The Mononuclear Phagocyte System: The Relationship between Monocytes and

Macrophages

David A. Hume1, Katharine M Irvine1 and Clare Pridans2

1. Mater Research Institute-University of Queensland, Translational Research Institute, 37

Kent Street, Woolloongabba, Qld 4102, Australia

2. University of Edinburgh Centre for Inflammation Research, 47 Little France Crescent,

Edinburgh EH16 4TJ, UK

Correspondence to David.Hume@uq.edu.au

Abstract

The mononuclear phagocyte system (MPS) is defined as a cell lineage in which committed

marrow progenitors give rise to blood monocytes and tissue macrophages. Here we discuss

the concept of self-proscribed macrophage territories and homeostatic regulation of tissue

macrophage abundance through growth factor availability. Recent studies have questioned

the validity of the MPS model and argued that tissue-resident macrophages are a separate

lineage seeded during development and maintained by self-renewal. We discuss the

limitations of inbred mouse models of monocyte-macrophage homeostasis and summarise

evidence that during postnatal life monocytes replace resident macrophages in all major

organs and adopt their tissue-specific gene expression. We conclude that the MPS remains

a valid and accurate framework for understanding macrophage development and

homeostasis.

Key words

monocyte, macrophage, heterogeneity, ontogeny, homeostasis, CSF1R

1

Macrophage ontogeny and the mononuclear phagocyte system

The mononuclear phagocyte system (MPS) was originally proposed as a new classification of

macrophages, blood monocytes and their precursors, based upon criteria of morphology,

function, origin and kinetics [1, 2]. The concept was based upon many lines of evidence that

in adult mice resident tissue macrophage populations are maintained through replacement

by monocytes in the steady state. Monocytes and macrophages share expression of many

surface markers and dependence upon specific growth factors and transcriptional regulators

including the lineage-specific factor, PU.1 (reviewed in [3]). The classical dendritic cell (DC)

was initially excluded from the MPS [1]. There is still ongoing nomenclature debate

surrounding the definition of DCs. For example, one study [4] reported that phagocytes in

the T cell areas of lymph nodes previously considered to be DCs could be separated from

classical DC -- as defined by Steinman and Cohn and by current criteria—and should be

considered to be macrophages. The lineage relationship between DC and macrophages has

been previously reviewed [5, 6]. For the purpose of this Opinion piece, we will consider the

MPS to be confined to definitive monocytes and macrophages.

Over the past 5 years, there has been a cascade of studies of macrophage ontogeny in

multiple organs that claim to have challenged the MPS concept and to replace it with a view

that resident macrophages are maintained entirely by self-renewal. It is therefore timely to

reconsider the validity of the MPS as a model for understanding macrophage biology. We

will argue that the MPS concept remains as valid today as when it was originally conceived.

Embryonic development of the MPS

Macrophages develop in the mouse embryo in three waves, starting with phagocytes

derived from the yolk sac independently of blood monocytes, then blood monocytes

produced in the fetal liver, and finally, definitive monocytes derived from bone marrow

2

hematopoietic stem cells (HSC) [7]. Yolk sac-derived macrophages enter the mouse embryo

via the emerging vasculature [8]. In vivo imaging of a macrophage-restricted fluorescent

reporter gene also revealed the first appearance of these phagocytes in the chick yolk sac,

their rapid proliferation, entry into the vasculature, and immediate engagement in

phagocytosis [9]. In the chick, yolk sac cells from an enhanced green fluorescent protein

(EGFP)-positive donor could be transplanted in ovo [12]. These cells gave rise to

macrophages throughout the embryo, but they were lost by the time of hatch. Similar

transplantation of bone marrow cells gave rise to long-lived chimerism indicating that

definitive HSC rather than fetal progenitors are the precursors of adult resident

macrophages [10]. The earliest yolk sac-derived phagocytes are produced in mice that lack

the transcription factor C-MYB which is required for definitive hematopoiesis [11]. The yolk

sac may also be the initial origin of a MYB-dependent erythro-myeloid progenitor (EMP) that

gives rise to the second wave of monocyte-macrophage generation in the foetal liver, which

in turn seeds resident tissue macrophages in many major organs [12, 13]. EMP are distinct

from the definitive HSC which first arise in the aorta-gonad-mesonephros (AGM) region of

the mouse embryo around 9.5 to 10.5 days post coitum and which also seed the liver [12,

13]. AGM-derived HSC also contribute significantly to the development of foetal monocytes

and most tissue macrophage populations [14].

Monocyte Development

Based upon thymidine labelling, van Furth & Cohn [2] demonstrated that the proliferating

monocyte precursor pool resided in the bone marrow. This population, the committed

monocyte progenitor, was identified and characterised in mouse bone marrow downstream

of a shared monocyte-DC progenitor (MDP) [15]. Blood monocytes are not homogeneous.

They have been divided into subsets, based upon surface markers; LY6C in mice, CD43 in

3

rats, CD163 in pigs and CD14/CD16 in humans. A proposed nomenclature defines the

populations as classical (LY6Chi in mouse and CD14Hi in human) and non-classical (LY6lo,

CD14lo) monocytes. The subsets are distinguished also by differential expression of the

chemokine receptors CCR2 and CX3CR1 [16]. The transit time of mouse blood monocytes

was first estimated to be around 32 hours [2], similar to current estimates of the short half-

life of LY6Chi monocytes [17]. The non-classical monocytes in both mouse and human have a

much longer half-life in the circulation and have a specific function in homeostasis [17, 18].

The monocyte subsets are actually a differentiation series. A detailed comparative analysis

of the transcriptome and epigenome of individual mouse LY6Chi and LY6Clo monocytes

revealed the role of the transcription factor, CEBP in the differentiation process [19]. A

similar study of human monocyte differentiation [20] showed that in contrast to mice,

CEBP was already highly-expressed in CD14hi monocytes. CD14+, CD16+ intermediate

monocytes expressed intermediate levels of all transcripts that defined the extremes [20].

In mice, the generation of LY6Clo monocytes depends upon the transcription factor NR4A1.

Indeed, a conserved subdomain of a NR4A1 enhancer in the mouse genome was required

specifically for NR4A1 expression in monocytes, and for the development of the LY6C lo

subpopulation [21]. This enhancer was shown to bind CEBP and another transcription

factor, KLF2 [21]. Unlike, CEBPB, KLF2 mRNA was also highly-induced during human

monocyte differentiation [20]. The LY6Chi monocyte subset in mice has been further

subdivided. One report [22] defined a subset based upon the marker TREML4 [27]. LY6Chi

monocytes that expressed TREML4 were committed to the LY6C lo monocyte fate whereas

those that lacked TREML4 were able to acquire antigen-presenting cell function in response

to GM-CSF and IL4, Others associated the distinct fates with underlying gene expression of

transcription factors, PU.1 (SFPI1) and NR4A1[23]. There is some evidence that subsets of

4

mouse LY6Chi monocytes arise from distinct committed marrow progenitor subsets, the

granulocyte-macrophage progenitor (GMP) and the monocyte dendritic cell progenitor

(MDP) [24]. These authors characterised the LY6C+ monocytic progeny of the GMP as

“neutrophil-like” sharing some lineage-associated genes with neutrophils. The presence of

monocyte-like cells amongst the “polymorphonuclear” fraction of blood in mice and

humans has been previously described [25]. The more recent study showed that neutrophil-

like monocytes are mobilised independently by different microbial stimuli and have distinct

functions in mouse models of inflammation.

Tissue-specific macrophage heterogeneity, adaptation and function

Resident tissue macrophages also adapt to each tissue environment to perform specific

functions. The gene expression profile of mouse embryonic macrophages is initially

relatively organ-independent [26]. The acquisition of tissue-specific macrophage

phenotypes in mice correlates with organogenesis and maturation of target organs [26, 27].

