large particle phagocytosis via focal exocytosis. Dev Cell
(2013) 26(5):511–24. doi:10.1016/j.devcel.2013.08.003
42. Py BF, Jin M, Desai BN, Penumaka A, Zhu H, Kober M, et al. Caspase-11 controls interleukin-1beta release through degradation of TRPC1. Cell Rep (2014)
6(6):1122–8. doi:10.1016/j.celrep.2014.02.015
43. Alonso-Torre SR, Trautmann A. Calcium responses elicited by nucleotides in
macrophages. Interaction between two receptor subtypes. J Biol Chem (1993)
268(25):18640–7.
44. Brough D, Le Feuvre RA, Wheeler RD, Solovyova N, Hilfiker S, Rothwell NJ, et al.
Ca2+ stores and Ca2+ entry differentially contribute to the release of IL-1 beta
and IL-1 alpha from murine macrophages. J Immunol (2003) 170(6):3029–36.
doi:10.4049/jimmunol.170.6.3029
45. Hanley PJ, Musset B, Renigunta V, Limberg SH, Dalpke AH, Sus R, et al. Extracellular ATP induces oscillations of intracellular Ca2+ and membrane potential
and promotes transcription of IL-6 in macrophages. Proc Natl Acad Sci U S A
(2004) 101(25):9479–84. doi:10.1073/pnas.0400733101
46. del Rey A, Renigunta V, Dalpke AH, Leipziger J, Matos JE, Robaye B, et al. Knockout mice reveal the contributions of P2Y and P2X receptors to nucleotideinduced Ca2+ signaling in macrophages. J Biol Chem (2006) 281(46):35147–55.
doi:10.1074/jbc.M607713200
47. Zhou X, Yang W, Li J. Ca2+- and protein kinase C-dependent signaling
pathway for nuclear factor-kappaB activation, inducible nitric-oxide synthase
expression, and tumor necrosis factor-alpha production in lipopolysaccharidestimulated rat peritoneal macrophages. J Biol Chem (2006) 281(42):31337–47.
doi:10.1074/jbc.M602739200
48. Chun J, Prince A. Activation of Ca2+-dependent signaling by TLR2. J Immunol
(2006) 177(2):1330–7. doi:10.4049/jimmunol.177.2.1330
49. Malik ZA, Thompson CR, Hashimi S, Porter B, Iyer SS, Kusner DJ. Cutting edge:
Mycobacterium tuberculosis blocks Ca2+ signaling and phagosome maturation
in human macrophages via specific inhibition of sphingosine kinase. J Immunol
(2003) 170(6):2811–5. doi:10.4049/jimmunol.170.6.2811
50. Roy D, Liston DR, Idone VJ, Di A, Nelson DJ, Pujol C, et al. A process for controlling intracellular bacterial infections induced by membrane injury. Science
(2004) 304(5676):1515–8. doi:10.1126/science.1098371
51. Czibener C, Sherer NM, Becker SM, Pypaert M, Hui E, Chapman ER, et al.
Ca2+ and synaptotagmin VII-dependent delivery of lysosomal membrane

November 2014 | Volume 5 | Article 580 | 268

Desai and Leitinger

52.
53.

54.

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

to nascent phagosomes. J Cell Biol (2006) 174(7):997–1007. doi:10.1083/jcb.
200605004
Canton J. Phagosome maturation in polarized macrophages. J Leukoc Biol (2014)
96(5):729–38. doi:10.1189/jlb.1MR0114-021R
Evans JH, Falke JJ. Ca2+ influx is an essential component of the
positive-feedback loop that maintains leading-edge structure and activity in
macrophages. Proc Natl Acad Sci U S A (2007) 104(41):16176–81. doi:10.1073/
pnas.0707719104
Chen Y, Corriden R, Inoue Y, Yip L, Hashiguchi N, Zinkernagel A, et al. ATP
release guides neutrophil chemotaxis via P2Y2 and A3 receptors. Science (2006)
314(5806):1792–5. doi:10.1126/science.1132559
Idzko M, Hammad H, van Nimwegen M, Kool M, Willart MA, Muskens F,
et al. Extracellular ATP triggers and maintains asthmatic airway inflammation
by activating dendritic cells. Nat Med (2007) 13(8):913–9. doi:10.1038/nm1617
Kadl A, Meher AK, Sharma PR, Lee MY, Doran AC, Johnstone SR, et al. Identification of a novel macrophage phenotype that develops in response to atherogenic phospholipids via Nrf2. Circ Res (2010) 107(6):737–46. doi:10.1161/
CIRCRESAHA.109.215715
Hogquist KA, Nett MA, Unanue ER, Chaplin DD. Interleukin 1 is processed
and released during apoptosis. Proc Natl Acad Sci U S A (1991) 88(19):8485–9.
doi:10.1073/pnas.88.19.8485
Perregaux D, Gabel CA. Interleukin-1 beta maturation and release in response
to ATP and nigericin. Evidence that potassium depletion mediated by these
agents is a necessary and common feature of their activity. J Biol Chem (1994)
269(21):15195–203.
Laliberte RE, Eggler J, Gabel CA. ATP treatment of human monocytes
promotes caspase-1 maturation and externalization. J Biol Chem (1999)
274(52):36944–51. doi:10.1074/jbc.274.52.36944
Qu Y, Misaghi S, Newton K, Gilmour LL, Louie S, Cupp JE, et al. Pannexin-1 is
required for ATP release during apoptosis but not for inflammasome activation.
J Immunol (2011) 186(11):6553–61. doi:10.4049/jimmunol.1100478
Sawada K, Echigo N, Juge N, Miyaji T, Otsuka M, Omote H, et al. Identification of a vesicular nucleotide transporter. Proc Natl Acad Sci U S A (2008)
105(15):5683–6. doi:10.1073/pnas.0800141105
Sakaki H, Tsukimoto M, Harada H, Moriyama Y, Kojima S. Autocrine regulation of macrophage activation via exocytosis of ATP and activation of P2Y11
receptor. PLoS One (2013) 8(4):e59778. doi:10.1371/journal.pone.0059778
Csoka B, Selmeczy Z, Koscso B, Nemeth ZH, Pacher P, Murray PJ, et al. Adenosine promotes alternative macrophage activation via A2A and A2B receptors.
FASEB J (2012) 26(1):376–86. doi:10.1096/fj.11-190934
Bouma MG, Stad RK, van den Wildenberg FA, Buurman WA. Differential regulatory effects of adenosine on cytokine release by activated human monocytes.
J Immunol (1994) 153(9):4159–68.
Sajjadi FG, Takabayashi K, Foster AC, Domingo RC, Firestein GS. Inhibition of
TNF-alpha expression by adenosine: role of A3 adenosine receptors. J Immunol
(1996) 156(9):3435–42.
McWhinney CD, Dudley MW, Bowlin TL, Peet NP, Schook L, Bradshaw M, et al.
Activation of adenosine A3 receptors on macrophages inhibits tumor necrosis factor-alpha. Eur J Pharmacol (1996) 310(2–3):209–16. doi:10.1016/00142999(96)00272-5
Baharav E, Bar-Yehuda S, Madi L, Silberman D, Rath-Wolfson L, Halpren M,
et al. Antiinflammatory effect of A3 adenosine receptor agonists in murine
autoimmune arthritis models. J Rheumatol (2005) 32(3):469–76.
Ohta A, Sitkovsky M. Role of G-protein-coupled adenosine receptors in downregulation of inflammation and protection from tissue damage. Nature (2001)
414(6866):916–20. doi:10.1038/414916a
Csoka B, Nemeth ZH, Virag L, Gergely P, Leibovich SJ, Pacher P, et al. A2A
adenosine receptors and C/EBPbeta are crucially required for IL-10 production

www.frontiersin.org

ATP and macrophage function

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

by macrophages exposed to Escherichia coli. Blood (2007) 110(7):2685–95.
doi:10.1182/blood-2007-01-065870
Orr AG, Orr AL, Li XJ, Gross RE, Traynelis SF. Adenosine A(2A) receptor mediates microglial process retraction. Nat Neurosci (2009) 12(7):872–8.
doi:10.1038/nn.2341
Schingnitz U, Hartmann K, Macmanus CF, Eckle T, Zug S, Colgan SP, et al.
Signaling through the A2B adenosine receptor dampens endotoxin-induced
acute lung injury. J Immunol (2010) 184(9):5271–9. doi:10.4049/jimmunol.
0903035
Csoka B, Nemeth ZH, Rosenberger P, Eltzschig HK, Spolarics Z, Pacher P, et al.
A2B adenosine receptors protect against sepsis-induced mortality by dampening excessive inflammation. J Immunol (2010) 185(1):542–50. doi:10.4049/
jimmunol.0901295
Hasko G, Pacher P, Deitch EA, Vizi ES. Shaping of monocyte and macrophage
function by adenosine receptors. Pharmacol Ther (2007) 113(2):264–75. doi:10.
1016/j.pharmthera.2006.08.003
Ferrante CJ, Pinhal-Enfield G, Elson G, Cronstein BN, Hasko G, Outram S, et al.
The adenosine-dependent angiogenic switch of macrophages to an M2-like phenotype is independent of interleukin-4 receptor alpha (IL-4Ralpha) signaling.
Inflammation (2013) 36(4):921–31. doi:10.1007/s10753-013-9621-3
Koscso B, Csoka B, Kokai E, Nemeth ZH, Pacher P, Virag L, et al. Adenosine
augments IL-10-induced STAT3 signaling in M2c macrophages. J Leukoc Biol
(2013) 94(6):1309–15. doi:10.1189/jlb.0113043
Mediero A, Frenkel SR, Wilder T, He W, Mazumder A, Cronstein BN. Adenosine
A2A receptor activation prevents wear particle-induced osteolysis. Sci Transl
Med (2012) 4(135):135ra65. doi:10.1126/scitranslmed.3003393
Tanke T, van de Loo JW, Rhim H, Leventhal PS, Proctor RA, Bertics PJ. Bacterial lipopolysaccharide-stimulated GTPase activity in RAW 264.7 macrophage
membranes. Biochem J (1991) 277(Pt 2):379–85.
Proctor RA, Denlinger LC, Leventhal PS, Daugherty SK, van de Loo JW,
Tanke T, et al. Protection of mice from endotoxic death by 2-methylthioATP. Proc Natl Acad Sci U S A (1994) 91(13):6017–20. doi:10.1073/pnas.91.13.
6017
Hu Y, Fisette PL, Denlinger LC, Guadarrama AG, Sommer JA, Proctor RA, et al.
Purinergic receptor modulation of lipopolysaccharide signaling and inducible
nitric-oxide synthase expression in RAW 264.7 macrophages. J Biol Chem (1998)
273(42):27170–5. doi:10.1074/jbc.273.42.27170
Davalos D, Grutzendler J, Yang G, Kim JV, Zuo Y, Jung S, et al. ATP mediates
rapid microglial response to local brain injury in vivo. Nat Neurosci (2005)
8(6):752–8. doi:10.1038/nn1472

