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Originally published online as doi:10.1189/jlb.0307191 on May 2, 2007

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(Journal of Leukocyte Biology. 2007;82:244-252.)
© 2007 by Society for Leukocyte Biology

Monocyte subpopulations and their differentiation patterns during infection

Dalit Strauss-Ayali, Sean M. Conrad and David M. Mosser1

Department of Cell Biology and Molecular Genetics, University of Maryland, College Park, Maryland, USA

1 Correspondence: Department of Cell Biology and Molecular Genetics, University of Maryland, College Park, MD 20742, USA. E-mail: dmosser{at}umd.edu

ABSTRACT

The term "monocyte" implies a single, homogenous population of cells with uniform physiology. Recent evidence from a number of laboratories indicates that it is likely that blood monocytes may consist of several subpopulations of cells, which differ in size, nuclear morphology, granularity, and functionality. The aim of this review is to give a summary of the new findings in the emerging field of monocyte heterogeneity. We provide a short description of the differentiation patterns of blood monocyte subpopulations, with an emphasis on how these subpopulations can be influenced by infection. We provide a comparison among the main monocyte subpopulations in humans, mice, and rats and illustrate some of the common features of these cells and some of the important interspecies distinctions. We will also discuss the bone marrow precursors of these cells and the differentiation patterns of these subsets in different tissues in response to infection. Most of the data about monocyte trafficking during infection are necessarily derived from murine models, and comparisons between mouse and man must be made with caution. However, these models may provide interesting springboards to permit us to speculate about the topic of monocyte heterogeneity in humans.

Key Words: macrophages • chemokine receptors • GR1 • inflammation

MONOCYTE SUBPOPULATIONS

It has been almost 20 years since the existence of different monocyte subpopulations in humans was initially proposed [1 , 2 ]. Human monocytes, which were until then, identified only by the expression of CD14, could be classified further, based on differential expression levels of CD14 and CD16 (Fc{gamma}RIII). The CD14hiCD16– cells were described originally as "classical" monocytes as a result of the fact that they were 90–95% of the total monocytes in a healthy person. The terms "nonclassical" or "proinflammatory" monocytes were associated with the Fc{gamma}RIII-positive (CD14+CD16+) subpopulation, which produced lower levels of IL-10 in response to a TLR4 agonist and higher amounts of TNF-{alpha} protein in response to TLR4 and TLR2 agonists [3 4 5 ]. In a healthy adult, this subpopulation consists of 5–10% of the total monocytes in the blood. In a model of transendothelial trafficking, these monocytes were subsequently shown to be able to differentiate into dendritic cells (DCs), suggesting that they might represent the physiologically relevant blood precursors of migrating DCs [6 ]. Further studies show that monocyte subpopulations also differ by means of other surface molecules (Fig. 1 and see Table 1 ). Among them are the chemokine receptors CCR5 and CX3CR1, which are highly expressed on the CD14+CD16+ cells, while CCR2 and lower levels of CX3CR1 are characteristic of the (classical) CD14hiCD16– cells [7 8 9 ]. The expression of specific chemokine receptors and adhesion molecules might have important, functional consequences for these cells, resulting not only in differences in migration patterns [7 , 9 ] but also conferring differential susceptibility to infections [10 ]. For example, organisms such as HIV and West Nile virus, which can exploit these types of receptors to establish infections, may therefore infect specific monocyte subpopulations preferentially [11 , 12 ].


Figure 1
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  		Figure 1. Schematic representation of the main monocyte subpopulations in the human, mouse, and rat. Monocytes from human, mouse, and rat are diagrammed schematically, showing the expression of surface markers on each. Note differences in size and granularity on the various cell populations. The expression patterns of the chemokine receptors CCR2 and CX3CR1, as well as of the adhesion molecule CD62 ligand (CD62L), are depicted on these monocyte subpopulations. (Note: Gr1 consists of two epitopes, Ly6C and LybG; of these, Ly6C is unique to monocytes.)