A recent study identified the transcription factor ZEB2 as a regulator in the acquisition and

maintenance of tissue-specific macrophage adaptation in multiple tissues (see below) [28].

Even within individual adult tissues, macrophages are heterogeneous in terms of surface

markers. For example, the spleen contains functionally distinct macrophage populations in

germinal centres, T cell areas, the marginal zone and the red pulp [29]. In other cases,

macrophage heterogeneity may be a reflection of life history (e.g. the time course of

maturation since entry or since division [30-32]). The past few years have seen a cascade of

detailed functional studies of new macrophage populations and sub-populations within

specific tissues, some of which are summarised in Table 1.

5

Gut Macrophages

Monocytes respond to the local environment to take up tissue-appropriate functions [33].

The largest macrophage population in the mouse, those of the intestinal lamina propria,

turns over continuously and is replaced by blood monocytes in the adult mouse (reviewed in

[34]). The TGF-dependent pathway of monocyte differentiation into macrophages of the

colonic lamina propria [35] requires the rapid down-regulation of inflammatory signalling

modules that would otherwise mediate activation in response to the luminal contents and

rapid up-regulation of receptors involved in apoptotic cell recognition [35]. The

environment of the intestinal wall differs between the small intestine and colon with the

influence of the microbiome, and locally between the lamina propria, crypts, submucosa,

isolated lymphoid follicles and the muscularis externa. For example, a specialised

population of macrophages expressing CD169 (Siglec1) is enriched at the base of crypts [36].

These crypt-associated macrophages are essential to support and regulate the

differentiation of crypt-derived cell types, including Paneth cells and antigen-sampling M

cells [37]. Intestinal macrophages have been segregated based upon expression of the

phosphatidyl serine receptor, TIM4 (TIMD4) and surface CD4. Long-lived TIM4+/CD4+

macrophages constitute a significant proportion of cells in the wall of the mouse small and

large intestine [38] and in mouse Peyer’s patches [39]. A recent study used an inducible YFP

lineage trace to tag long-lived macrophages in the mouse intestinal wall and separated

those of the lamina propria and muscularis [40]. The large majority of macrophages in both

locations were monocyte-derived after 20 weeks. The longer-lived cells were located in the

submucosa and muscularis externa [40]. Shaw et al. [38] compared expression profiles of

blood monocytes with the three populations isolated from the gut, TIM4 -/CD4-, CD4+/TIM4-

and CD4+/TIM4+. Our view of their primary data indicates that all of the populations share

6

high expression of monocyte-enriched transcription factors (IRF8, IRF4, CEBPB, KLF2, KLF4,

NR4A1) discussed above. From monocytes to CD4+/TIM4+ macrophages, there was

progressive upregulation of transcription factors involved in differentiation of other

macrophage populations, notably NR1H3 (LXRA) (implicated in TIMD4 regulation in liver;

[28]) and BATF3 (a key factor in regulation of antigen-presenting cell differentiation in

monocytes [22]). We suggest that like blood monocyte “subsets”, the isolated gut

macrophages are actually a differentiation series.

Liver and Lung Macrophages

The macrophages of the liver (Kupffer cells, KC) and lung each have unique transcriptomic

profiles [26]. Both populations were proposed to be seeded primarily from foetal liver-

derived monocytes and maintained by self-renewal [12, 13]. The liver also contains a

functionally-distinct population of macrophages associated with the outer capsule that

monitors the peritoneal cavity [41]. Three separate studies used depletion of KC in adult

mice to demonstrate that bone marrow-derived monocytes could occupy the vacant

sinusoidal location and adopt the transcriptomic profile and clearance functions of the cells

they replaced [42-44]. A small number of key genes distinguished recent arrivals but the

functional significance of this difference is not clear [42].

In the lung, alveolar macrophages (AM) are unique, and occupy a location within the

airways separated from several distinct interstitial macrophage populations that arise from

different progenitors at different stages of perinatal development [45-48]. Unlike most

tissue macrophages, alveolar macrophages (AM) depend upon signals from the GM-CSF

receptor encoded by the Csf2ra and Csf2rb genes. Nevertheless, monocytes and

macrophages from yolk sac, blood monocytes or bone marrow transplanted into Csf2r-/-

7

mice can reconstitute the missing population and acquire the differentiated AM phenotype

[49].

Microglia in the Brain

Brain macrophage populations, both microglia and perivascular, are established during

embryonic development in adults are maintained independently of monocytes. There is a

postnatal surge of monocyte migration, as evidenced from a novel retroviral lineage trace

injected into the murine foetal liver [50]. The infiltrating monocytes respond to the wave of

apoptotic neuronal cell death that occurs through the brain at that time [51]. Microglia

undergo apoptosis and are replaced by proliferation constantly in the adult mouse brain

[50]. The resident non-microglial macrophages of the brain, associated with the

vasculature and meninges, were also maintained independently of monocytes [52].

Nevertheless, donor LY6Chi monocytes replaced microglia and brain macrophages in

irradiated mouse chimeras [52, 53] and occupied the vacant niche following neonatal bone

marrow transplantation, using macrophage-deficient, PU.1 knockout mice [54]. Donor bone

marrow-derived monocytes, and even macrophages from other locations, could also

establish the microglial network and express microglia-specific transcriptomic profile when

transplanted into a CSF1R -/- mouse at birth [55]. Even partial microglial depletion was

sufficient to enable monocyte-derived cells to enter the brain and occupy the microglial

niche [56]. As in the liver, subtle differences were detected in the gene expression profiles

between recent arrivals and resident microglia generated during embryonic development.

Long-lived and short-lived tissue macrophages.

The turnover and function of relevant tissue resident macrophage in other mouse tissues

mice is summarised in Box 1. In every tissue studied there is evidence of macrophage

heterogeneity and open questions as to whether that heterogeneity reflects progress along

8

a developmental pathway (life history), specialised adaptation to a distinct sub-environment

within the tissue or even cellular migration. In the case of the gut for example, the long-

lived cells were restricted to the submucosa [40], but it is not clear whether they have

resided permanently in that location or migrated from the lamina propria. Without

evidence of location it is unclear whether the subpopulations occupy distinct locations or

are randomly distributed amongst the regular territories shown in Figure 2. For example,

four distinct populations of macrophages were described in the heart by two groups [14,

57]. The two reports differ on their relative abundance and the question of whether they

are replaced by blood monocytes with time. But neither study provides information as to

the locations of those subpopulations in the heart. In overview, resident macrophages are

long-lived. But where lineage trace experiments have been performed in most of these

organs, at least some of the macrophages of foetal origin appear to be replaced gradually by

blood monocytes over the life course and/or can be replaced by monocytes following

ablation (Key Figure, Figure 3).

The CSF1/CSF1R signalling axis controls macrophage homeostasis

The proliferation, differentiation and survival of MPS cells in most organs of the mouse is

dependent upon signals from the macrophage colony-stimulating factor receptor (CSF1R)

initiated by its two growth factor ligands, macrophage colony-stimulating factor (CSF1) and

interleukin (IL)-34. A knockout of the CSF1R gene in mice produces extensive loss of tissue

macrophages and has numerous pleiotropic impacts on development [6, 58]. The

expression of CSF1R mRNA provides one of the earliest markers of the appearance of

macrophages in mouse yolk sac [11, 59]. CSF1R signalling is not absolutely required for

monocytopoiesis but is required for monocyte differentiation to the LY6C lo monocyte subset

[60]. In both embryonic [13] and adult tissues [60] most CSF1R-dependent macrophage

9

populations in mice are rapidly depleted by administration of a blocking anti-CSF1R

antibody.