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 15 August 2014; accepted: 29 October 2014; published online: 27 November
2014.
Citation: Desai BN and Leitinger N (2014) Purinergic and calcium signaling in macrophage function and plasticity. Front. Immunol. 5:580. doi:
10.3389/fimmu.2014.00580
This article was submitted to Molecular Innate Immunity, a section of the journal
Frontiers in Immunology.
Copyright © 2014 Desai and Leitinger. This is an open-access article distributed under
the terms of the Creative Commons Attribution License (CC BY). The use, distribution
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are credited and that the original publication in this journal is cited, in accordance with
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does not comply with these terms.

November 2014 | Volume 5 | Article 580 | 269

PERSPECTIVE ARTICLE
published: 09 October 2014
doi: 10.3389/fimmu.2014.00478

Nitric oxide synthase: non-canonical expression patterns
Joshua T. Mattila 1 * and Anita C. Thomas 2
1
2

Department of Microbiology and Molecular Genetics, University of Pittsburgh, Pittsburgh, PA, USA
Bristol Heart Institute and Bristol CardioVascular, Bristol Royal Infirmary, University of Bristol, Bristol, UK

Edited by:
Charles Dudley Mills, BioMedical
Consultants, USA
Reviewed by:
Gaurav K. Gupta, Harvard Medical
School, USA
Bo Shen, University of Illinois at
Chicago, USA
*Correspondence:
Joshua T. Mattila, Department of
Microbiology and Molecular Genetics,
University of Pittsburgh, 5065
Biomedical Science Tower 3, 3501
Fifth Avenue, Pittsburgh, PA 15260,
USA
e-mail: jmattila@pitt.edu

Science can move ahead by questioning established or canonical views and, so it may be
with the enzymes, nitric oxide synthases (NOS). Nitric oxide (NO) is generated by NOS isoforms that are often described by their tissue-specific expression patterns. NOS1 (nNOS)
is abundant in neural tissue, NOS2 is upregulated in activated macrophages and known as
inducible NOS (iNOS), and NOS3 (eNOS) is abundant in endothelium where it regulates vascular tone.These isoforms are described as constitutive or inducible, but in this perspective
we question the broad application of these labels. Are there instances where “constitutive” NOS (NOS1 and NOS3) are inducibly expressed; conversely, are there instances
where NOS2 is constitutively expressed? NOS1 and NOS3 inducibility may be linked to
post-translational regulation, making their actual patterns activity much more difficult to
detect. Constitutive NOS2 expression has been observed in several tissues, especially the
human pulmonary epithelium where it may regulate airway tone. These data suggest that
expression of the three NOS enzymes may include non-established patterns. Such information should be useful in designing strategies to modulate these important enzymes in
different disease states.
Keywords: NOS1, NOS2, NOS3, iNOS, eNOS, nNOS, nitric oxide, nitric oxide synthase

INTRODUCTION
Nitric oxide synthases (NOS) are enzymes that catalyze the conversion of l-arginine to l-citrulline and nitric oxide (NO), a free
radical involved in homeostatic and immunological functions.
There are three NOS isoforms and each isoform is associated with
a set of characteristics and expression pattern. These expression
patterns have been used to define the isoform’s nomenclature.
NOS1 is often called nNOS because of its expression in neurons and the brain. NOS2 is referred to as iNOS, because its
expression can be induced by cellular activation. NOS3 is often
referred to as eNOS because of its association with the endothelium. The purpose of this Perspective is to examine the concept
of inducible and constitutive NOS expression, and suggest that
although the current paradigm is supported in many instances,
the constitutive versus inducible dichotomy has been applied too
broadly and may restrict our understanding of these enzymes’
functions in health and disease. A complete examination of potential NO-mediated physiological functions and NOS-expressing
cells throughout an organism’s tissues is beyond the scope of this
work. Our focus will be on immunologically relevant cells (e.g.,
lymphocytes, macrophages, and the epithelium), but we will also
include some non-typical NOS-expressing cells (osteoclasts and
cancer). Moreover, to avoid any confusion associated with the
tissue-origin nomenclature, we will identify each NOS isoform
by its numeric descriptor (e.g., NOS1, NOS2, NOS3).

BASIC NOS BIOCHEMISTRY
All three NOS enzymes are catalytically active when dimerized
and require two substrates, l-arginine, molecular oxygen, in combination with several co-factors including nicotinamide-adeninedinucleotide phosphate (NADPH), flavin adenine dinucleotide

Frontiers in Immunology | Inflammation

(FAD), flavin mononucleotide (FMN), and (6R)5,6,7,8tetrahydro-l-biopterin (BH4) to generate NO (1). Two NOS
isoforms, NOS1 and NOS3, are commonly associated with constitutive expression. NOS1 and NOS3 activity is calcium dependent and requires interaction between the NOS enzyme and
calmodulin-bound calcium to facilitate the catalysis of l-arginine
and production of NO. In addition to the required co-factors
and enzyme substrates, NOS1 and NOS3 are regulated through
a variety of post-translational mechanisms including phosphorylation, myristoylation, and palmitoylation, and modification of
subcellular localization (2, 3). NOS1 and NOS3 are commonly
associated with the “low” levels of NO production that mediate
intracellular signaling processes (NOS1) and vascular homeostasis (NOS3). In addition to NO production, NOS3 can function
in an “uncoupled” manner and produce ROS when the available stores of BH4 are removed or oxidized, l-arginine depleted,
or the NOS3 inhibitor asymmetric dimethyl-l-arginine overexpressed (1, 4). NOS1 and NOS3 are most commonly found in
non-immunological cells (e.g., neurons, muscle, endothelium),
and, because their NO output is relatively low, these isoforms
are considered to be less immunologically important than their
inducible, immunologically relevant counterpart, NOS2.
Inducible expression of NOS has long been associated with
immunological functions. Immune cells use NO, often in conjunction with reactive oxygen intermediates (ROI), to kill pathogens
and cancer cells (5, 6). NO acts non-specifically on a variety of targets and can kill targets at micromolar concentrations (7). This
lack of specificity can cause collateral damage to normal cells
and tissues and consequently, NO production is tightly regulated.
NOS2 is minimally expressed or is not abundant intracellularly
in macrophages unless immune-related stimulation and gene

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NOS expression varies by situation

transcription occurs (hence its label as the “inducible” NOS isoform). Once transcribed, NOS2 has a high-affinity binding site for
calmodulin and can function in a calcium-independent manner
suggesting that any time it is expressed it is likely to be active.
While the factors inducing and regulating NOS2 have been extensively studied in rodent models, NOS2 has been more difficult to
study in primates. There has been controversy regarding its importance in human immune responses (8), or even whether NOS2 is
expressed in human macrophages (9–11). There are several reasons why NOS2 expression has been difficult to identify in primate
macrophages, including the different signals required for induction, inappropriate culture conditions, or intrinsic differences in
NOS expression, but it is increasingly clear that NOS2 is expressed
by human macrophages and has implications for human disease
(12). A variety of immune cells other than macrophages [ranging
from memory T cells (13, 14) to chondrocytes (15)] also express
NOS2 in response to stimulation, suggesting that NOS2 expression
is more flexible and extensive than previously reported.

IS NOS EXPRESSION “TRUE TO FORM?”
While NOS1, NOS2, and NOS3 have been associated with particular cell types and expression patterns, questions remain about
whether these associations have been applied too strictly. Although
the concept of constitutive (NOS1, NOS3) vs. inducible (NOS2)
expression appears to be convenient, is it biologically plausible that
NOS1 or NOS3 expression can be inducible under some circumstances, and conversely, can NOS2 be constitutively expressed in
other circumstances? This issue has important clinical and therapeutic implications that need to be considered when designing
new immunomodulatory therapies that rely on NOS expression to
fight cancer or infectious diseases, or exploring current therapies
for unanticipated effects. The answers are complicated by inconsistent data from experiments using different cell lines, animal
models or clinical samples, and experimental techniques, but there
are likely to be some generally applicable concepts and examples

that we can use as guidelines. The remainder of this review will be
focused on identifying the evidence for inducible NOS1 and NOS3,
and constitutive NOS2 expression (summarized in Table 1).