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  		Table 1. Cell Surface Marker Expression on the Two Major Monocyte Subpopulations in Mice, Humans, and Ratsa

Mouse blood monocytes are identified primarily by their expression of CD115 (M-CSF receptor), CD11b [membrane-activating complex 1 (Mac-1)], and the F4/80 antigen. Only recently have murine monocyte subpopulations been found to share striking similarities to the subpopulations described previously in humans [8 ]. In the mouse, one population of monocytes was found to express intermediate levels of CX3CR1 and high levels of granulocyte antigen 1 (Gr1). These cells also express CCR2. This population of murine monocytes (CX3CR1+CCR2+Gr1hi) shares morphological characteristics and chemokine receptor expression patterns with the classical human CD14hiCD16– (Fig. 1) . However, because of their rapid recruitment to areas of damage or inflamed tissues, this population of murine monocytes was called "inflammatory" monocytes [8 ]. Thus, although these two populations share morphological characteristics, current studies suggest that they may have different functions. This difference has led to some confusion in extrapolating information from the murine system into the human system and vice versa. The other main mouse monocyte subpopulation was found to express high levels of CX3CR1 and low levels of Gr1 and CCR2. These monocytes appear to be orthologs of the human CD14+CD16+ counterparts described above (Fig. 1) . Murine CX3CR1hiCCR2–Gr1low monocytes and human CD14+CD16+ (CX3CR1hiCCR2–) monocytes were found to be smaller in size and less granular. In the mouse, the CX3CR1hiCCR2–Gr1low cells tended to persist longer in blood and normal tissues [8 ]. For this reason, these monocyte subpopulations have also been referred to as resident monocytes [8 ].

After the depletion of mouse monocytes using clodronate liposomes, the first monocytes that appear in the blood are the CX3CR1+CCR2+Gr1hi (Ly6Chi) [13 ]. It was therefore suggested that these cells are the precursors of the CX3CR1hiCCR2–Gr1low(Ly6Clow) cells, possibly through an intermediate, Gr1-expressing, monocyte subpopulation. It was suggested that Gr1 expression might be lost as the cells mature [13 ].

Two subpopulations of monocytes have also been identified in the rat. Subsets have been identified on the basis of chemokine receptor expression and CD43 levels [14 ] (Fig. 1 and see Table 1 ). Rat monocytes, which were CD43hi, were also CX3CR1+CD11c+ and CCR2–, and the rat CD43low monocytes were CX3CR1lowCD11c– but were CCR2+. The latter CD43lowCX3CR1lowCCR2+ monocytes were larger and were recruited to inflamed tissues, whereas the CD43hiCX3CR1+CCR2–monocytes were not recruited to these sites. Using this rat model, it was further shown that under steady-state conditions, the CD43lowCX3CR1lowCCR2+ cells differentiate into CD43hiCX3CR1+CCR2– without cell division [14 ]. Therefore, on the basis of size and chemokine receptor expression, the CD43hi cells resemble CX3CR1hiCCR2–Gr1low in mice and CD14+CD16+ in humans, whereas the CD43low cells resemble CX3CR1+CCR2+Gr1hi and CD14hiCD16– in mouse and human, respectively.

Thus, with regard to a number of criteria, including size, granularity, chemokine receptor expression, and function, two populations of human, murine, and rat monocytes can be readily delineated (Fig. 1 and Table 1 ). Nevertheless, there are also some clear distinctions, which are awaiting resolution. For example, unlike the human situation, where the vast majority (~90%) of the monocytes are of the classical phenotype (CD14hiCD16–), the two major mouse subpopulations are generally, equally represented in blood, and the nonclassical CD43hiCX3CR1+CCR2– monocytes are the bigger population (80–90%) in rat blood. Moreover, the in vitro proinflammatory phenotype of the human CD14+CD16+ cells has not been confirmed yet for their mouse and rat counterparts. The CD16+ human monocyte subpopulation may also have better antigen presentation ability than the mouse and rat subpopulations.

It is reasonable to suspect that these two monocyte populations may represent the "tip of the iceberg." As we develop new markers and better analytical techniques, it may well be that these populations can be subdivided further. For example, human monocytes have also been classified based on their CD64 (Fc{gamma}RI) expression [16 ], as well as on expression of the adhesion molecule CD56 [20 , 21 ]. An additional minor murine subpopulation was also characterized by the intermediate expression of Gr1 and the expression of the chemokine receptors CCR7 and CCR8 [13 , 22 ]. This subpopulation was suggested to be most predisposed to become lymphatic-migrating DCs [22 ].