Consistent with tissue macrophage depletion by CSF1R blockade, administration of CSF1, or

a novel long-acting form, CSF1-Fc, produces a massive expansion of blood monocyte and

macrophage numbers in all tissues in mice, rats and pigs [61-64]. CSF1R-dependence may

be one distinction between monocyte-macrophages and a separate classical DC lineage

dependent upon FLT3 ligand [65]. However, many populations of classical DC defined by

other markers can express CSF1R mRNA and can bind labelled CSF1-Fc [62]. Overall, FLT3-

deficiency, and consequent loss of DC, can be partly compensated in mice by CSF1R signals

[66]. So, there is not likely to be a strict separation of these related lineages in every tissue

based upon growth factor-dependence.

Taken together, these findings led to the concept of homeostatic self-regulation of

monocyte and macrophage numbers by the availability of CSF1 [6]. Since monocytes and

macrophages consume CSF1, they control the availability of their own growth factor. When

monocytes or macrophages are depleted, the available growth factor concentration rises.

Increased CSF1 availability may then act globally or locally. CSF1 can be made in three

isoforms including cell surface, secreted glycoprotein and proteoglycan forms with distinct

functions in complementing phenotypic impacts of the CSF1op/op mutation [67]. A

proteoglycan form might integrate within a local-acting trophic extracellular matrix. The

consumption of CSF1 by LY6Chi monocytes can autoregulate the generation of the LY6C lo

subpopulation [6, 17, 62] and their depletion or absence (for example in CCR2-/- mice) can

lead to increased circulating CSF1, which might in turn promote compensatory resident

macrophage self-renewal. The ability of tissue macrophages to self-renew in the absence of

10

monocytes cannot, therefore, be taken as evidence against a role for monocytes as

progenitors in the steady state.

The homeostatic control of blood and tissue macrophage populations by CSF1 at a systemic

level is linked in turn to regulation of the size of the liver. KC are the largest macrophage

population in direct contact with the blood, and the main site of circulating CSF1 clearance

[62]. CSF1-Fc treatment of mice produced a rapid increase in the size of the liver through

extensive hepatocyte proliferation [61, 64]. Liver hyperplasia is an indirect effect of the

increased liver macrophage numbers, since CSF1R is not expressed by hepatocytes [61, 64].

Nevertheless, fluorescent CSF1 administered to mice does access most tissue macrophage

populations and CSF1-Fc treatment expanded the numbers of macrophages in all organs

except the brain [62]. So, the circulating CSF1 provides a link between all tissue macrophage

populations that could regulate the size of the MPS.

The central importance of the CSF1R signalling axis in macrophage homeostasis requires

that the gene is lineage-restricted in its expression [3]. Figure 2 shows a stylised diagram of

the chromatin structure of the mouse CSF1R locus derived from our own data and published

chromatin accessibility and chromatin immunoprecipitation-sequencing (ChIP-Seq) data on

isolated macrophage populations [68, 69]. The CSF1R locus contains numerous active

enhancers bound selectively by multiple macrophage-specific transcription factors. Based

upon the identification of key regulatory elements, CSF1R-EGFP [70], CSF1R-enhanced cyan

fluorescent protein (ECFP) [71] and recently CSF1R-mApple [62] reporter transgenes have

enabled visualisation of monocytes and macrophages in situ. In the CSF1R-ECFP mouse, the

CSF1R promoter has been mutated to remove a distal promoter element. The so-called

MacBlue reporter transgene identified the earliest foetal yolk sac macrophages arising in

Reichert’s membrane, and abundant macrophages all over the embryo at subsequent

11

developmental stages [71]. However, in adult mice, transgene expression was absent from

the majority of tissue macrophages.

The expression of the original CSF1R-EGFP transgene in isolated peritoneal, lung and bone

marrow-derived macrophages depends upon a highly-conserved intronic enhancer, the Fms

Intronic Regulatory Element (FIRE) [70] (Figure 1). FIRE is the only binding site detected in

the mouse CSF1R locus for the key regulatory transcription factor, IRF8 [68], and also

mediates the regulatory effects of RUNX1 [3]. Homozygous FIRE mutant mice with a germ

line deletion of FIRE (CSF1RFIRE/FIRE) exhibit ablated CSF1R mRNA and protein expression in

bone marrow and blood monocytes, but unlike CSF1R-/- mice, were not monocyte-deficient

[72]. CSF1RFIRE/FIRE mice also lacked microglia, Langerhans cells, and resident kidney, heart

and peritoneal macrophages but CSF1R mRNA was expressed normally in all other

macrophage populations. We conclude that adult tissue macrophages differ from each

other, from those in the embryo, and from blood monocytes, through a distinctive

transcriptional regulatory framework that includes control of CSF1R expression [72]. By

contrast to CSF1R-/- mice, which have severe brain development defects and die before

weaning [58], CSF1RFIRE/FIRE mutant mice have no overt defects in brain development

despite lacking microglia. This suggests that the loss of peripheral macrophage populations

may contribute to the brain phenotype in CSF1R -/- mice, and that the numerous

developmental functions attributed to microglia (see [73] may be at least partly redundant.

Autoregulation of tissue macrophage patterning: macrophage territories

Resident macrophages are an abundant cellular component of every organ in the body. The

availability of reporter genes driven by macrophage-expressed promoters (e.g. CSF1R,

CX3CR1) enables visualisation of their distribution in 3 dimensions using confocal

microscopy, which permits a more global overview of the MPS in each location. Examples

12

include analyses of the function of renal medullary macrophages using CX3CR1-EGFP mice

[74] and of diverse tissues in the CSF1R-mApple mouse [62]. Three features are evident

from a global perspective as shown in a diverse set of tissues in Figure 2: the abundance of

macrophages, the similarity in relative density amongst diverse tissues, and their regular

spacing. The similarity in relative density of macrophages amongst tissues is supported by

analysis of the contribution of macrophage-specific transcripts (e.g. CSF1R, ADGRE1) to total

tissue mRNA in a wide range of organs [27].

Guilliams & Scott [33] proposed the existence of a spatially-defined tissue macrophage

niche, and niche competition, to explain the relative inability of monocytes to replace

resident macrophages in some tissues. The evidence for such a niche is lacking. In the

muscularis externa of the intestine, macrophages are intimately associated with neurons

and implicated in the regulation of intestinal motility [40] but this relationship does not

determine macrophage localisation. During development, monocytes enter the muscularis

before enteric neurons, and their numbers and distribution are unchanged in mice that lack

an enteric nervous system [75]. Apparent associations with other cells in the muscularis,

including interstitial cells of Cajal, endothelial cells and fibroblasts, which are the main local

source of CSF1, also failed to explain the regular patterning. An alternative view is that the

generation of regular spacing is actually macrophage-autonomous. One mechanism that is

known to generate a regular pattern is self-avoidance or mutual repulsion [75]. If that is the

mechanism the macrophage niche is better described as a self-proscribed territory.

Dynamic imaging of microglia in the mouse brain supports the concept of a territory

demonstrating engagement of each cell in active surveillance of its immediate environment

through highly motile filopodia [76]. Notably, the imaging provided evidence of active

repulsion when processes from neighbouring cells encountered one another. Some

13

peripheral macrophages may be quite motile within larger territories [77]. Others, such as

the peritubular renal macrophages [74], have static cell bodies within non-overlapping

territories, but like microglia continuously survey their local milieu through highly motile

filipodia. The phenomenon of contact inhibition of phagocyte locomotion has been

described in detail in drosophila [78]. If the territory model is correct, monocyte infiltration

during inflammation may depend upon the loss of resident occupants of the territory [33] or

specific (and perhaps transient) down-regulation of their repulsive functions. Furthermore,

if macrophages can establish territories in a cell-autonomous manner they may actually

initiate and define the regular spatial organisation of cells of other lineages in their vicinity.