INDUCIBLE EXPRESSION OF “CONSTITUTIVE” NOS
ISOFORMS
There are a few instances of NOS1 expression that are clearly
associated with upregulation in response to external stimuli. Classically, homeostatic NOS1 expression has been associated with
neuronal signaling, although inflammatory stimuli can increase
neuronal NOS1 expression, potentially leading to NO-mediated
damage (30, 31). The relative contribution of NOS1 to pathology in this context is often confounded by co-induction of
NOS2 expression. NOS1 splice variants are expressed in skeletal, cardiac, and smooth muscle cells and can generate NO that
increases blood vessel dilatation and improved blood flow to
nearby muscle tissue (32). The paucity of data on NOS1 expression in monocyte-derived macrophages has suggested, perhaps
incorrectly, that NOS1 has little expression or importance for
tissue macrophages. That said, NOS1 expression has been identified in human bronchoalveolar lavage cells (16), lung cancer
(33), and alveolar and epithelioid macrophages from humans
with tuberculosis (17). Although these observations do not
necessarily indicate that NOS1 is upregulated in these cells,
they demonstrate non-canonical NOS1 expression, and suggest that NOS1 may be immunologically important in unanticipated ways. Significantly, recent data indicate that NOS1
activity may be regulated post-transcriptionally, with important consequences for macrophage activation and function. In
unprimed murine bone marrow-derived macrophages, immune
complexes can stimulate calcium-dependent NOS1 and NOS3
activity that leads to increased phagocytosis by these cells (18)
indicating the upregulated activity of the “constitutive” NOS isoforms that may have unappreciated roles in immunity. There
may be other systems and cell types where post-transcriptional

Table 1 | Examples of non-canonical NOS expression in non-cancerous cells and tissues.
Isoform

Cell type and reference

Species

Pathology

Expression pattern

NOS1

Bronchial epithelial cells (16)

Human

No

Ca2+ flux-dependent induction

NOS1

Alveolar macrophages (17)

Human

Tuberculosis

Induced-immune stimulation?

NOS1

Epithelioid macrophages (17)

human

Tuberculosis

Induced-immune stimulation?

NOS1

BMDa macrophage (18)

mouse

N/Ab

Ca2+ flux-dependent induction

NOS2

Colonic epithelium (19, 20)

Human

No

Constitutive expression

NOS2

Lung epithelium (21–23)

Human, macaque

No

Constitutive expression

NOS2

Brain, spinal tissue (24, 25)

Rat

No

Constitutive expression

NOS3

Alveolar macrophages (17, 26)

Human, macaque

Tuberculosis

Induced-immune stimulation?

NOS3

Epithelioid macrophages (17)

Human, macaque

Tuberculosis

Induced-immune stimulation?

NOS3

RAW264.7 macrophagesc (27)

Mouse

N/A

Ca2+ flux-dependent induction

NOS3

BMDa macrophages (28)

Mouse

N/A

LPS-stimulated activity

NOS3

Osteoclasts (29)

human

No

Ca2+ flux-dependent induction

a

BMD, macrophages differentiated from bone marrow-derived monocytes.

b

N/A, not applicable.

c

RAW264.7 macrophages are a murine macrophage-like cell line.

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Mattila and Thomas

regulation of NOS1 expression through Ca2+ -dependent or other
modulatory mechanisms can confer inducible-like characteristics to this “constitutively” expressed isoform. However, identifying these mechanisms will require a deeper understanding
of cellular dynamics and responses in vivo, and this cannot be
obtained using immunohistochemistry or studying isolated cells
or cell lines.
There is considerable evidence indicating NOS3 expression is
inducible under the right conditions. Forstermann et al. found
that expression of NOS3 could be modulated by a range of stimuli, and that there appeared to be a species-specific difference
in NOS3 regulation (34). More recently, reports have identified
that NOS3 expression can be induced in human and macaque
macrophages (17, 26), but the significance of the presence and
inducibility of this isoform in macrophages remains to be elucidated. The macrophage-like murine cell line RAW264.7 is known
for its ability to produce significant quantities of NO via an iNOSdependent mechanism following interferon gamma and LPS stimulation, but it also constitutively expresses calcium-sensitive NOS3
and produces low levels of NO in a calcium-dependent manner (27), reminiscent of NOS1-mediated NO production (18).
The quantity of NO produced at steady state was approximately
20-fold less than that produced by NOS2 following stimulation,
suggesting that its function was not directly bactericidal. A later
study using murine bone marrow-derived macrophages identified NOS3-generated NO as an important factor in capacitating macrophage activation by enabling increased NK-κB activity,
NOS2 expression, and NO production (28). Interestingly, mice
lacking NOS3 produced less NOS2 protein and, subsequently,
less NO following immune stimulation than control mice (28).
In addition, it was observed that NOS2 induction led to diminished NOS3 expression, suggesting that there was an inverse feedback loop regulating NOS2- and NOS3-mediated NO production.
NOS3 expression has also been observed in macrophages from
non-human primates (26) and humans (17, 26) in the context of
Mycobacterium tuberculosis infection, suggesting that NOS3 may
be important in primate pulmonary immune responses. As previously mentioned, dysregulated NOS3 can produce ROS instead of
NO and it cannot be ruled out that macrophage NOS3 does not
generate non-traditional products instead of NO in these situations, particularly in an environments rich in l-arginine-utilizing
enzymes (e.g., lung and tuberculous granuloma (26, 35)). There
is also evidence that NOS3 may be important in bone remodeling and can be regulated by controlling access to Ca2+ - and
NOS2-mediated NO production (29). Unstimulated osteoclasts
(macrophage-like cells responsible for bone remodeling) constitutively express both NOS2 and NOS3, with bone resorption
associated with Ca2+ -dependent NOS3-mediated NO production and inhibition of osteoclast function mediated by NOS2
(36). As with NOS1, it may be difficult to identify upregulated
NOS3-mediated NO production in instances where this increase
is attributable to post-translational events. There is also some
evidence that NOS1 and NOS3 activity can be upregulated posttranslationally by stimuli-specific release of Ca2+ . This type of
activation has important consequences in the regulation of many
physiological processes, ranging from macrophage activation to
bone homeostasis.

Frontiers in Immunology | Inflammation

NOS expression varies by situation

CONSTITUTIVE EXPRESSION OF THE “INDUCIBLE” NOS2
ISOFORM
Nitric oxide synthase 2 has become the paradigm of an inducible
immunoresponsive gene, particularly in rodent systems. The highaffinity calmodulin-binding domain of NOS2 enables it to function in conditions where Ca2+ is unavailable, suggesting that
dimerized NOS2 is always active and capable of generating NO
when the appropriate co-factors are present (1, 37). The ease at
which NOS2 expression is induced varies across and there are significant differences in species-specific expression patterns (9, 38,
39) and even differences between individuals in genetically diverse
populations (40). In mice, which are often viewed as the paradigm
for inducible NOS2 expression, some strains have macrophages
that readily express NOS2 when stimulated, whereas other strains
have more restrained NOS2 expression (41). NOS2 expression
in primate systems appears to have different requirements for
its induction that can result in NO concentrations that differ
by several orders of magnitude (39). That said, although NOS2
expression is generally inducible, in some circumstances, NOS2
can be constitutively expressed. Some of the best-described examples of constitutive NOS2 expression occur in the human colonic
epithelium (19, 20) and pseudostratified columnar epithelia in the
human (21–23) and non-human primate lung (Figure 1). In the
lung, constitutive NOS2 expression by these cells is robust and
likely to be responsible for the majority of exhaled NO in human
breath (22). NOS2 expression from these cells is thought to help
regulate ciliary beat (16) and airway tone or reactivity (22). Rat
epithelium can also express NOS1 (21), suggesting that there are
likely to be species-specific differences in epithelial NOS expression. It should also be noted that neither the lung nor the colonic
epithelia are sterile environments, and there remains the possibility that NOS2 expression occurs in response to stimulation by
the normal microbiota associated with these tissues. Neural tissue

FIGURE 1 | NOS2 is strongly expressed by ciliated pseudostratified
columnar epithelial cells in the cynomolgus macaque lung.
Formalin-fixed paraffin-embedded lung tissue sections were stained for
(A) NOS2 (green) or (B) NOS3 (green) in combination with CD163 (red), a
hemoglobin scavenger receptor expressed on macrophages and epithelial
cells, and imaged by widefield epifluorescence microscopy. Intense NOS2
expression can be observed in the basal cells underlying the ciliated cells,
with less intense staining in the ciliated cells. NOS3 staining is associated
with cells in the lamina propria but not ciliated epithelial cells. This staining
is characteristic of ciliated epithelia of both uninfected and Mycobacterium
tuberculosis-infected macaques (pictured). DAPI-stained nuclei are
indicated in blue.

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Mattila and Thomas

is much less likely to be associated with bacteria, and there is
evidence that NOS2 in rodents is constitutively expressed at low
levels in brain and spinal tissue (24, 25). This can be upregulated
above basal levels by inflammatory stimuli where it may be associated with disease in models of pathological conditions including
Alzheimer’s disease and arthritis-associated arthralgia (30, 31).
In addition to constitutive expression in normal tissues, constitutive NOS2 expression has been identified in tumors, including melanoma (42), prostate cancer (43), colorectal cancer (44),
breast cancer (45), bladder cancer (46), head and neck cancer
(47), and esophageal adenocarcinoma (48). In these pathologies,
NOS2 is often associated with poor prognosis, potentially related
to increased angiogenesis, metastatic ability, aggressive growth,
resistance to apoptosis, and chemotherapy (49, 50). The mechanistic basis for why tumor progression is sometimes associated
with NOS2 expression is not fully understood, but could include
additional mutation by NOS-mediated DNA strand breakage,
and immunosuppression of T-cell responses through both NOdependent and NO-independent mechanisms (49). Research in
this area is not without controversy and there is evidence that
NOS2-generated NO has protective effects in cancer, possibly
reflecting differences in a tumor’s inflammatory state, the type
of infiltrating immune cells, tumor location, tumor type, and the
stage of disease, as well as differences in whether there are high
or low levels of NO in the tumor microenvironment (49, 50).
Although poorly understood at present, a better understanding
of how NOS2 expression influences the tumor environment may
lead to the development of novel interventional strategies and
improved clinical treatment (50, 51).