BLOOD

A considerable increase in the number of the CD14+CD16+ monocytes had been described for a variety of systemic, infectious agents in humans, including hemolytic uremic syndrome, bacterial sepsis, HIV, and experimental SIV infections in monkeys. All of these infections are accompanied by a pronounced increase in the CD14+CD16+ monocyte subpopulation [23 24 25 26 27 28 ]. It is interesting that elevation in the number of the CD14+CD16+ monocytes has also been described during localized infections. In a localized skin infection caused by the parasite Leishmania braziliensis, a significant increase in the levels of CD16 expression on monocytes was observed, and the increase in the CD14+CD16+ monocytes correlated, not only with larger lesion size but also with elevated levels of plasma active TGF-β [29 ]. As TGF-β has been shown previously to induce CD16 expression on monocytes [30 ], the authors suggested that the production of TGF-β was responsible for the induction of CD16. An alternative which was not explored, is that IL-10, a cytokine that plays a key role in the pathogenesis of human visceral leishmaniasis [31 , 32 ], may be involved, as this cytokine has also been shown to induce elevated CD16 expression in cultured human monocytes [33 ]. During erysipelas, a skin infection caused by β-hemolytic Group A streptococci, an elevation in CD14+CD16+ monocytes was observed, and the appearance of these cells was associated with lower intracellular TNF-{alpha} production relative to cells from healthy patients [34 ]. In contrast, during pulmonary tuberculosis, an elevation in CD16 expression on monocytes was associated with elevated serum TNF-{alpha} levels [35 ]. It may therefore be an oversimplification to state unequivocally that the CD14+CD16+ monocytes act as proinflammatory monocytes during all in vivo infections.

In stark contrast to the expansion of the human CD14+CD16+ (CX3CR1hiCCR2–) monocyte subpopulation during infections, in the mouse the CX3CR1+CCR2+Gr1hi subpopulation is the population that is elevated during infections. For example, during systemic Listeria monocytogenes infection, the proportion of this population increased considerably [13 , 36 ], and almost all cell-associated bacteria were found in association with this subpopulation. It is interesting that infection with an avirulent L. monocytogenes mutant, which did not produce listeriolysin O and therefore, could neither escape the phagosomes nor replicate intracellularly, did not induce this increase [36 ].

The s.c. infection of susceptible BALB/c mice with Leishmania major promastigotes causes a visceralizing form of disease, which is different than the local, self-resolving, cutaneous disease caused in resistant C57BL/6 mice [37 ] and in most natural human infections. Experimental infections in BALB/c mice resulted in an increase in the Gr1hi monocyte subpopulation during the course of the infection (Fig. 2A and 2B) . In contrast, low-dose, intradermal L. major infection in C57BL/6 mice [39 ] did not result in detectable changes in the monocyte subpopulations (Fig. 2C and 2D) . Thus, in this model, systemic but not local infections caused dramatic alterations in monocyte subpopulations. Sunderkotter et al. [13 ] described an increase in the proportion of the Gr1hi subpopulation in C57BL/6 mice infected s.c. with a supraphysiological dose L. major promastigotes, suggesting that in the mouse, the extent of infection can determine and influence the development of monocyte subpopulations.


Figure 2
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  		Figure 2. Gr1 expression on mouse monocyte subpopulations during Leishmania infection. (A and B) BALB/c mice were infected s.c. in the foot-pad with 105 L. major promastigotes and monitored for 6 weeks after infection. (C and D) C57BL/6 mice were infected intradermally in the ear with 2 x 103 L. major promastigotes and monitored for 6 weeks after infection. Blood was obtained from these mice, stained with anti-CD115, anti-F4/80, and anti-Gr1, and gated as described previously [38 ]. (A and C) Representative histogram plots of mononuclear cells in the low, side-scatter range, gated upon CD115 and F4/80 positivity before infection and at 6 weeks after the infection. (B and D) Quantitation of the changes in the Gr1hi and Gr1low subpopulations during the infection.

BONE MARROW

Blood monocytes originate in the bone marrow from a common myeloid progenitor, which also gives rise to neutrophils. Recently, a specific macrophage and DC progenitor (MDP) population was identified in the mouse [40 ]. These cells were found to express the c-kit and the fractalkine receptor (CD117int/CX3CR1+) and were lacking any lineage-specific markers. This MDP was reported to give rise to monocytes, several subsets of macrophages, and steady-state DCs in vivo. It lacks, however, the potential to differentiate into polymorphonuclear, lymphoid, erythroid, or megakaryocytic cells. The MDP precursors have been shown recently to differentiate into CD11b+Gr1hi and CD11b+Gr1low bone marrow monocytes, which are the bone marrow-resident intermediates to the blood monocytes [41 ]. The previous thought that myeloid differentiation occurred only in one direction from the bone marrow to the periphery [15 ] has been replaced recently with the idea that Gr1hi monocytes can shuttle back from the blood to the bone marrow and even convert from CD11b+Gr1hi into CD11b+Gr1low bone marrow monocytes [41 ]. This finding can explain previous observations regarding the disappearance of adoptively transferred Gr1hi monocytes from the circulation in the absence of inflammation [8 , 13 ].