It is beyond the scope of the current topic to review the many examples of macrophage

function in processes such as branching morphogenesis, vascularisation and patterning.

Is there any need to revise the MPS concept?

The MPS framework has been described in the recent literature as a dogma or paradigm

(e.g. [38, 42]). The alternative proposal is that the majority of resident tissue macrophages

are derived from foetal monocytes and “maintained by self-renewal with minimal input

from circulating monocytes”. A new nomenclature was suggested in which tissue

macrophages and monocyte-derived cells were separate entities [5]. There are many

caveats to the interpretation of lineage trace models of monocyte-macrophage

homeostasis, most of which suggest that macrophage turnover is under-estimated in

current models (Box 2). But in any case, van Furth and Cohn [2] did not claim that

macrophages were “continuously replaced” by monocytes. They explicitly recognised that

resident macrophages were long-lived compared to monocytes. At the time, there was

limited knowledge of the development and function of macrophages in the embryo but the

first seeding of tissue macrophages from the embryonic circulation during development was

14

recognised within the MPS concept. The relative contribution of macrophage-specific

transcripts to total mRNA in most organs is established around the time of birth in mice [27].

Postnatal organ growth therefore creates many new territories to be filled either by

monocyte infiltration or by continued proliferation of the foetal liver-derived initial

residents. The creation of those new territories is itself partly CSF1R- and macrophage-

dependent as evidenced by the fact that the CSF1R knockout in both mice and rats has

severe postnatal growth retardation [58, 79]. Many of the recent studies discussed above

and in Box 1 show that blood monocytes can, and do, enter tissues to replace tissue

macrophages progressively in the postnatal period. In fact, no significant modification of

the MPS framework or reinterpretation of the original studies is required to include foetal

liver-derived monocytes as the initial precursors of tissue macrophages and to accept that

cells seeded in late gestation and early postnatal period may persist for months.

How often are resident macrophages replaced in an adult animal?

Although much emphasis has been placed upon resident macrophage proliferation and self-

renewal arguably the important parameter is the frequency of cell death amongst resident

macrophages. van Furth and Cohn recognised that extravasation of monocytes was a

probabilistic process and reported the half-life of monocytes [2]. Cell death is also a

probabilistic rather than a deterministic process [80] so cellular longevity is also best

described in terms of half-life. van Furth & Cohn [2] calculated the turnover time of 40 days

for peritoneal macrophages. However, if cell death is probabilistic turnover/replacement

will be governed by an exponential decay function and some cells will be very long-lived by

chance. Those cells will be exposed for a longer period to the local tissue environment. Like

the LY6Clo monocytes in the blood, they will have time to acquire more differentiated

functions in response to that environment. So, what happens when a resident macrophage

15

dies? One possibility is that a neighbouring macrophage divides in response to increased

local growth factor availability and one daughter cell migrates to occupy the vacant

territory. A second possibility is that the apoptotic macrophage produces a chemotactic

signal to attract a monocyte to engulf the dying cell. The infiltrating monocyte then

occupies the vacant territory and gradually responds to the local environment to acquire the

appropriate adaptation. Such a model is supported by the detailed analysis of targeted

ZEB2 deletion in different macrophage populations [28]. The reduced longevity of the

resident macrophages associated with loss of ZEB2 led to the loss of tissue-specific

macrophage adaptation and replacement of resident macrophages by marrow-derived cells.

Is the C57Bl/6 mouse a generalisable model of monocyte-macrophage homeostasis?

Macrophage turnover is clearly influenced by both genotype and environment. All of the

studies summarised in Table 1 and Box 1 have been carried out using inbred, specified

pathogen free (SPF), C57Bl/6J mice on a constant diet using individually ventilated cages.

They often focus on one sex, usually male because of the complex impact of the short

oestrous cycle in females on monocyte-macrophage turnover [8]. Is “the steady state”

described in the C57Bl/6 mouse model actually one of many mouse steady states with

distinct impacts on MPS homeostasis? The unique features of C57Bl/6 mice and relevant

variations amongst mouse strains are discussed in Box 3.

Transcription regulation and homeostasis in the MPS also differs between species and the

mouse is not a predictive model for larger animals including the rat [81, 82]. The

differences are not small. For example, the set of glucocorticoid (GC)-inducible genes in

mouse and human macrophages has a minimal overlap as a consequence of widespread

gain and loss of GC-responsive enhancers [83]. Rats [84], pigs [85, 86] and sheep [87] and of

course humans can each provide alternative models in which to study mammalian CSF1R

16

function, monocyte homeostasis and macrophage differentiation. We have recently

knocked out the CSF1R gene by homologous recombination in the rat genome [79]. The

CSF1R-/- rats are macrophage and osteoclast-deficient, but they are viable as adults and lack

many of the pleiotropic effects seen in mutant mice. Based upon these and many other

findings, it is reasonable to question whether precise details about MPS homeostasis in the

adult steady state, derived from studies of a single inbred mouse strain under laboratory

conditions, apply equally to other strains, to outbred male and female mice, or to other

species, living their normal life course in a changing environment.

Concluding Remarks

Ralph van Furth died early in 2018 in the 50th year since his development of the MPS

concept with Zanvil Cohn. In our opinion, the research in the 5 years since a previous

review of macrophage homeostasis [6] can be summarised by a well-known French

aphorism, “plus ça change, plus c'est la même chose” (the more things change, the more

they stay the same). The studies reviewed here remain consistent with the original MPS

lineage concept. Many questions remain (see outstanding questions), including the key

issue of how often macrophage territories become vacant and accessible to monocytes in

the face of real environmental challenges such as infection, injury, stress and ageing over

the life course of different species. To address these questions, there is a clear need to go

beyond inbred mouse models.

Acknowledgements

DAH and KMI are supported by The Mater Foundation. We thank David Langlais and Eyal

David for providing annotated chromatin data used in the drawing of Figure 1.

17

18

Legends to Figures

Figure 1. Complex transcriptional regulation of the mouse CSF1R locus

The mouse CSF1R locus contains multiple regulatory elements that are each in an open

chromatin conformation in macrophages and bound selectively by macrophage-specific

transcription factors. The Figure shows a diagrammatic representation of the mouse CSF1R

and neighbouring PDGFRB loci. Red arrows indicate the position of enhancers identified by

detection of open chromatin (Dnase hypersensitive sites (DHS), Assay for Transpose-

accessible chromatin (ATAC-Seq); Formaldehyde-assisted isolation of regulatory elements

(FIRE-Seq), histone modifications (H3K4me1, H3K27Ac) or active transcription of eRNA; see

Refs [3, 27, 68, 88]. The green arrow highlights the position of the Fms Intronic Regulatory

Element (FIRE) and the purple arrow is the major macrophage transcription start site. Chip-

Seq data for various transcription factor binding sites was integrated from [68] and other

published datasets (reviewed in [3]). The circles below highlights confirmed binding sites for

each of the named transcription factors, corresponding to the indicated enhancers.

Figure 2. The distribution of macrophages in mouse tissues

Representative images of the distribution of macrophages in diverse tissues of the CSF1R-

mApple mouse [62] taken by whole mount imaging using a spinning disc confocal

microscope. All images are the same magnification. The scale bar is 20 in all panels. The

low power perspective shows the remarkably similar density of macrophages (red) in

diverse organs and their regular spacing throughout the tissue regardless of underlying

structures indicating the lack of overlap of individual “territories”. Note the equally high

density of macrophages in liver and testis, and in several organs (abdominal wall, skeletal

muscle, salivary gland, trachea, bladder) that have not yet been widely studied. Asterisk in

19

brain meninges marks a vessel, in testis marks a seminiferous tubule (note cells spread on

the surface) and in liver the centrilobular region where Kupffer cells are excluded.