CONCLUDING STATEMENT
A better understanding of the properties and expression patterns
of the different NOS isoforms has shed light on the diverse range
of physiological roles that these enzymes fulfill. We now know that
there are instances where functions of these enzymes diverge from
the dichotomous constitutive or inducible expression patterns they
are often associated with. We should take this opportunity to study
the full range of possible NOS function. Recognizing the possibility that NOS enzymes may act in non-canonical ways can only
increase our understanding of how tissues respond to disease and
give us new opportunities for developing innovative therapeutic
strategies.

AUTHOR CONTRIBUTIONS
Joshua T. Mattila and Anita C. Thomas contributed to drafting the
manuscript.

ACKNOWLEDGMENTS
We gratefully acknowledge JoAnne L. Flynn (University of Pittsburgh, Pittsburgh, PA, USA) for permission to image data on
cynomolgus macaque lung. Joshua T. Mattila is supported in
part by NIH RO1A103785 (JoAnne L. Flynn) and grants from
the Bill and Melinda Gates Foundation (JoAnne L. Flynn). Anita
C. Thomas is supported by funding from the British Heart
Foundation.

REFERENCES
1. Forstermann U, Sessa WC. Nitric oxide synthases: regulation and function. Eur
Heart J (2012) 33(7):829–37. doi:10.1093/eurheartj/ehr304

www.frontiersin.org

NOS expression varies by situation

2. Musicki B, Ross AE, Champion HC, Burnett AL, Bivalacqua TJ. Posttranslational
modification of constitutive nitric oxide synthase in the penis. J Androl (2009)
30(4):352–62. doi:10.2164/jandrol.108.006999
3. Venema RC. Post-translational mechanisms of endothelial nitric oxide synthase
regulation by bradykinin. Int Immunopharmacol (2002) 2(13–14):1755–62.
doi:10.1016/S1567-5769(02)00185-6
4. Forstermann U, Munzel T. Endothelial nitric oxide synthase in vascular disease: from marvel to menace. Circulation (2006) 113(13):1708–14. doi:10.1161/
CIRCULATIONAHA.105.602532
5. MacMicking J, Xie QW, Nathan C. Nitric oxide and macrophage function. Annu
Rev Immunol (1997) 15:323–50. doi:10.1146/annurev.immunol.15.1.323
6. Xu W, Liu LZ, Loizidou M, Ahmed M, Charles IG. The role of nitric oxide in
cancer. Cell Res (2002) 12(5–6):311–20. doi:10.1038/sj.cr.7290133
7. Wink DA, Hines HB, Cheng RY, Switzer CH, Flores-Santana W, Vitek MP, et al.
Nitric oxide and redox mechanisms in the immune response. J Leukoc Biol
(2011) 89(6):873–91. doi:10.1189/jlb.1010550
8. Murray PJ, Wynn TA. Obstacles and opportunities for understanding
macrophage polarization. J Leukoc Biol (2011) 89(4):557–63. doi:10.1189/jlb.
0710409
9. Gross TJ, Kremens K, Powers LS, Brink B, Knutson T, Domann FE, et al.
Epigenetic silencing of the human NOS2 gene: rethinking the role of nitric
oxide in human macrophage inflammatory responses. J Immunol (2014)
192(5):2326–38. doi:10.4049/jimmunol.1301758
10. Schneemann M, Schoeden G. Macrophage biology and immunology: man is
not a mouse. J Leukoc Biol (2007) 81(3):579. doi:10.1189/jlb.1106702
11. Schneemann M, Schoedon G. Species differences in macrophage NO production
are important. Nat Immunol (2002) 3(2):102. doi:10.1038/ni0202-102b
12. Nathan C, Shiloh MU. Reactive oxygen and nitrogen intermediates in the relationship between mammalian hosts and microbial pathogens. Proc Natl Acad
Sci U S A (2000) 97(16):8841–8. doi:10.1073/pnas.97.16.8841
13. Obermajer N, Wong JL, Edwards RP, Chen K, Scott M, Khader S, et al. Induction and stability of human Th17 cells require endogenous NOS2 and cGMPdependent NO signaling. J Exp Med (2013) 210(7):1433–45. doi:10.1084/jem.
20121277
14. Esaki T, Hayashi T, Muto E, Yamada K, Kuzuya M, Iguchi A. Expression
of inducible nitric oxide synthase in T lymphocytes and macrophages of
cholesterol-fed rabbits. Atherosclerosis (1997) 128(1):39–46. doi:10.1016/S00219150(96)05976-X
15. Schmidt N, Pautz A, Art J, Rauschkolb P, Jung M, Erkel G, et al. Transcriptional
and post-transcriptional regulation of iNOS expression in human chondrocytes.
Biochem Pharmacol (2010) 79(5):722–32. doi:10.1016/j.bcp.2009.10.012
16. Asano K, Chee CB, Gaston B, Lilly CM, Gerard C, Drazen JM, et al. Constitutive
and inducible nitric oxide synthase gene expression, regulation, and activity in
human lung epithelial cells. Proc Natl Acad Sci U S A (1994) 91(21):10089–93.
doi:10.1073/pnas.91.21.10089
17. Choi HS, Rai PR, Chu HW, Cool C, Chan ED. Analysis of nitric oxide synthase
and nitrotyrosine expression in human pulmonary tuberculosis. Am J Respir
Crit Care Med (2002) 166(2):178–86. doi:10.1164/rccm.2201023
18. Huang Z, Hoffmann FW, Fay JD, Hashimoto AC, Chapagain ML, Kaufusi
PH, et al. Stimulation of unprimed macrophages with immune complexes
triggers a low output of nitric oxide by calcium-dependent neuronal nitricoxide synthase. J Biol Chem (2012) 287(7):4492–502. doi:10.1074/jbc.M111.
315598
19. Perner A, Andresen L, Normark M, Rask-Madsen J. Constitutive expression of
inducible nitric oxide synthase in the normal human colonic epithelium. Scand
J Gastroenterol (2002) 37(8):944–8. doi:10.1080/003655202760230919
20. Roberts PJ, Riley GP, Morgan K, Miller R, Hunter JO, Middleton SJ. The physiological expression of inducible nitric oxide synthase (iNOS) in the human colon.
J Clin Pathol (2001) 54(4):293–7. doi:10.1136/jcp.54.4.293
21. Kobzik L, Bredt DS, Lowenstein CJ, Drazen J, Gaston B, Sugarbaker D, et al.
Nitric oxide synthase in human and rat lung: immunocytochemical and histochemical localization. Am J Respir Cell Mol Biol (1993) 9(4):371–7. doi:10.1165/
ajrcmb/9.4.371
22. Lane C, Knight D, Burgess S, Franklin P, Horak F, Legg J, et al. Epithelial inducible
nitric oxide synthase activity is the major determinant of nitric oxide concentration in exhaled breath. Thorax (2004) 59(9):757–60. doi:10.1136/thx.2003.
014894
23. Guo FH, De Raeve HR, Rice TW, Stuehr DJ, Thunnissen FB, Erzurum
SC. Continuous nitric oxide synthesis by inducible nitric oxide synthase in

October 2014 | Volume 5 | Article 478 | 273

Mattila and Thomas

normal human airway epithelium in vivo. Proc Natl Acad Sci U S A (1995)
92(17):7809–13. doi:10.1073/pnas.92.17.7809
24. Tang Q, Svensson CI, Fitzsimmons B, Webb M, Yaksh TL, Hua XY. Inhibition of spinal constitutive NOS-2 by 1400W attenuates tissue injury and
inflammation-induced hyperalgesia and spinal p38 activation. Eur J Neurosci
(2007) 25(10):2964–72. doi:10.1111/j.1460-9568.2007.05576.x
25. Ruscheweyh R, Goralczyk A, Wunderbaldinger G, Schober A, Sandkuhler J. Possible sources and sites of action of the nitric oxide involved in synaptic plasticity at spinal lamina I projection neurons. Neuroscience (2006) 141(2):977–88.
doi:10.1016/j.neuroscience.2006.04.010
26. Mattila JT, Ojo OO, Kepka-Lenhart D, Marino S, Kim JH, Eum SY, et al. Microenvironments in tuberculous granulomas are delineated by distinct populations
of macrophage subsets and expression of nitric oxide synthase and arginase
isoforms. J Immunol (2013) 191(2):773–84. doi:10.4049/jimmunol.1300113
27. Schmidt HH, Warner TD, Nakane M, Forstermann U, Murad F. Regulation and
subcellular location of nitrogen oxide synthases in RAW264.7 macrophages. Mol
Pharmacol (1992) 41(4):615–24.
28. Connelly L, Jacobs AT, Palacios-Callender M, Moncada S, Hobbs AJ. Macrophage
endothelial nitric-oxide synthase autoregulates cellular activation and proinflammatory protein expression. J Biol Chem (2003) 278(29):26480–7. doi:10.
1074/jbc.M302238200
29. Evans DM, Ralston SH. Nitric oxide and bone. J Bone Miner Res (1996)
11(3):300–5. doi:10.1002/jbmr.5650110303
30. Wu J, Lin Q, Lu Y, Willis WD, Westlund KN. Changes in nitric oxide synthase
isoforms in the spinal cord of rat following induction of chronic arthritis. Exp
Brain Res (1998) 118(4):457–65. doi:10.1007/s002210050302
31. Di Girolamo G, Farina M, Riberio ML, Ogando D, Aisemberg J, de los Santos
AR, et al. Effects of cyclooxygenase inhibitor pretreatment on nitric oxide production, nNOS and iNOS expression in rat cerebellum. Br J Pharmacol (2003)
139(6):1164–70. doi:10.1038/sj.bjp.0705315
32. Mungrue IN, Bredt DS. nNOS at a glance: implications for brain and brawn.
J Cell Sci (2004) 117(Pt 13):2627–9. doi:10.1242/jcs.01187
33. Lewko B, Zoltowska A, Stepinski J, Kaminski M, Skokowski J, Roszkiewicz A,
et al. Nitric oxide synthase type 1 expression in human lung cancer and its
relation to p53. Med Sci Monit (2001) 7(2):218–21.
34. Forstermann U, Boissel JP, Kleinert H. Expressional control of the ‘constitutive’ isoforms of nitric oxide synthase (NOS I and NOS III). FASEB J (1998)
12(10):773–90.
35. Pessanha AP, Martins RA, Mattos-Guaraldi AL, Vianna A, Moreira LO. Arginase1 expression in granulomas of tuberculosis patients. FEMS Immunol Med Microbiol (2012) 66(2):265–8. doi:10.1111/j.1574-695X.2012.01012.x
36. Brandi ML, Hukkanen M, Umeda T, Moradi-Bidhendi N, Bianchi S, Gross
SS, et al. Bidirectional regulation of osteoclast function by nitric oxide synthase isoforms. Proc Natl Acad Sci U S A (1995) 92(7):2954–8. doi:10.1073/pnas.
92.7.2954
37. Lowenstein CJ, Padalko E. iNOS (NOS2) at a glance. J Cell Sci (2004)
117(Pt 14):2865–7. doi:10.1242/jcs.01166
38. Jungi TW, Adler H, Adler B, Thony M, Krampe M, Peterhans E. Inducible
nitric oxide synthase of macrophages. Present knowledge and evidence for
species-specific regulation. Vet Immunol Immunopathol (1996) 54(1–4):323–30.
doi:10.1016/S0165-2427(96)05690-5
39. Vitek MP, Brown C, Xu Q, Dawson H, Mitsuda N, Colton CA. Characterization of NO and cytokine production in immune-activated microglia and peritoneal macrophages derived from a mouse model expressing the human NOS2
gene on a mouse NOS2 knockout background. Antioxid Redox Signal (2006)
8(5–6):893–901. doi:10.1089/ars.2006.8.893