Murine blood monocytes have been implicated in the dissemination of several pathogens such as L. monocytogenes and Salmonella typhimurium [36 , 42 ]. The observation that the Gr1hi monocytes can shuttle back to the bone marrow raises the possibility that they can transport pathogens from the blood to the bone marrow or alternatively, that they can acquire antigens or infections in the bone marrow after leaving the blood. The former possibility would be consistent with the observation that Gr1hi bone marrow monocytes can spread L. monocytogenes infection to the brain [43 ]. The latter possibility would be consistent with the observation that CD11b+Gr1hi bone marrow monocytes were able to acquire antigens from neutrophils and B cells while in the bone marrow [38 ] and can possibly get infected in a similar way.

CCR2 expression on CD11b+Ly6Chi (Gr1hi) bone marrow monocytes appears to be necessary for the emigration of these cells to the blood, as these cells accumulate in the bone marrow of CCR2–/– mice [44 ]. Furthermore, during initial L. monocytogenes infection, the number of these CD11b+Ly6Chi (Gr1hi) cells in the bone marrow increased considerably in mice lacking CCR2 relative to wild-type mice, suggesting that the infection induces the generation of these cells, which are arrested in the bone marrow as a result of the lack of CCR2 expression. However, it is possible that CCR2 is not the only inducer of monocyte migration from the bone marrow during other infections. Over the course of a Leishmania amazonensis infection, we observed a gradual increase in the levels of Gr1hi monocytes in wild-type and CCR2–/– mice (Fig. 3 ). This suggests that during L. amazonensis infection, some CD11b+Ly6Chi monocytes can leave the bone marrow, even in the absence of CCR2 expression.


Figure 3
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  		Figure 3. Gr1hi blood monocytes show a transient increase during L. amazonensis infection of C57BL/6 and CCR2–/– mice, which were infected with 106 L. amazonensis parasites s.c. in the foot-pad and monitored for 9 weeks after the infection. Mononuclear cells in the low, side-scatter range were gated on CD11b and F4/80 positivity at different times after the infection, and the percentage of Gr1hi blood monocytes was quantitated. Note that the percentage of the Gr1hi blood monocytes increased temporarily after infection in C57BL/6 and also in CCR2 –/– mice.

PERITONEUM

Monocyte migration into the peritoneum has been studied widely under a variety of conditions, and therefore, this tissue has been considered as a reference from which to compare and contrast macrophage dynamics in all other tissues. After much early controversy, it is now well-established that during steady-state, peritoneal macrophages are self-maintained without significant recruitment of monocytic cells [45 , 46 ]. Furthermore, the grafting of CX3CR1+Gr1hi or the CX3CR1hiGr1– monocyte subpopulations did not result in substantial accumulation of donor cells in the peritoneum [8 ]. During inflammation, however, a different picture emerges.

It was demonstrated several years ago that the increased number of peritoneal macrophages during inflammation is a result of the recruitment of blood monocytes into the peritoneum [47 ]. Engraftment of bone marrow MDPs resulted in the accumulation of donor-derived CD11b+F4/80+ macrophages in the peritoneal exudates of thioglycollate-injected mice [40 ]. Moreover, thioglycollate-induced inflammation resulted in recruitment of donor-derived CD115+ bone marrow monocytes into the peritoneum, and most of them are Gr1hi [41 ].

Using a nonlethal, low-dose, i.p. infection with the parasite Toxoplasma gondii, Mordue and Sibley [48 ] identified the early recruitment of CD68+Gr1hi cells, which were characterized further as macrophages. These cells were able to produce IL-12, reactive nitrogen intermediates, and controlled parasite replication when stimulated in vitro, suggesting that they contributed to the control of parasite replication. CCR2–/– and MCP-1–/– mice had considerably lower numbers of the CD68+Gr1hi cells and failed to control parasite numbers, despite normal production of IFN-{gamma} [49 ]. It was therefore suggested that in this infection model, the ability of mice to resist acute toxoplasmosis is dependent on the successful recruitment of effector macrophages from Gr1hi monocytes via CCR2 and MCP-1.