Key Figure, Figure 3. The life history of macrophage territories in mice

As tissues grow in during embryonic and postnatal development, new macrophage

territories are created. These territories are occupied by blood monocytes produced by the

fetal liver or bone marrow and/or by local macrophage proliferation. When organ growth

ceases, the rate of macrophage replacement is determined primarily by longevity.

Apoptotic cell death of resident macrophages produces both chemotactic signals for

monocytes and a local increase in CSF1 availability (red dots). Replacement may occur

through division and occupation of the territory by a neighbouring macrophage or by

monocytic infiltration. When a monocyte occupies a vacant territory it gradually acquires

the tissue-specific adaptations specific to its location.

20

TABLE 1

Recent highlights of identification of novel mouse tissue macrophage populations

Tissue Summary of key findings References

Lymph

node

CX3CR1+, MERTK+ macrophages of T cell zones, seeded in

utero but slowly replaced by monocytes. Function in

clearance of apoptotic T cells.

[4]

Skin Dermal macrophage population, including cells that

internalise melanosomes. Identified the population that

internalises tattoo pigment particles, and the ability of

incoming monocyte-derived macrophages to recapture

particles released from dying cells.

[89]

Peripheral

nerves

Macrophage population associated with sympathetic

nervous system, mediates clearance of norepinephrine and

contributes to regulation of thermogenesis.

[90]

Artery CX3CR1-dependent population of macrophages in the

artery wall with distinct gene expression profile; seeded

embryonically and postnatally from blood monocytes.

Express LYVE-1 and regulate arterial tone

[91, 92]

Pancreas Distinct macrophage populations identified in islets of

Langerhans and inter-acinar stroma. Mixtures of cells

derived from foetal seeding and definitive hematopoiesis.

Contribute to differentiation of pancreatic beta cells.

[93] [94]

Testis Two populations of macrophages described in the testis, [95]

21

interstitial and peritubular, with distinct developmental

origins

Bone

marrow

Radiation resistant, self-repopulating hematopoietic island

macrophages essential for optimal engraftment in bone

marrow transplantation

[96]

Bone CD169+ macrophages of the bone surface essential for

osteoblast maintenance and regulation of ossification.

[97]

Gut Distinct muscularis and submucosal macrophages regulate

vascular and enteric neuronal function and gastrointestinal

motility. Population of macrophages associated with

intestinal crypts regulates Paneth cells and stem cell

fate/differentiation

[98] [37, 40]

Kidney Macrophage function and location (concentration in

medulla) is regulated dynamically by the renal salt

gradient. Renal medullary macrophages interact intimately

with peritubular capillary endothelial cells.

[74, 99]

Brown

adipose

Macrophages of brown adipose regulate sympathetic

innervation and thermogenesis

[100]

Text Boxes

22

Box 1. Turnover of Tissue Resident Macrophages in Mice.

The past 3 years has seen a deluge of publications on the developmental origins and

turnover of tissue macrophages in mice. They include studies of macrophages of the

peritoneum [31, 101], skin [89], lymph node [4], intestinal submucosa and muscularis [38,

40], pancreas [93], artery [91], kidney [74, 102, 103], heart [14, 57] and testis [95]. The

conclusion of one study [31] that large F4/80hi peritoneal macrophages are slowly replaced

through differentiation of monocyte-derived F4/80lo/MHCII+ progenitors is consistent with

the original study of van Furth and Cohn [2]. A separate study [101] suggested that the

F4/80lo monocyte-derived peritoneal macrophages are continuously replenished, have a

unique requirement for the transcription factor IRF4. Dermal macrophages, like those of

the gut, are believed to be continuously replaced from blood monocytes [89]. A novel

dermal macrophage population accumulates melanin granules (melanophages) and can be

reconstituted from monocyte sources following ablation [89]. Incoming monocyte-derived

dermal macrophages were found to recapture tattoo ink particles locally released following

induced macrophage ablation [89]. Another example of clinical interest is the specialised

macrophages that form the centre of hematopoietic islands of the bone marrow. These

cells are capable of self-renewal even in the context of irradiation and bone marrow

transplantation in mouse models, and are essential for successful engraftment [96]. One of

the archetypal functions of macrophages, mediated by a complex array of receptors

including MERTK and TIM4, is the recognition and clearance of apoptotic cells. Two recent

studies have focused on novel populations of macrophages engaged with T cell

efferocytosis. One study [104] identified F4/80+/TIM4+ macrophages in the thymus and

demonstrated their dependence (in common with Ly6Clo monocytes) on the transcription

factor NR4A1. Several lines of evidence suggested that these cells were maintained

23

primarily by self-renewal. By contrast, a newly recognised and abundant population of

macrophages (T cell zone macrophages, or TZM) in T cell areas of mouse lymph node and

Peyer’s patch (and likely also spleen) lacked expression of the F4/80 marker and appeared

to be replaced slowly by blood monocytes [4]. One macrophage niche in the gut wall that

has not been studied in detail is the isolated lymphoid follicles (ILF) [105]. These are more

abundant than commonly-recognised; their macrophage populations are so numerous that

they can be identified in an en face view of the mouse intestine with a macrophage reporter

transgene [71]. At least some of the submucosal TIM4+ intestinal macrophages discussed

might be engaged in efferocytosis of lymphocytes in ILF.

Box 2: Lineage-trace models of macrophage self-renewal

The conclusion that many tissue macrophages are maintained by self-renewal,

independently of blood monocytes, relies in part upon the interpretation of lineage tracers

in inbred reporter mice, in which precursor cells can be pulse-labelled in the embryo or the

adult. Different promoters used to drive constitutive or tamoxifen-inducible cre-

recombinase, linked to different Cre-dependent reporter genes (CSF1R, S100A4, FLT3,

CX3CR1, RUNX1, KIT, TNFRSF11A) have produced different outcomes, interpreted in

different ways to support distinct roles for the yolk sac-derived EMP and foetal monocytes

versus HSC-derived foetal or adult monocytes. The interpretation of lineage traces has been

discussed by others [7, 106-109]. These studies have been reviewed critically previously

specifically questioning the widespread use of tamoxifen-inducible lineage tracers to study

macrophage homeostasis [6]. Tamoxifen is not a neutral agonist. In our hands, maternal

tamoxifen treatment produced >4-fold increase in tissue macrophage numbers in the

11.5dpc embryo [110] which could bias the relative contribution of embryonic macrophages

24

to the postnatal pool. Aside from the study of microglia noted above [50], there is limited

and sometimes contradictory evidence of resident macrophage self-renewal by cell division

in adults. Epelman et al [14] reported around 1-2% labelling of the different cardiac

macrophage populations with bromodeoxyuridine. On the other hand, Molawai et al. [57]

provided evidence that self-renewal declines rapidly with age and resident macrophages are

progressively replaced by monocyte-derived cells.

Box 3: Is the male C57Bl/6 mouse model representative of monocyte homeostasis?

There are many caveats to the study of male C57Bl/6 mice in an SPF animal facility as a

model for monocyte-macrophage homeostasis. Male and female mice are clearly different

in their macrophage homeostasis. Macrophage proliferation and self-renewal is sensitive to

estrogen [111] which brings further into question the widespread use of the estrogen

receptor agonist tamoxifen in inducible lineage trace studies (e.g. [17, 40]). Thion et al [73]

described transcriptomic differences in microglia of male and female mice, and differential

impacts of the microbiome on microglial phenotype in each sex. There is emerging evidence

that commonly-used housing conditions produce chronic temperature and social stress in

laboratory mice [112]. Williams et al [113] reported direct effects of cold stress versus

thermoneutrality on monocyte retention in the bone marrow.

All of the original studies that led to the MPS concept and its subsequent development were

performed on Swiss outbred mice and there was substantial variation between animals

[114]. Blood leukocyte numbers vary by an order of magnitude between mouse strains.