Frontiers in Immunology | Inflammation

NOS expression varies by situation

40. Weinberg JB. Nitric oxide production and nitric oxide synthase type 2 expression
by human mononuclear phagocytes: a review. Mol Med (1998) 4(9):557–91.
41. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages
and the Th1/Th2 paradigm. J Immunol (2000) 164(12):6166–73. doi:10.4049/
jimmunol.164.12.6166
42. Grimm EA, Ellerhorst J, Tang CH, Ekmekcioglu S. Constitutive intracellular
production of iNOS and NO in human melanoma: possible role in regulation of growth and resistance to apoptosis. Nitric Oxide (2008) 19(2):133–7.
doi:10.1016/j.niox.2008.04.009
43. Klotz T, Bloch W, Volberg C, Engelmann U, Addicks K. Selective expression of inducible nitric oxide synthase in human prostate carcinoma. Cancer
(1998) 82(10):1897–903. doi:10.1002/(SICI)1097-0142(19980515)82:10<1897:
:AID-CNCR12>3.0.CO;2-O
44. Cianchi F, Cortesini C, Fantappie O, Messerini L, Schiavone N, Vannacci A,
et al. Inducible nitric oxide synthase expression in human colorectal cancer: correlation with tumor angiogenesis. Am J Pathol (2003) 162(3):793–801.
doi:10.1016/S0002-9440(10)63876-X
45. Vakkala M, Kahlos K, Lakari E, Paakko P, Kinnula V, Soini Y. Inducible nitric
oxide synthase expression, apoptosis, and angiogenesis in in situ and invasive
breast carcinomas. Clin Cancer Res (2000) 6(6):2408–16.
46. Wolf H, Haeckel C, Roessner A. Inducible nitric oxide synthase expression in human urinary bladder cancer. Virchows Arch (2000) 437(6):662–6.
doi:10.1007/s004280000296
47. Brennan PA, Dennis S, Poller D, Quintero M, Puxeddu R, Thomas GJ. Inducible
nitric oxide synthase: correlation with extracapsular spread and enhancement
of tumor cell invasion in head and neck squamous cell carcinoma. Head Neck
(2008) 30(2):208–14. doi:10.1002/hed.20675
48. Wilson KT, Fu S, Ramanujam KS, Meltzer SJ. Increased expression of inducible
nitric oxide synthase and cyclooxygenase-2 in Barrett’s esophagus and associated
adenocarcinomas. Cancer Res (1998) 58(14):2929–34.
49. Lechner M, Lirk P, Rieder J. Inducible nitric oxide synthase (iNOS) in tumor
biology: the two sides of the same coin. Semin Cancer Biol (2005) 15(4):277–89.
doi:10.1016/j.semcancer.2005.04.004
50. Crowell JA, Steele VE, Sigman CC, Fay JR. Is inducible nitric oxide synthase a
target for chemoprevention? Mol Cancer Ther (2003) 2(8):815–23.
51. Fitzpatrick B, Mehibel M, Cowen RL, Stratford IJ. iNOS as a therapeutic
target for treatment of human tumors. Nitric Oxide (2008) 19(2):217–24.
doi:10.1016/j.niox.2008.05.001

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 01 August 2014; accepted: 19 September 2014; published online: 09 October
2014.
Citation: Mattila JT and Thomas AC (2014) Nitric oxide synthase: non-canonical
expression patterns. Front. Immunol. 5:478. doi: 10.3389/fimmu.2014.00478
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Mattila and Thomas. This is an open-access article distributed under
the terms of the Creative Commons Attribution License (CC BY). The use, distribution
or reproduction in other forums is permitted, provided the original author(s) or licensor
are credited and that the original publication in this journal is cited, in accordance with
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October 2014 | Volume 5 | Article 478 | 274

PERSPECTIVE ARTICLE
published: 02 September 2014
doi: 10.3389/fimmu.2014.00420

Metabolic reprograming in macrophage polarization
Silvia Galván-Peña and Luke A. J. O’Neill *
School of Biochemistry and Immunology, Trinity Biomedical Science Institute, Trinity College Dublin, Dublin, Ireland

Edited by:
Laurel L. Lenz, National Jewish
Health, USA
Reviewed by:
Kottarappat N. Dileepan, The
University of Kansas Medical Center,
USA
Bo Shen, University of Illinois at
Chicago, USA
*Correspondence:
Luke A. J. O’Neill , School of
Biochemistry and Immunology, Trinity
Biomedical Science Institute, Trinity
College Dublin, 152-160 Pearse
Street, Dublin 2, Ireland
e-mail: laoneill@tcd.ie

Studying the metabolism of immune cells in recent years has emphasized the tight link
existing between the metabolic state and the phenotype of these cells. Macrophages in
particular are a good example of this phenomenon. Whether the macrophage obtains its
energy through glycolysis or through oxidative metabolism can give rise to different phenotypes. Classically activated or M1 macrophages are key players of the first line of defense
against bacterial infections and are known to obtain energy through glycolysis. Alternatively activated or M2 macrophages on the other hand are involved in tissue repair and
wound healing and use oxidative metabolism to fuel their longer-term functions. Metabolic
intermediates, however, are not just a source of energy but can be directly implicated in
a particular macrophage phenotype. In M1 macrophages, the Krebs cycle intermediate
succinate regulates HIF1α, which is responsible for driving the sustained production of
the pro-inflammatory cytokine IL1β. In M2 macrophages, the sedoheptulose kinase carbohydrate kinase-like protein is critical for regulating the pentose phosphate pathway. The
potential to target these events and impact on disease is an exciting prospect.
Keywords: metabolism, macrophage, HIF, glycolysis, PGC-1β

INTRODUCTION
Early insights into the metabolic status of macrophages date back
to the pioneering work carried out by G. C Hard more than 40 years
ago. Hard showed that activated murine peritoneal macrophages
had lower levels of oxygen consumption than resting ones as well
as higher levels of glycolysis (1). This study provided the first evidence of a significant metabolic change in the macrophage as a
consequence of activation. Studies by Newsholme and colleagues
in the 1980s provided further evidence supporting this idea, as
they were able to show that enzymes involved in glucose metabolism have higher enzymatic activities in macrophages, resulting in
high rates of glucose and glutamine consumption (2).
Shortly afterward, in the early 1990s, a role for IL4 in
macrophage activation was described, as well as the concept of
alternative activation (3, 4). At this stage, a distinction was made
between classically activated macrophages, also known as M1,
and alternatively activated macrophages, also referred to as M2.
M1 macrophages are activated by bacterial-derived products such
as lipopolysaccharide (LPS), as well as by signals associated with
infection such as IFNγ. This type of activation results in a highly
inflammatory macrophage with high phagocytic and bactericidal
potential. M2 macrophages on the other hand can be activated
by parasitic products as well as signals associated with parasitic
infections, such as the cytokines IL4 and IL13. This gives rise to a
macrophage with anti-parasitic and tissue repair functions (5).
Also during this period, further research was carried out on the
metabolic changes associated with macrophage activation. Bustos and Sobrino suggested for the first time that the inhibition
of cytokine production in macrophages caused by glucocorticoids could be due to the inhibition of the glycolytic enzymes
PFK1 and PFK2, thus directly implicating impaired metabolism
with impaired function (6). A key discovery was, however, in arginine metabolism. Inés María Corraliza and colleagues were able

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to show that different enzymes responsible for the metabolism of
arginine would be induced in a macrophage depending on the
type of activation. In an M1 macrophage, nitric oxide synthase
(iNOS) is upregulated, resulting in the catabolism of arginine to
citrulline and nitric oxide, the latter playing key role in the intracellular killing of pathogens. In an M2 macrophage on the other
hand, arginase-1 (Arg1) is induced, which results in the production
of urea, polyamines, and ornithine, which are important for the
wound healing actions of this macrophage population (7, 8). The
differential metabolism of arginine is as of today, one of the most
reliable discriminating factors between M1 and M2 macrophages.
In fact, it is the only factor identified so far that can be used to detect
M2 macrophage polarization in human samples (9). This provides
an example of how studying the metabolic status of macrophages
has proven more useful than studying function alone as well as
unveiling the potential for therapeutic targeting in disease.