The newly migrated Gr1hi monocytes may not always give rise to effector macrophages. It was shown recently that upon transfer of CX3CR1+Gr1hi blood monocytes into an inflamed peritoneum, a major part of the migrating donor monocytes had differentiated to express CD11c and MHC Class II molecules. These cells had the ability to stimulate naïve T cells, suggesting that they have differentiated into functional DCs [8 ]. Thus, it appears that under conditions of inflammation and infection, the Gr1hi monocytes, which can give rise to macrophages or DCs, are the major, early infiltrating monocyte subpopulation, migrating into the peritoneum. The signals that control this differentiation in vivo have yet to be defined completely.

SPLEEN

Bone narrow MDPs were found to give rise to steady-state CD11c+CD8{alpha}+ and CD11c+CD8{alpha}– splenic DCs and to splenic resident macrophages [40 ]. Thus, similar to what had been described previously in the peritoneum under steady-state conditions, splenic, conventional DCs (cDCs) are replenished primarily by local progenitors and only minimally through blood monocyte intermediates. However, the origins of the different splenic macrophage populations in steady-state appear to be somewhat more complex than in the peritoneum [50 , 51 ]. That is probably because the spleen has at least four different macrophage populations with different phenotypes and different functions [52 ]. Each population may be maintained by different mechanisms. Local proliferation of macrophage precursors appears to be particularly important for white pulp and metallophilic macrophages; however, monocyte influx may account for the other macrophage populations in the spleen [53 ]. A detailed analysis of each of the splenic macrophage populations after the transfer of specific monocyte subpopulations has yet to be done.

The maintenance of DCs in the spleen during steady-state appears to be largely dependent on local MDP proliferation. Gr1hi-grafted bone marrow monocytes failed to give rise to CD11chi splenic DCs [40 , 41 ]. Moreover, intrasplenic transfer of MDPs, as well as the characterization of an intrasplenic, steady-state DC precursor, which was found to be different from blood monocytes, provided further evidence that the role of blood monocytes in the generation of steady-state, splenic cDCs was minimal [41 , 54 ]. Recently, Naik and colleagues [54 ] confirmed that Ly6Chi (Gr1hi) blood monocytes could not generate splenic cDCs. They did, however, demonstrate that the Ly6Clow (Gr1low) subpopulation of monocytes was capable of generating low numbers of cDC under steady-state.

Similar to the peritoneum, the origin of splenic macrophages and DCs is changed upon inflammation [54 ]. In the absence of detailed studies pertaining to macrophages, it is logical to assume that the Gr1hi blood monocytes migrate into the spleen and contribute at least partially to the splenic macrophage populations. Under inflammatory conditions, transferred Ly6Chi (Gr1hi) blood monocytes were also able to differentiate into splenic DCs [54 ]. However, these DCs appeared to be distinct from the resident steady-state, splenic DCs (CD11chi CD11bintMac-3–), as they had low expression of CD8 and CD4 and were CD11cintCD11bhiMac-3+.

The i.v. inoculation of mice with the gram-negative bacteria L. monocytogenes, which infects the spleen and the liver primarily, gave rise to a novel population of DCs, which was recruited to the spleen in a CCR2-dependent manner. These DCs produced TNF and inducible NO synthase and were termed TipDCs [55 ]. These cells were characterized as CD11bintCD11cintGr1+F4/80–DEC205– and were found to be lacking in CCR2–/– mice, suggesting that they migrate into the spleen in response to MCP-1. It is interesting that the phenotype of these infection-induced TipDCs resembles closely the phenotype of the inflammatory, monocyte-derived, splenic DCs described above. The authors hypothesized originally that during L. monocytogenes infection, splenic macrophages, which become activated after clearing the bacteria, further secrete MCP-1, which recruits CCR2+Ly6Chi (Gr1hi) monocytes from the blood. These cells can then become TipDCs in the spleen. However, as mentioned earlier, a later study demonstrated that although CCR2 was required for the migration of Ly6Chi (Gr1hi) bone marrow monocytes to the bloodstream during this infection [44 ], it was not required for the migration of monocytes from the blood into the spleen. Nevertheless, the CX3CR1+CCR2+Gr1hi blood monocytes can still be the potential precursors for TipDCs, even if the MCP-1–CCR2 axis is not mandatory for the recruitment of these monocytes to the spleen in this infection model.