C57Bl/6 mice, for example, are notably granulocyte-deficient [115]. Macrophage gene

expression and function certainly differs between inbred mouse strains each of which is

homozygous for various loss of function alleles. Those differences relate to monocyte-

macrophage homeostasis. For example, female C57Bl/6 mice develop spontaneous

25

osteoporosis associated with dysregulated osteoclastogenesis that can be reverted with

anti-CSF1R treatment [116]. Macrophages from C57Bl/6 mice lack expression of cathepsin

E, which influences cell migration, and express C1Q components and IFN target genes

(attributed functions in microglia, [73]) at very high levels relative to other strains [117].

Campbell et al. [118] recently compared the response of C57Bl/6 and BALB/c mice to

parasitic nematode infection. Their previous work had demonstrated that in C57Bl/6 mice

the Th2 lymphokine, IL4, promotes rapid proliferation of resident macrophages in this

infection model whereas monocyte recruitment was minimal [63]. In their recent study, the

response of BALB/c mice was instead dominated by monocyte recruitment. Even in the

unstimulated state, the F4/80lo monocyte-derived, peritoneal macrophage population was

much more abundant in the BALB/c than the C57Bl/6 mice. The clear implication is that

monocyte and macrophage homeostasis is mouse strain-dependent and conclusions based

upon any single mouse strain are not generalisable.

26

27

Reference

1. van Furth, R. et al. (1972) The mononuclear phagocyte system: a new classification of

macrophages, monocytes, and their precursor cells. Bull World Health Organ 46 (6), 845-52.

2. van Furth, R. and Cohn, Z.A. (1968) The origin and kinetics of mononuclear phagocytes. J

Exp Med 128 (3), 415-35.

3. Rojo, R. et al. (2017) Transcriptional mechanisms that control expression of the

macrophage colony-stimulating factor receptor locus. Clin Sci (Lond) 131 (16), 2161-2182.

4. Baratin, M. et al. (2017) T Cell Zone Resident Macrophages Silently Dispose of Apoptotic

Cells in the Lymph Node. Immunity 47 (2), 349-362 e5.

5. Guilliams, M. et al. (2014) Dendritic cells, monocytes and macrophages: a unified

nomenclature based on ontogeny. Nat Rev Immunol 14 (8), 571-8.

6. Jenkins, S.J. and Hume, D.A. (2014) Homeostasis in the mononuclear phagocyte system.

Trends Immunol 35 (8), 358-67.

7. Hoeffel, G. and Ginhoux, F. (2018) Fetal monocytes and the origins of tissue-resident

macrophages. Cell Immunol 330, 5-15.

8. Stremmel, C. et al. (2018) Yolk sac macrophage progenitors traffic to the embryo during

defined stages of development. Nat Commun 9 (1), 75.

9. Balic, A. et al. (2014) Visualisation of chicken macrophages using transgenic reporter

genes: insights into the development of the avian macrophage lineage. Development 141

(16), 3255-65.

10. Garceau, V. et al. (2015) The development and maintenance of the mononuclear

phagocyte system of the chick is controlled by signals from the macrophage colony-

stimulating factor receptor. BMC Biol 13, 12.

28

11. Schulz, C. et al. (2012) A lineage of myeloid cells independent of Myb and hematopoietic

stem cells. Science 336 (6077), 86-90.

12. Gomez Perdiguero, E. et al. (2015) Tissue-resident macrophages originate from yolk-sac-

derived erythro-myeloid progenitors. Nature 518 (7540), 547-51.

13. Hoeffel, G. et al. (2015) C-Myb(+) erythro-myeloid progenitor-derived fetal monocytes

give rise to adult tissue-resident macrophages. Immunity 42 (4), 665-78.

14. Epelman, S. et al. (2014) Embryonic and adult-derived resident cardiac macrophages are

maintained through distinct mechanisms at steady state and during inflammation. Immunity

40 (1), 91-104.

15. Hettinger, J. et al. (2013) Origin of monocytes and macrophages in a committed

progenitor. Nat Immunol 14 (8), 821-30.

16. Mildner, A. et al. (2016) Murine Monocytes: Origins, Subsets, Fates, and Functions.

Microbiol Spectr 4 (5).

17. Yona, S. et al. (2013) Fate mapping reveals origins and dynamics of monocytes and tissue

macrophages under homeostasis. Immunity 38 (1), 79-91.

18. Patel, A.A. et al. (2017) The fate and lifespan of human monocyte subsets in steady state

and systemic inflammation. J Exp Med 214 (7), 1913-1923.

19. Mildner, A. et al. (2017) Genomic Characterization of Murine Monocytes Reveals

C/EBPbeta Transcription Factor Dependence of Ly6C(-) Cells. Immunity 46 (5), 849-862 e7.

20. Schmidl, C. et al. (2014) Transcription and enhancer profiling in human monocyte

subsets. Blood 123 (17), e90-9.

21. Thomas, G.D. et al. (2016) Deleting an Nr4a1 Super-Enhancer Subdomain Ablates

Ly6C(low) Monocytes while Preserving Macrophage Gene Function. Immunity 45 (5), 975-

987.

29

22. Briseno, C.G. et al. (2016) Distinct Transcriptional Programs Control Cross-Priming in

Classical and Monocyte-Derived Dendritic Cells. Cell Rep 15 (11), 2462-74.

23. Menezes, S. et al. (2016) The Heterogeneity of Ly6C(hi) Monocytes Controls Their

Differentiation into iNOS(+) Macrophages or Monocyte-Derived Dendritic Cells. Immunity 45

(6), 1205-1218.

24. Yanez, A. et al. (2017) Granulocyte-Monocyte Progenitors and Monocyte-Dendritic Cell

Progenitors Independently Produce Functionally Distinct Monocytes. Immunity 47 (5), 890-

902 e4.

25. Biermann, H. et al. (1999) Murine leukocytes with ring-shaped nuclei include

granulocytes, monocytes, and their precursors. J Leukoc Biol 65 (2), 217-31.

26. Mass, E. et al. (2016) Specification of tissue-resident macrophages during organogenesis.

Science 353 (6304). aaf4238

27. Summers, K.M. and Hume, D.A. (2017) Identification of the macrophage-specific

promoter signature in FANTOM5 mouse embryo developmental time course data. J Leukoc

Biol 102 (4), 1081-1092.

28. Scott, C.L. et al. (2018) The Transcription Factor ZEB2 Is Required to Maintain the Tissue-

Specific Identities of Macrophages. Immunity 49 (2), 312-325 e5.

29. Gordon, S. and Pluddemann, A. (2017) Tissue macrophages: heterogeneity and

functions. BMC Biol 15 (1), 53.

30. Bain, C.C. et al. (2014) Constant replenishment from circulating monocytes maintains the

macrophage pool in the intestine of adult mice. Nat Immunol 15 (10), 929-937.

31. Bain, C.C. et al. (2016) Long-lived self-renewing bone marrow-derived macrophages

displace embryo-derived cells to inhabit adult serous cavities. Nat Commun 7,

ncomms11852.

30

32. Matcovitch-Natan, O. et al. (2016) Microglia development follows a stepwise program to

regulate brain homeostasis. Science 353 (6301), aad8670.

33. Guilliams, M. and Scott, C.L. (2017) Does niche competition determine the origin of

tissue-resident macrophages? Nat Rev Immunol 17 (7), 451-460.

34. Mowat, A.M. et al. (2017) Barrier-tissue macrophages: functional adaptation to

environmental challenges. Nat Med 23 (11), 1258-1270.

35. Schridde, A. et al. (2017) Tissue-specific differentiation of colonic macrophages requires

TGFbeta receptor-mediated signaling. Mucosal Immunol 10 (6), 1387-1399.