GLYCOLYTIC M1 VERSUS OXIDATIVE M2 MACROPHAGES
Although studies into the metabolism of immune cells date back
a few decades, it has only been in recent years that the tight link
between metabolism and function has become apparent. The clear
metabolic differences existing between M1 and M2 macrophages
exemplify this idea. An M1 macrophage is part of the first line of
defense of the innate immune system, which takes place within
hours to days, as opposed to an M2 macrophage, which plays a
bigger role within the resolution phase and thus has longer-term
functions. Their metabolism is unsurprisingly a clear reflection of
those functions.
In M1 macrophages, aerobic glycolysis is induced upon activation, which involves an increase in glucose uptake as well as the
conversion of pyruvate to lactate (Figure 1). At the same time,
the activities of the respiratory chain are attenuated, allowing
for reactive-oxygen species (ROS) production. Further evidence

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Galván-Peña and O’Neill

FIGURE 1 | Metabolic profile of an M1 macrophage is shown.
Classically activated macrophages induce an aerobic glycolytic program
that results in lactate production and increased levels of intermediates of
the Krebs cycle. The HIF1α transcription factor also becomes activated
and can drive production of pro-inflammatory cytokines. The key

for this is provided when treating macrophages with the electron transport chain inhibitors rotenone and antimycin A as this
mimics the effects of toll-like receptor (TLR) agonists in driving
ROS production from the mitochondria (10). Furthermore, the
pentose phosphate pathway is also induced following classical activation. This pathway is key for the generation of NADPH for the
NADPH oxidase, which is important for ROS production, but also
for nitric oxide synthesis (11). Altogether, these metabolic events
can provide the cell with rapid energy and reducing equivalents,
which are required for bactericidal activity. M2 macrophages on
the other hand obtain much of their energy from fatty acid oxidation and oxidative metabolism, which can be sustained for longer.
Following activation, they can induce expression of constituents
of the electron transport chain that will perform oxidative phosphorylation as well as driving the pyruvate into the Krebs cycle
(Figure 2). The pentose phosphate pathway is also more limited in
M2 macrophages. Blocking oxidative metabolism not only blocks
the M2 phenotype but also drives the macrophage into an M1 state.
Similarly, forcing oxidative metabolism in an M1 macrophage
potentiates the M2 phenotype (12, 13). These key metabolic differences between differentially activated macrophages are widely

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Metabolic reprogramming in macrophage polarization

functional consequences are bacterial killing, mostly through the
production of ROS and NO, and inflammation, which occurs via cytokine
production. G6P, glucose-6-phosphate; F6P, fructose-6-phosphate; R5P,
ribulose-5-phosphate; S7P, sedoheptulose phosphate; NO, nitric oxide;
ROS, reactive-oxygen species.

accepted; however, the switches responsible for orchestrating these
different profiles at the molecular level remain largely unknown
and how exactly the cell’s metabolic status regulates polarization
is not yet well understood.
Following classical activation, there is a switch in the expression of 6-phosphofructose-2-kinase/fructose-2,6-bisphosphatase
(PFK2) isoforms from the liver-form (L-PFK2) to the more active
ubiquitous form (u-PFK2), leading to fructose-2,6-bisphosphaste
accumulation, which pushes the glycolytic flux. This switching
occurs at the transcriptional level with the L-PFK2 gene, PFKB3,
being induced following activation (12). Additionally, there seems
to be a requirement for downregulation of the carbohydrate
kinase-like protein (CARKL) for the development of an M1
phenotype. CARKL catalyzes the production of sedoheptulose7-phosphate, an intermediate of the pentose phosphate pathway
(Figure 1). Besides expression levels of CARKL rapidly decreasing following classical activation, CARKL-expressing cells show
defects in LPS-induced superoxide production. Furthermore,
overexpression of CARKL results in a decrease in the production of pro-inflammatory cytokines in accordance with an M2
phenotype. Altogether, this would suggest that CARKL may help

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FIGURE 2 | Metabolic profile of an M2 macrophage is shown. Alternatively
activated macrophages trigger a metabolic program including the electron
transport chain as well as fatty acid β-oxidation, which is orchestrated by

drive the macrophage metabolism toward increased pentose phosphate pathway activity and increased redox state, thus supporting
M1 polarization (14). Finally, activation of macrophages with LPS
results in increased levels of Krebs cycle intermediates such as succinate and malate. Succinate, in particular, was shown to drive IL1β
production through HIF1α, a response that could be blocked by
inhibition of glycolysis using 2-deoxyglucose (15). This exemplifies how the macrophage metabolism is not simply needed for providing the energy required but can also have a direct involvement
in the transcriptional regulation of the immune response.
Following alternative activation, the PFKB1 gene instead of the
PFKB3 is expressed, resulting in higher levels of the liver isoform
of PFK2 and lower levels of fructose-2,6-bisphosphate. The lower
glycolytic levels are compensated with an increase in oxidative
phosphorylation. Following macrophage activation with IL4, there
is massive induction of an oxidative metabolic program, ranging
from fatty acid uptake and oxidation, to oxidative phosphorylation and mitochondrial respiration. The mechanism behind this
increase is somewhat better understood than that of glycolysis in
M1 macrophages. Following IL4 treatment, the transcription factor STAT6, which is responsible for mediating the transcriptional
responses of this cytokine, becomes activated. Active STAT6 can
induce the coactivator protein peroxisome proliferator-activated
receptor (PPAR)γ-coactivator-1β (PGC-1β). PGC-1β can induce
mitochondrial respiration as well as mitochondrial biogenesis.
Furthermore, together with the transcription factors, nuclear respiratory factor 1 (NRF-1) and estrogen-related receptor-α (ERRα),
it drives the production of key mitochondrial components, such
as cytochrome c and ATP synthase (16, 17). It is therefore not

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Metabolic reprogramming in macrophage polarization

STAT6 and PGC-1β. Arg1 also drives the production of polyamines and
ornithine. The key functional consequences are tissue repair and anti-parasitic
responses.

surprising that PGC-1β is considered as the key player responsible
for the metabolic switch in M2 macrophages (Figure 2). In fact,
knockdown of PGC-1β impairs not only the metabolic profile of
M2 macrophages but also their functions (13). Furthermore, while
PGC-1β is the key trigger, PPARs, particularly PPARγ and PPARδ,
have a key role in maintaining the phenotype. PPARδ is responsible
for orchestrating the effector functions of alternative activation,
for instance, expression of the macrophage galactose-type C-type
lectin 1 (MGL-1) as well as costimulatory molecules and other
factors involve in the anti-inflammatory response. PPARγ on the
other hand, is involved in the transcription of different factors
required for β-oxidation of fatty acids (18, 19).
Recently, the protein TNF-alpha-induced protein 8-like 2
(TIPE2) has also been associated with an M2 phenotype, through
the induction of arginine metabolism, which as already mentioned, is the most distinguished metabolic feature of M2
macrophages. Interestingly, TIPE2 exerts such function following long-term classical activation of macrophages with LPS and
not alternative activation. Thus, TIPE2 uses the switching to arginine metabolism to negatively regulate inflammation, and can
therefore re-program a classically activated macrophage into its
anti-inflammatory counterpart (20).

HYPOXIA-INDUCIBLE FACTOR IN MACROPHAGE
POLARIZATION
Macrophages, as well as other immune cells, are usually found in
inflamed sites, which are characterized by low oxygen levels. The
transcription factor HIF thus plays an important role as one of
the key mediators in the adaptation of macrophages to hypoxic

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Metabolic reprogramming in macrophage polarization

conditions. This heterodimeric protein is composed of two subunits, an α and a β subunit. Three isoforms of the oxygen-sensitive
α subunit have been identified. The HIF1α isoform is expressed
ubiquitously, and is tightly linked to the inflammatory response
and microbicidal activities. HIF2α on the other hand, is expressed
in a more limited fashion, but it is present in myeloid cells (21,
22). There is evidence in the literature suggesting a role for the two
HIFα isoforms, 1 and 2, in macrophage polarization. While HIF1α
has been associated with classical macrophage activation, HIF2α
has been recently linked to an M2 phenotype. These differential
roles are, however, far from clear.
HIF1α expression can be driven by different classical activators
through NF-κB, resulting in the production of pro-inflammatory
cytokines and other mediators of the M1 phenotype, such as glycolytic enzymes and glucose transporters. HIF2α expression, on
the other hand, occurs independently of NF-κB, which would be in
accordance with alternative activation. Interestingly though, both
isoforms seem to be important in maintaining levels of the NFκB subunit p65 (23). A key mediator regulated by HIF1α is the
M1 marker iNOS. Under hypoxic conditions, nitric oxide production through iNOS is HIF1α-dependent thus implicating HIF1α
in bacterial clearance (24). In fact, HIF1α−/− macrophages have
impaired capacity to clear both Gram-positive and Gram-negative
bacteria. Nevertheless, superoxide production during the respiratory burst, which is also required for bacterial clearance, seems
to be a HIF1α-independent event (25). This is, interestingly, not
the only HIF1α-independent event that occurs following classical
activation. A critical event in the reprograming of metabolism to
glycolysis is the switch from L-PFK2 to u-PFK2, which also occurs
independently of HIF1α (12). This would suggest the presence of
some other yet unidentified factor responsible for mediating the
metabolic switch in M1 macrophages, either independently or in
association with HIF1α.
The potential role of HIF2α in promoting the M2 phenotype,
although promising, remains obscure. HIF2α has been shown to
regulate transcription of the M2 marker, Arg1. This finding is
supported by the half-life of both proteins, as both the mRNAs
for HIF2α as well as Arg1, have relatively long half-lives. The
mRNAs for HIF1α and iNOS, however, are relatively short-lived
(24). This would agree with the initial idea of how the metabolism of polarized macrophages goes hand in hand with the timing
of their functions, and would support the association of HIF1α
and HIF2α with M1 and M2 macrophages, respectively. There are,
however, incongruences regarding the role of HIF2α. For instance,
HIF2α also controls IL1β production, which is associated with an
M1 phenotype rather than M2 (15). Additionally, HIF2α has also
been associated with NF-κB activity as mentioned above, which is
also associated with an M1 phenotype (23). Studies have shown,
however, that both isoforms seem to have redundant and overlapping functions, even though when one is knocked down, the
other does not seem to be able to compensate (23). This highlights
the fact that there are still major gaps in our understanding of the
differential activities of the two isoforms.