SKIN

The skin is a tissue which is in close contact with the environment, and as such, possesses a considerable number of mononuclear phagocytes. Langerhans cells (LC) are located in the suprabasal layer of the epidermis, and dermal DCs and dermal macrophages reside in the dermis. Under steady-state, the epidermal LC population is maintained by local radio-resistant precursors, and cells are replaced by circulating precursors only during major skin injuries [56 ]. The majority of dermal DCs is similarly maintained in quiescent skin, independently of circulating precursors; however, dermal macrophages were mixed almost completely 6 months after the generation of parabiotic mice [57 ], suggesting that their renewal is dependent on circulating precursors. A similar conclusion was reached in humans, where a small percentage (2–3%) of the dermal DCs is constitutively cycling in human quiescent skin.

In inflammatory conditions, after UV irradiation of mice skin, the direct precursors of LC and dermal macrophages were found to be the CCR2+Gr1hi blood monocytes [58 ]. It is interesting that using an in vitro model of reconstructed skin, it was reported that human CD14+CCR2+ monocytes differentiated into LC [59 ]. The existence of the CSF-1 receptor is apparently critical for the mouse LC repopulation [58 ]. During inflammation, locally proliferating, dermal DCs are replaced by circulating, dermal DC precursors in a CCR2-dependent manner; however, the exact nature of these precursors has not been determined yet [57 ].

During L. major infection of mice, a population of dermal macrophages, which was characterized by the expression of CD11b+F4/80+Gr1–CCR2+, could be identified in infected ears (Fig. 4A ) [60 ]. We have identified a 50% increase in the number of these cells 7 days after infection with transgenic L. major parasites secreting murine MCP-1 [60 ]. This increase in the CCR2+ macrophage migration correlated with a significant decrease in the number of viable parasites in the lesion relative to wild-type L. major-infected ears. Sorted CCR2+ macrophages from transgenic L. major-infected ears contained few if any parasites, while macrophages from wild-type, infected ears were heavily infected (Fig. 4B) . It has been shown previously that L. major infection induces increased MCP-1 production in resistant mouse strains but not in susceptible BALB/c mice [61 ]. We have detected a considerable increase in the number of the CD11b+Gr1–CCR2+ cells in L. major-infected ears from resistant C57BL/6 mice in comparison with susceptible BALB/c mice (D. Strauss-Ayali et al., unpublished observations), suggesting that a CCR2-dependent migration of macrophages can limit infection in this model. Studies to identify the precursors of these cells are under way.


Figure 4
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  		Figure 4. CCR2+ macrophages are recruited to the ears of mice infected with MCP-1 transgenic L. major parasites. BALB/c mice were infected in the ear with 5 x 104 wild-type (WT) L. major parasites or MCP-1 transgenic L. major parasites [60 ]. Ear cells were isolated 7 days postinfection and gated on CD11b and F4/80. (A) Gated CD11b+F4/80+ cells are negative for Gr1 and positive for CCR2 expression (green, isotype control; red/blue, cells from L. major wild-type and MCP-1 transgenic, infected ears, respectively). (B) CCR2+ macrophages were sorted from the ears of BALB/c mice 21 days postinfection with wild-type L. major parasites or MCP-1 transgenic L. major parasites and were cytospun and Giemsa-stained to examine macrophage infectivity.

LUNGS

The lung is a site of primary exposure to environmental stimuli as well as to potentially harmful pathogens. Lung-specific DC and macrophage populations are the major accessory cell populations with immunoregulatory potential in this environment [62 , 63 ]. Although earlier studies suggested a major role for local proliferation of precursors as the source of alveolar macrophages in steady-state [64 ], a recent study has found that the CX3CR1hiCCR2–Gr1low blood monocyte subpopulation also has the potential to differentiate into lung macrophages [65 ]. In this study, it was also shown that both mouse monocyte subpopulations, Gr1low and Gr1hi, could give rise to pulmonary DCs [65 ].