36. Asano, K. et al. (2015) Intestinal CD169(+) macrophages initiate mucosal inflammation

by secreting CCL8 that recruits inflammatory monocytes. Nat Commun 6, 7802.

37. Sehgal, A. et al. (2018) The role of CSF1R-dependent macrophages in control of the

intestinal stem-cell niche. Nat Commun 9 (1), 1272.

38. Shaw, T.N. et al. (2018) Tissue-resident macrophages in the intestine are long lived and

defined by Tim-4 and CD4 expression. J Exp Med 215 (6), 1507-1518.

39. Bonnardel, J. et al. (2015) Innate and adaptive immune functions of peyer's patch

monocyte-derived cells. Cell Rep 11 (5), 770-84.

40. De Schepper, S. et al. (2018) Self-Maintaining Gut Macrophages Are Essential for

Intestinal Homeostasis. Cell.175 (2), 400-415

41. Sierro, F. et al. (2017) A Liver Capsular Network of Monocyte-Derived Macrophages

Restricts Hepatic Dissemination of Intraperitoneal Bacteria by Neutrophil Recruitment.

Immunity 47 (2), 374-388 e6.

42. Beattie, L. et al. (2016) Bone marrow-derived and resident liver macrophages display

unique transcriptomic signatures but similar biological functions. J Hepatol 65 (4), 758-768.

31

43. David, B.A. et al. (2016) Combination of Mass Cytometry and Imaging Analysis Reveals

Origin, Location, and Functional Repopulation of Liver Myeloid Cells in Mice.

Gastroenterology 151 (6), 1176-1191.

44. Scott, C.L. et al. (2016) Bone marrow-derived monocytes give rise to self-renewing and

fully differentiated Kupffer cells. Nat Commun 7, 10321.

45. Cohen, M. et al. (2018) Lung Single-Cell Signaling Interaction Map Reveals Basophil Role

in Macrophage Imprinting. Cell 175 (4), 1031-1044.e18.

46. Garbi, N. and Lambrecht, B.N. (2017) Location, function, and ontogeny of pulmonary

macrophages during the steady state. Pflugers Arch 469 (3-4), 561-572.

47. Gibbings, S.L. et al. (2017) Three Unique Interstitial Macrophages in the Murine Lung at

Steady State. Am J Respir Cell Mol Biol 57 (1), 66-76.

48. Tan, S.Y. and Krasnow, M.A. (2016) Developmental origin of lung macrophage diversity.

Development 143 (8), 1318-27.

49. van de Laar, L. et al. (2016) Yolk Sac Macrophages, Fetal Liver, and Adult Monocytes Can

Colonize an Empty Niche and Develop into Functional Tissue-Resident Macrophages.

Immunity 44 (4), 755-68.

50. Askew, K. et al. (2017) Coupled Proliferation and Apoptosis Maintain the Rapid Turnover

of Microglia in the Adult Brain. Cell Rep 18 (2), 391-405.

51. Perry, V.H. et al. (1985) Immunohistochemical localization of macrophages and microglia

in the adult and developing mouse brain. Neuroscience 15 (2), 313-26.

52. Goldmann, T. et al. (2016) Origin, fate and dynamics of macrophages at central nervous

system interfaces. Nat Immunol 17 (7), 797-805.

53. Mildner, A. et al. (2007) Microglia in the adult brain arise from Ly-6ChiCCR2+ monocytes

only under defined host conditions. Nat Neurosci 10 (12), 1544-53.

32

54. Beers, D.R. et al. (2006) Wild-type microglia extend survival in PU.1 knockout mice with

familial amyotrophic lateral sclerosis. Proc Natl Acad Sci U S A 103 (43), 16021-6.

55. Bennett, F.C. et al. (2018) A Combination of Ontogeny and CNS Environment Establishes

Microglial Identity. Neuron 98 (6), 1170-1183 e8.

56. Cronk, J.C. et al. (2018) Peripherally derived macrophages can engraft the brain

independent of irradiation and maintain an identity distinct from microglia. J Exp Med 215

(6), 1627-1647.

57. Molawi, K. et al. (2014) Progressive replacement of embryo-derived cardiac

macrophages with age. J Exp Med 211 (11), 2151-8.

58. Chitu, V. and Stanley, E.R. (2017) Regulation of Embryonic and Postnatal Development

by the CSF-1 Receptor. Curr Top Dev Biol 123, 229-275.

59. Lichanska, A.M. et al. (1999) Differentiation of the mononuclear phagocyte system

during mouse embryogenesis: the role of transcription factor PU.1. Blood 94 (1), 127-38.

60. MacDonald, K.P. et al. (2010) 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 116 (19), 3955-63.

61. Gow, D.J. et al. (2014) Characterisation of a novel Fc conjugate of macrophage colony-

stimulating factor. Mol Ther 22 (9), 1580-92.

62. Hawley, C.A. et al. (2018) Csf1r-mApple Transgene Expression and Ligand Binding In Vivo

Reveal Dynamics of CSF1R Expression within the Mononuclear Phagocyte System. J Immunol

200 (6), 2209-2223.

63. Jenkins, S.J. et al. (2013) IL-4 directly signals tissue-resident macrophages to proliferate

beyond homeostatic levels controlled by CSF-1. J Exp Med 210 (11), 2477-91.

33

64. Stutchfield, B.M. et al. (2015) CSF1 Restores Innate Immunity After Liver Injury in Mice

and Serum Levels Indicate Outcomes of Patients With Acute Liver Failure. Gastroenterology

149 (7), 1896-1909 e14.

65. Perie, L. and Naik, S.H. (2015) Toward defining a 'lineage'--The case for dendritic cells.

Semin Cell Dev Biol 41, 3-8.

66. Durai, V. et al. (2018) Altered compensatory cytokine signaling underlies the discrepancy

between Flt3(-/-) and Flt3l(-/-) mice. J Exp Med 215 (5), 1417-1435.

67. Nandi, S. et al. (2006) Developmental and functional significance of the CSF-1

proteoglycan chondroitin sulfate chain. Blood 107 (2), 786-95.

68. Langlais, D. et al. (2016) The macrophage IRF8/IRF1 regulome is required for protection

against infections and is associated with chronic inflammation. J Exp Med 213 (4), 585-603.

69. Lavine, K.J. et al. (2014) Distinct macrophage lineages contribute to disparate patterns of

cardiac recovery and remodeling in the neonatal and adult heart. Proc Natl Acad Sci U S A

111 (45), 16029-34.

70. Sasmono, R.T. et al. (2003) A macrophage colony-stimulating factor receptor-green

fluorescent protein transgene is expressed throughout the mononuclear phagocyte system

of the mouse. Blood 101 (3), 1155-63.

71. Sauter, K.A. et al. (2014) The MacBlue binary transgene (csf1r-gal4VP16/UAS-ECFP)

provides a novel marker for visualisation of subsets of monocytes, macrophages and

dendritic cells and responsiveness to CSF1 administration. PLoS One 9 (8), e105429.

72. Rojo, R. et al. (2018) Germ-line deletion of a conserved macrophage-specific enhancer

within the mouse Csf1r locus has a selective impact on tissue macrophage populations

including microglia. Nature Immunology In press.

34

73. Thion, M.S. et al. (2018) Microbiome Influences Prenatal and Adult Microglia in a Sex-

Specific Manner. Cell 172 (3), 500-516 e16.

74. Stamatiades, E.G. et al. (2016) Immune Monitoring of Trans-endothelial Transport by

Kidney-Resident Macrophages. Cell 166 (4), 991-1003.

75. Avetisyan, M. et al. (2018) Muscularis macrophage development in the absence of an

enteric nervous system. Proc Natl Acad Sci U S A 115 (18), 4696-4701.