that have been carried out using human macrophages have identified major interspecies differences. For instance, classic murine
M2 macrophage markers, such as Ym1 or Fizz1, lack human
homologs and can therefore not be used as markers in human
macrophages (26).
Interestingly, a recent proteomic analysis of differentially activated human macrophages suggests that the major functional differences between the two lie within metabolic pathways. The study
identifies major metabolic enzymes such as glucose-6-phosphate
dehydrogenase, fructose-1,6-bisphosphatase 1 (Fbp1), alpha enolase, and fructose bisphosphate aldolase A as being differentially
expressed in human M1 and M2 macrophages (27). In agreement
with murine studies, human M1 macrophages also upregulate glycolysis to give rise to a pro-inflammatory phenotype characterized
by the production of cytokines such as IL12p40, TNFα, or IL6.
However, oxidative metabolism and fatty acid oxidation do not
seem to predominate in human M2 macrophages, but instead,
gluconeogenesis, driven by Fbp1, seems to play a major role (27).
This finding is supported by a subsequent study suggesting that
fatty acid oxidation is dispensable in human M2 macrophages.
D. Namgaladze and B. Brüne show that IL4-induced human M2
macrophages do not induce PGC-1β, the key transcription factor responsible for driving the fatty acid oxidation program. In
contrast with murine studies, the use of a fatty acid oxidation
inhibitor does not impair the ability of human macrophages to
produce high levels of CCL18 and Mrc1 and low levels of IL1-β
and IL6, suggesting that they maintain the M2 phenotype (28).
Another major aspect that seems to differ considerably between
murine and human macrophages is the role of iNOS and Arg1
in M1 and M2 macrophages, respectively. Attempts to demonstrate significant production of NO by human macrophages in
culture have mostly failed. When successfully detected, it has only
been after a period of stimulation of a few days and in much
smaller amounts than that detected in murine macrophages (29).
Furthermore, a recent report indicates that epigenetic modifications silence the nos2 gene in humans, suggesting that there is
no role for iNOS in M1-mediated inflammation (30). Intriguingly, macrophage-derived NO production has been reported in
cases of acute inflammation such as those presented by rheumatoid arthritis patients as well as those suffering from malaria (31,
32). On the other hand, it is not just the role of iNOS that has
been questioned, but also that of Arg1. Neither Cameron et al. nor
Sheemann et al. could detect any arginase activity from human
macrophages in culture (33, 34). However, Anika Geelhaar-Karsch
and colleagues have recently shown that patients suffering from
classical Whipple’s disease, which is associated with elevated levels
of M2 macrophages, present with higher levels of arginase activity
as well as Arg1-derived products, such as urea (9). Interestingly,
this could only be detected in plasma and fresh biopsies and not in
macrophages in culture. Therefore, although the major differences
existing between mice and humans in this regard are undisputable,
the switch toward iNOS versus Arg1 may still play an important
role in human diseases.

POLARIZATION OF HUMAN MACROPHAGES

FINAL PERSPECTIVES

Most current knowledge of macrophage polarization comes from
murine studies; however, our understanding of this topic in human
macrophages remains quite poor. Furthermore, the limited studies

The metabolic aspects behind macrophage activation have long
been an area of interest for many. Metabolism as a key aspect
of macrophage polarization, however, is an intriguing area within

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Galván-Peña and O’Neill

macrophage biology that has only started to develop more recently.
Although we are still very much in the dark regarding our understanding of the metabolic molecular events driving macrophage
polarization, the evidence discussed suggests that the role of metabolic intermediates is much more important than expected. The
key question is why M1 and M2 macrophages would have such
different metabolic profiles? It is possible that M1 macrophages
are mainly found in hypoxic environments and therefore have to
rely on glycolysis, produced via HIF1α, for their ATP production.
Glycolysis can also be rapidly induced, which is perhaps needed for
the rapid activation that occurs in M1 macrophages during infection. The attenuation in the respiratory chain will also allow M1
macrophages to produce ROS, as will the NADPH produced by
the pentose phosphate pathway, which is required for the NADPH
oxidase. For M2 macrophages, acute activation is less of an issue, as
their main function is in wound healing and anti-parasitic defense.
M2 macrophages also do not generate ROS and therefore have a
fully functional respiratory redox chain, allowing for oxidation
of fatty acids. β-oxidation of fatty acids has, in fact, been shown
to be anti-inflammatory, possibly because of a decrease in the
production of prostaglandins, although this is not fully understood (35). Perhaps, the more sustained role of M2 macrophages
mainly involves the metabolism of fat reserves with less ROS being
a safe-ground against injury during tissue repair.
The translation of these discoveries to human diseases is an
intriguing prospect, especially, since there are diseases that have
been associated with one particular macrophage phenotype or
another. For instance, patients presenting with chronic venous
ulcers suffer from chronic inflammation as a result of failing to
switch from M1 macrophages to M2 (36). On the other hand,
those suffering from classical Whipple’s disease, a result of chronic
infection caused by Tropheryma whipplei, fail to clear the infection due to the lack of inflammation and excess presence of M2
macrophages (37). Interestingly, there are also reports suggesting
that the distribution of M1 and M2 macrophages varies between
males and females. In fact, the higher incidence of asthma in
female mice was associated with higher levels of M2 macrophages
when compared to male mice (38). Since females are known to
present with higher incidence of not only asthma but also other
autoimmune diseases, it would be interesting to speculate whether
gender-associated differences in macrophage polarization might
play a role.
Finally, manipulation of macrophage polarization has already
proven to be somewhat successful clinically. Administration of
the classical M1 macrophage activator IFNγ had beneficial effects
in patients with ovarian carcinoma (39, 40). Therefore, our current understanding of the metabolic status of differentially activated macrophages holds great potential for clinical applications,
although further research is required in order to capitalize clinically on the observations made to date in both murine and human
systems.

REFERENCES
1. Hard GC. Some biochemical aspects of the immune macrophage. Br J Exp Pathol
(1970) 51(1):97–105.
2. Newsholme P, Curi R, Gordon S, Newsholme EA. Metabolism of glucose, glutamine, long-chain fatty acids and ketone bodies by murine macrophages. Biochem
J (1986) 239(1):121–5.