During bacterial pneumonia caused by various pathogens, the initial, inflammatory reaction involves resident alveolar macrophages, which is followed by the infiltration of mononuclear phagocytes into the lung. For instance, during pneumonia caused by Streptococcus pneumoniae, the number of lung macrophages was found to increase considerably [66 ]. Contrary to steady-state alveolar macrophages, which are CD11chiMHCIIintCD11b–, these cells expressed high levels of CD11b. It is most likely that these cells were derived from blood monocytes, which had up-regulated their CD11c expression during transit from the circulation into the infected tissue. These cells may increase their CD11b levels further in response to inflammation-induced GM-CSF. These lung macrophages lacked Gr1 expression. It is not clear whether they were derived from CX3CR1hiCCR2–Gr1low monocytes or from the CX3CR1+CCR2+Gr1hi monocytes and had down-regulated their Gr1 expression upon entry into the lung. Similarly, it is not clear whether the restoration of the steady-state lung macrophage population after infection occurred by the death of the CD11bhi macrophages or by the down-regulation of CD11b levels on macrophages as a result of a decrease in lung GM-CSF. During the initial stages of granulomatous response as a result of Mycobacterium tuberculosis infection of mice, several populations of macrophages and DC-like cells were characterized by the differential expression of the CD11b and CD11c surface molecules [67 ]. Similarly to the pneumococcal infection, the macrophages were suggested to originate from blood CD11b+CD11c– monocytes; however, an up-regulation of CD11b expression on resident lung macrophages could not be ruled out.

CONCLUDING REMARKS

Blood monocyte subpopulations appear similar in humans, mice, and rats, especially when based on the expression of some chemokine receptors (CCR2, CX3CR1), adhesion molecules (CD62L), and differences in size and granularity. However, during steady-state, the proportions of the classical versus nonclassical monocyte subpopulations in the blood of humans, mice, and rats are quite different. Moreover, during inflammation and systemic infections in humans, the number of the nonclassical CD14+CD16+CCR2– monocytes increases quite dramatically, and in the mouse, it is typically the classical population of Gr1+CCR2+ monocytes, which are elevated during infection. Monocyte dynamics in the rat are similar to those in the mouse. As much of the data about the function of human monocyte subpopulations are based on in vitro studies, it is necessary to get more in vivo data in the human to provide better understanding of the relationship between the monocyte subpopulations and the cells that they give rise to in tissue.

Regardless of the monocyte precursors, which give rise to macrophages, it is important to remember that the ultimate fate of the tissue macrophage is dictated primarily by the conditions that they encounter in tissue. Cytokines, chemokines, tissue metabolites, and inflammatory mediators can influence the differentiation of macrophages in tissue. Furthermore, these cells remain remarkably plastic and capable of reversing their phenotype in the face of changing conditions [68 ]. Thus, it may well be that the same monocyte subpopulation can differentiate into different types of macrophages depending on the tissue environment and the pathogens that are encountered in this environment. We would suggest that the ultimate tissue response is the result of an orchestrated interplay between the various monocyte subsets that are induced to leave the blood and the differentiative plasticity of macrophages in tissue.

ACKNOWLEDGEMENTS

This study was supported by National Institutes of Health grant AI055576.

Received March 28, 2007; accepted April 10, 2007.