76. Nimmerjahn, A. et al. (2005) Resting microglial cells are highly dynamic surveillants of

brain parenchyma in vivo. Science 308 (5726), 1314-8.

77. Poche, R.A. et al. (2015) Macrophages engulf endothelial cell membrane particles

preceding pupillary membrane capillary regression. Dev Biol 403 (1), 30-42.

78. Wood, W. and Martin, P. (2017) Macrophage Functions in Tissue Patterning and Disease:

New Insights from the Fly. Dev Cell 40 (3), 221-233.

79. Pridans, C. et al. (2018) Targetted disruption of the rat Csf1r locus generates viable adult

animals with selective macrophage and microglial deficiencies. J.Immunol. 201(9), 2683-

2699.

80. Hodgkin, P.D. (2018) Modifying clonal selection theory with a probabilistic cell. Immunol

Rev 285 (1), 249-262.

81. Hagai, T. et al. (2018) Gene expression variability across cells and species shapes innate

immunity. Nature. 563(7730), 197-202

82. Schroder, K. et al. (2012) Conservation and divergence in Toll-like receptor 4-regulated

gene expression in primary human versus mouse macrophages. Proc Natl Acad Sci U S A 109

(16), E944-53.

35

83. Jubb, A.W. et al. (2016) Enhancer Turnover Is Associated with a Divergent

Transcriptional Response to Glucocorticoid in Mouse and Human Macrophages. J Immunol

196 (2), 813-822.

84. Pridans, C. et al. (2018) Macrophage colony-stimulating factor increases hepatic

macrophage content, liver growth, and lipid accumulation in neonatal rats. Am J Physiol

Gastrointest Liver Physiol 314 (3), G388-G398.

85. Kapetanovic, R. et al. (2013) The impact of breed and tissue compartment on the

response of pig macrophages to lipopolysaccharide. BMC Genomics 14, 581.

86. Sauter, K.A. et al. (2016) Macrophage colony-stimulating factor (CSF1) controls

monocyte production and maturation and the steady-state size of the liver in pigs. Am J

Physiol Gastrointest Liver Physiol 311 (3), G533-47.

87. Pridans, C. et al. (2016) A Csf1r-EGFP Transgene Provides a Novel Marker for Monocyte

Subsets in Sheep. J Immunol 197 (6), 2297-305.

88. Lavin, Y. et al. (2014) Tissue-resident macrophage enhancer landscapes are shaped by

the local microenvironment. Cell 159 (6), 1312-26.

89. Baranska, A. et al. (2018) Unveiling skin macrophage dynamics explains both tattoo

persistence and strenuous removal. J Exp Med 215 (4), 1115-1133.

90. Pirzgalska, R.M. et al. (2017) Sympathetic neuron-associated macrophages contribute to

obesity by importing and metabolizing norepinephrine. Nat Med 23 (11), 1309-1318.

91. Ensan, S. et al. (2016) Self-renewing resident arterial macrophages arise from embryonic

CX3CR1(+) precursors and circulating monocytes immediately after birth. Nat Immunol 17

(2), 159-68.

36

92. Lim, H.Y. et al. (2018) Hyaluronan Receptor LYVE-1-Expressing Macrophages Maintain

Arterial Tone through Hyaluronan-Mediated Regulation of Smooth Muscle Cell Collagen.

Immunity 49 (2), 326-341 e7.

93. Calderon, B. et al. (2015) The pancreas anatomy conditions the origin and properties of

resident macrophages. J Exp Med 212 (10), 1497-512.

94. Mussar, K. et al. (2017) A CCR2+ myeloid cell niche required for pancreatic beta cell

growth. JCI Insight 2 (15). 93834

95. Mossadegh-Keller, N. et al. (2017) Developmental origin and maintenance of distinct

testicular macrophage populations. J Exp Med 214 (10), 2829-2841.

96. Kaur, S. et al. (2018) Self-repopulating recipient bone marrow resident macrophages

promote long-term hematopoietic stem cell engraftment. Blood 132 (7), 735-749.

97. Batoon, L. et al. (2017) CD169(+) macrophages are critical for osteoblast maintenance

and promote intramembranous and endochondral ossification during bone repair.

Biomaterials. pii S014209612(17)30682-8

98. Gabanyi, I. et al. (2016) Neuro-immune Interactions Drive Tissue Programming in

Intestinal Macrophages. Cell 164 (3), 378-91.

99. Berry, M.R. et al. (2017) Renal Sodium Gradient Orchestrates a Dynamic Antibacterial

Defense Zone. Cell 170 (5), 860-874 e19.

100. Wolf, Y. et al. (2017) Brown-adipose-tissue macrophages control tissue innervation and

homeostatic energy expenditure. Nat Immunol 18 (6), 665-674.

101. Kim, K.W. et al. (2016) MHC II+ resident peritoneal and pleural macrophages rely on

IRF4 for development from circulating monocytes. J Exp Med 213 (10), 1951-9.

102. Lever, J.M. et al. (2018) Parabiosis reveals leukocyte dynamics in the kidney. Lab Invest

98 (3), 391-402.

37

103. Munro, D.A.D. and Hughes, J. (2017) The Origins and Functions of Tissue-Resident

Macrophages in Kidney Development. Front Physiol 8, 837.

104. Tacke, R. et al. (2015) The transcription factor NR4A1 is essential for the development

of a novel macrophage subset in the thymus. Sci Rep 5, 10055.

105. Donaldson, D.S. et al. (2015) Reciprocal regulation of lymphoid tissue development in

the large intestine by IL-25 and IL-23. Mucosal Immunol 8 (3), 582-95.

106. Epelman, S. et al. (2014) Origin and functions of tissue macrophages. Immunity 41 (1),

21-35.

107. Jung, S. (2018) Macrophages and monocytes in 2017: Macrophages and monocytes: of

tortoises and hares. Nat Rev Immunol 18 (2), 85-86.

108. Perdiguero, E.G. and Geissmann, F. (2016) The development and maintenance of

resident macrophages. Nat Immunol 17 (1), 2-8.

109. Varol, C. et al. (2015) Macrophages: development and tissue specialization. Annu Rev

Immunol 33, 643-75.

110. Rojo, R. et al. (2018) Maternal tamoxifen treatment expands the macrophage

population of early mouse embryos. . BioRxiv, 296749.

111. Pepe, G. et al. (2017) Self-renewal and phenotypic conversion are the main

physiological responses of macrophages to the endogenous estrogen surge. Sci Rep 7,

44270.

112. Gaskill, B.N. and Garner, J.P. (2017) Stressed out: providing laboratory animals with

behavioral control to reduce the physiological effects of stress. Lab Anim (NY) 46 (4), 142-

145.

113. Williams, J.W. et al. (2017) Thermoneutrality but Not UCP1 Deficiency Suppresses

Monocyte Mobilization Into Blood. Circ Res 121 (6), 662-676.

38

114. Crofton, R.W. et al. (1978) The origin, kinetics, and characteristics of the Kupffer cells in

the normal steady state. J Exp Med 148 (1), 1-17.

115. Petkova, S.B. et al. (2008) Genetic influence on immune phenotype revealed strain-

specific variations in peripheral blood lineages. Physiol Genomics 34 (3), 304-14.

116. Sauter, K.A. et al. (2014) Pleiotropic effects of extended blockade of CSF1R signaling in

adult mice. J Leukoc Biol 96 (2), 265-74.

117. Raza, S. et al. (2014) Analysis of the transcriptional networks underpinning the

activation of murine macrophages by inflammatory mediators. J Leukoc Biol 96 (2), 167-83.

118. Campbell, S.M. et al. (2018) Myeloid cell recruitment versus local proliferation

differentiates susceptibility from resistance to filarial infection. Elife 7, e30947

39