www.frontiersin.org

Metabolic reprogramming in macrophage polarization

3. Abramson SL, Gallin JI. IL4 inhibits superoxide production by human mononuclear phagocytes. J Immunol (1990) 144(2):625–30.
4. Stein M, Keshaw S, Harris N, Gordon S. Interleukin 4 potently enhances murine
macrophages mannose receptor activity: a marker of alternative immunologic
macrophage activation. J Exp Med (1992) 176(1):287–92. doi:10.1084/jem.176.
1.287
5. Odegaard JI, Chawla A. Alternative macrophage activation and metabolism.
Annu Rev Pathol (2011) 6:275–97. doi:10.1146/annurev-pathol-011110-130138
6. Bustos R, Sobrino F. Stimulation of glycolysis as an activation signal in rat peritoneal macrophages: effect of glucocorticoids on this process. Biochem J (1992)
282:299–303.
7. Corraliza IM, Soler G, Eichmann K, Modolell M. Arginase induction by suppressors of nitric oxide synthesis (IL4, IL10, PGE2 ) in murine bone-marrowderived macrophages. Biochem Biophys Res Commun (1995) 206(2):667–73.
doi:10.1006/bbrc.1995.1094
8. Munder M, Eichmann K, Modolell M. Alternative metabolic states in murine
macrophages reflected by the nitric oxide synthase/arginase balance: competitive regulation by CD4+ T cells correlates with Th1/Th2 phenotype. J Immunol
(1998) 160(11):5347–54.
9. Geelhard-Karsch A, Schinnerling K, Conrad K, Friebel J, Allers K, Schneider T, et al. Evaluation of arginine metabolism for the analysis of m1/m2
macrophage activation in human clinical specimens. Inflamm Res (2013)
62:865–9. doi:10.1007/s00011-013-0642-z
10. West AP, Brodsky IE, Rahner C, Woo DK, Erdjument-Bromage H, Tempst P,
et al. TLR signaling augments macrophage bactericidal activity through mitochondrial ROS. Nature (2011) 472(7344):476–80. doi:10.1038/nature09973
11. Aktan F. iNOS-mediated nitric oxide production and its regulation. Life Sci
(2004) 75(6):639–53. doi:10.1016/j.lfs.2003.10.042
12. Rodríguez-Prados JC, Través PG, Cuenca J, Rico D, Aragonés J, MartínSanz P, et al. Substrate fate in activated macrophages: a comparison between
innate, classic and alternative activation. J Immunol (2010) 185(1):605–14.
doi:10.4049/jimmunol.0901698
13. Vats D, Mukundan L, Odegaard JI, Zhang L, Smith KL, Morel CR, et al. Oxidative metabolism and PGC-1β attenuate macrophage-mediated inflammation.
Cell Metab (2006) 4(1):13–4. doi:10.1016/j.cmet.2006.08.006
14. Haschemi A, Kosma P, Gille L, Evans CR, Burant CF, Starkl P, et al. The sedoheptulose kinase CARKL directs macrophage polarization through control of
glucose metabolism. Cell Metab (2012) 15(6):813–26. doi:10.1016/j.cmet.2012.
04.023
15. Tannahill GM, Curtis AM, Adamik J, Palsson-McDermott EM, McGettrick AF,
Goel G, et al. Succinate is an inflammatory signal that induces IL-1β through
HIF-1α. Nature (2013) 496(7444):238–42. doi:10.1038/nature11986
16. St-Pierre J, Lin J, Krauss S, Tarr PT, Yang R, Newgard CB, et al. Bioenergetic analysis of peroxisome proliferator-activated receptor gamma coactivators
1alpha and 1beta (PGC-1α and PGC-1β) in muscle cells. J Biol Chem (2003)
278(29):26597–603. doi:10.1074/jbc.M301850200
17. Shao D, Liu Y, Liu X, Zhu L, Cui Y, Cui A, et al. PGC-1 beta-regulated mitochondrial biogenesis and function in myotubes is mediated by NRF-1 and ERR
alpha. Mitochondrion (2010) 10(5):516–27. doi:10.1016/j.mito.2010.05.012
18. Odegaard JI, Ricardo-Gonzalez RR, Goforth MH, Morel CR, Subramanian
V, Mukundan L, et al. Macrophage-specific PPARγ controls alternative activation and improves insulin resistance. Nature (2007) 447(7148):1116–20.
doi:10.1038/nature05894
19. Kang K, Reilly SM, Karabacak V, Gangl MR, Fitzgerald K, Hatano B, et al.
Adipocyte-derived Th2 cytokines and myeloid PPARδ regulate macrophage
polarization and insulin sensitivity. Cell Metab (2008) 7(6):485–95. doi:10.1016/
j.cmet.2008.04.002
20. Lou Y, Zhang G, Geng M, Zhang W, Cui J, Liu S. TIPE2 negatively regulates
inflammation by switching arginine metabolism from nitric oxide synthase to
arginase. PLoS One (2014) 9(5):e96508. doi:10.1371/journal.pone.0096508
21. Cramer T, Yamaniski R, Clausen BE, Förster I, Pawlinski R, Mackman N,
et al. HIF1α is essential for myeloid cell-mediated inflammation. Cell (2003)
112(5):645–57. doi:10.1016/S0092-8674(03)00154-5
22. Talks KL, Turley H, Gatter KC, Maxwell PH, Pugh CW, Ratcliffe PJ, et al. The
expression and distribution of the hypoxia-inducible factors HIF1α and HIF2α
in normal human tissues, cancers, and tumor-associated macrophages. Am
J Pathol (2000) 157(2):411–21. doi:10.1016/S0002-9440(10)64554-3
23. Fang HY, Hughes R, Murdoch C, Coffelt SB, Bismas SK, Harris AL, et al.
Hypoxia-inducible factors 1 and 2 are important transcriptional effectors

September 2014 | Volume 5 | Article 420 | 279

Galván-Peña and O’Neill

24.

25.

26.

27.

28.

29.
30.

31.

32.

33.

34.

in primary macrophages experiencing hypoxia. Blood (2009) 114(4):844–59.
doi:10.1182/blood-2008-12-195941
Takeda N, O’Dea EL, Doedens A, Kim JW, Weidemann A, Stockmann
C, et al. Differential activation and antagonistic function of HIF-α isoforms in macrophages are essential for NO homeostasis. Genes Dev (2010)
24(5):491–501. doi:10.1101/gad.1881410
Peyssonnaux C, Datta V, Cramer T, Doedens A, Theodorakis EA, Gallo RL, et al.
HIF1α expression regulates the bactericidal capacity of phagocytes. J Clin Invest
(2005) 115(7):1806–15. doi:10.1172/JCI23865
Raes G, Van den Bergh R, De Baetselier P, Ghassabeh GH, Scotton C, Locati M,
et al. Arginase-1 and Ym1 are markers for murine, but not human, alternatively
activated myeloid cells. J Immunol (2005) 174(11):6561. doi:10.4049/jimmunol.
174.11.6561 author reply 6561-6562,
Reales-Calderon JA, Aguilera-Montilla N, Corbi AL, Molero G, Gil C. Proteomic characterization of human proinflammatory M1 and anti-inflammatory
M2 macrophages and their response to Candida albicans. Proteomics (2014)
14(12):1503–18. doi:10.1002/pmic.201300508
Namgaladze D, Brüne B. Fatty acid oxidation is dispensable for
human macrophage IL4-induced polarization. Biochem Biophys Acta (2014)
1841(9):1329–35. doi:10.1016/j.bbalip.2014.06.007
Albina JE. On the expression of nitric oxide synthase by human macrophages.
Why no NO? J Leukoc Biol (1995) 58(6):643–9.
Gross TJ, Kremens K, Powers LS, Brink B, Knutson T, Domann FE, et al.
Epigenetic silencing of the human NOS2 gene: rethinking the role of nitric
oxide in human macrophage inflammatory responses. J Immunol (2014)
192(5):2326–38. doi:10.4049/jimmunol.1301758
St Clair EW, Wilkinson WE, Lang T, Sanders L, Misukonis MA, Gilkeson GS,
et al. Increased expression of blood mononuclear cell nitric oxide synthase
type 2 in rheumatoid arthritis patients. J Exp Med (1996) 184(3):1173–8.
doi:10.1084/jem.184.3.1173
Anstey NM, Weinberg JB, Hassanali MY, Mwaikambo ED, Manyenga D, Misukonis MA, et al. Nitric oxide in Tanzanian children with malaria: inverse relationship between malaria severity and nitric oxide production/nitric oxide synthase
type 2 expression. J Exp Med (1996) 184(2):557–67. doi:10.1084/jem.184.2.557
Cameron ML, Granger DL, Weinberg JB, Kozumbo WJ, Koren HS. Human
alveolar and peritoneal macrophages mediate fungistasis independently of Larginine oxidation to nitrite or nitrate. Am Rev Respir Dis (1990) 142:1313–9.
doi:10.1164/ajrccm/142.6_Pt_1.1313
Schneeman M, Schoedon G, Hofer S, Blau N, Guerrero L, Shaffner A. Nitric oxide
synthase is not a constituent of the antimicrobial armature of human mononuclear phagocytes. J Infect Dis (1993) 167:1358–63. doi:10.1093/infdis/167.6.1358

Frontiers in Immunology | Inflammation

Metabolic reprogramming in macrophage polarization

35. Steinberg GR, Schertzer JD. AMPK promotes macrophage fatty acid oxidative metabolism to mitigate inflammation: implications for diabetes and
cardiovascular disease. Immunol Cell Biol (2014) 92(4):340–5. doi:10.1038/icb.
2014.11
36. Sindrilaru A, Peters T, Wieschalka S, Baican C, Baican A, Peter H, et al. An
unrestrained proinflammatory M1 macrophage population induced by iron
impairs wound healing in humans and mice. J Clin Invest (2011) 121(3):985–97.
doi:10.1172/JCI44490
37. Moos V, Schmidt C, Geelhar A, Kunkel D,Allers K, Schinnerling K, et al. Impaired
immune functions of monocytes and macrophages in Whipple’s disease. Gastroenterology (2010) 138(1):210–20. doi:10.1053/j.gastro.2009.07.066
38. Melgert BN, Oriss TB, Qi Z, Dixon-McCarthy B, Geerlings M, Hylkema MN,
et al. Macrophages: regulators of sex differences in asthma? Am J Respir Cell Mol
Biol (2010) 42(5):595–603. doi:10.1165/rcmb.2009-0016OC
39. Allavena P, Peccatori F, Maggioni D, Erroi A, Sironi M, Colombo N, et al.
Intraperitoneal recombinant gamma interferon in patients with recurrent ascetic
ovarian carcinoma: modulation of cytotoxicity and cytokine production in
tumor-associated effectors and of major histocompatibility antigen expression
on tumor cells. Cancer Res (1990) 50(22):7318–23.
40. Colombo N, Peccatori F, Paganin C, Bini S, Brandely M, Mangioni C, et al.
Anti-tumor and immunomodulatory activity of intraperitoneal IFN-gamma in
ovarian carcinoma patients with minimal residual tumor after chemotherapy.
Int J Cancer (1992) 51(1):42–6. doi:10.1002/ijc.2910510109

Conflict of Interest Statement: The authors declare that the research was conducted
in the absence of any commercial or financial relationships that could be construed
as a potential conflict of interest.
Received: 11 July 2014; paper pending published: 04 August 2014; accepted: 18 August
2014; published online: 02 September 2014.
Citation: Galván-Peña S and O’Neill LAJ (2014) Metabolic reprograming in
macrophage polarization. Front. Immunol. 5:420. doi: 10.3389/fimmu.2014.00420
This article was submitted to Inflammation, a section of the journal Frontiers in
Immunology.
Copyright © 2014 Galván-Peña and O’Neill. This is an open-access article distributed
under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s)
or licensor are credited and that the original publication in this journal is cited, in
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permitted which does not comply with these terms.

September 2014 | Volume 5 | Article 420 | 280