REFERENCES

    1
  1. Ziegler-Heitbrock, H. W., Passlick, B., Flieger, D. (1988) The monoclonal antimonocyte antibody My4 stains B lymphocytes and two distinct monocyte subsets in human peripheral blood Hybridoma 7,521-527[Medline]
    2. 2
  2. Passlick, B., Flieger, D., Ziegler-Heitbrock, H. W. (1989) Identification and characterization of a novel monocyte subpopulation in human peripheral blood Blood 74,2527-2534[Abstract/Free Full Text]
    3. 3
  3. Frankenberger, M., Sternsdorf, T., Pechumer, H., Pforte, A., Ziegler-Heitbrock, H. W. (1996) Differential cytokine expression in human blood monocyte subpopulations: a polymerase chain reaction analysis Blood 87,373-377[Abstract/Free Full Text]
    4. 4
  4. Mizuno, K., Toma, T., Tsukiji, H., Okamoto, H., Yamazaki, H., Ohta, K., Ohta, K., Kasahara, Y., Koizumi, S., Yachie, A. (2005) Selective expansion of CD16highCCR2– subpopulation of circulating monocytes with preferential production of haem oxygenase (HO)-1 in response to accute inflammation Clin. Exp. Immunol. 142,461-470[Medline]
    5. 5
  5. Belge, K. U., Dayyani, F., Horelt, A., Siedlar, M., Frankenberger, M., Frankenberger, B., Espevik, T., Ziegler-Heitbrock, L. (2002) The proinflammatory CD14+CD16+DR++ monocytes are a major source of TNF J. Immunol. 168,3536-3542[Abstract/Free Full Text]
    6. 6
  6. Randolph, G. J., Sanchez-Schmitz, G., Liebman, R. M., Schakel, K. (2002) The CD16(+) (Fc{gamma}RIII(+)) subset of human monocytes preferentially becomes migratory dendritic cells in a model tissue setting J. Exp. Med. 196,517-527[Abstract/Free Full Text]
    7. 7
  7. Weber, C., Belge, K. U., von Hundelshausen, P., Draude, G., Steppich, B., Mack, M., Frankenberger, M., Weber, K. S., Ziegler-Heitbrock, H. W. (2000) Differential chemokine receptor expression and function in human monocyte subpopulations J. Leukoc. Biol. 67,699-704[Abstract]
    8. 8
  8. Geissmann, F., Jung, S., Littman, D. R. (2003) Blood monocytes consist of two principal subsets with distinct migratory properties Immunity 19,71-82[CrossRef][Medline]
    9. 9
  9. Ancuta, P., Rao, R., Moses, A., Mehle, A., Shaw, S. K., Luscinskas, F. W., Gabuzda, D. (2003) Fractalkine preferentially mediates arrest and migration of CD16+ monocytes J. Exp. Med. 197,1701-1707[Abstract/Free Full Text]
    10. 10
  10. Faure, S., Meyer, L., Costagliola, D., Vaneensberghe, C., Genin, E., Autran, B., Delfraissy, J. F., McDermott, D. H., Murphy, P. M., Debre, P., Theodorou, I., Combadiere, C. (2000) Rapid progression to AIDS in HIV+ individuals with a structural variant of the chemokine receptor CX3CR1 Science 287,2274-2277[Abstract/Free Full Text]
    11. 11
  11. Doranz, B. J., Rucker, J., Yi, Y., Smyth, R. J., Samson, M., Peiper, S. C., Parmentier, M., Collman, R. G., Doms, R. W. (1996) A dual-tropic primary HIV-1 isolate that uses fusin and the β-chemokine receptors CKR-5, CKR-3, and CKR-2b as fusion cofactors Cell 85,1149-1158[CrossRef][Medline]
    12. 12
  12. Glass, W. G., Lim, J. K., Cholera, R., Pletnev, A. G., Gao, J. L., Murphy, P. M. (2005) Chemokine receptor CCR5 promotes leukocyte trafficking to the brain and survival in West Nile virus infection J. Exp. Med. 202,1087-1098[Abstract/Free Full Text]
    13. 13
  13. Sunderkotter, C., Nikolic, T., Dillon, M. J., Van Rooijen, N., Stehling, M., Drevets, D. A., Leenen, P. J. (2004) Subpopulations of mouse blood monocytes differ in maturation stage and inflammatory response J. Immunol. 172,4410-4417[Abstract/Free Full Text]
    14. 14
  14. Yrlid, U., Jenkins, C. D., MacPherson, G. G. (2006) Relationships between distinct blood monocyte subsets and migrating intestinal lymph dendritic cells in vivo under steady-state conditions J. Immunol. 176,4155-4162[Abstract/Free Full Text]
    15. 15
  15. Gordon, S., Taylor, P. R. (2005) Monocyte and macrophage heterogeneity Nat. Rev. Immunol. 5,953-964[CrossRef][Medline]
    16. 16
  16. Grage-Griebenow, E., Zawatzky, R., Kahlert, H., Brade, L., Flad, H., Ernst, M. (2001) Identification of a novel dendritic cell-like subset of CD64(+)/CD16(+) blood monocytes Eur. J. Immunol. 31,48-56[CrossRef][Medline]
    17. 17
  17. Ziegler-Heitbrock, L. (2006) The CD14+ CD16+ blood monocytes: their role in infection and inflammation J. Leukoc. Biol. 81,584-592[CrossRef][Medline]
    18. 18
  18. Ahuja, V., Miller, S. E., Howell, D. N. (1995) Identification of two subpopulations of rat monocytes expressing disparate molecular forms and quantities of CD43 Cell. Immunol. 163,59-69[CrossRef][Medline]
    19. 19
  19. Grage-Griebenow, E., Flad, H. D., Ernst, M., Bzowska, M., Skrzeczynska, J., Pryjma, J. (2000) Human MO subsets as defined by expression of CD64 and CD16 differ in phagocytic activity and generation of oxygen intermediates Immunobiology 202,42-50[Medline]
    20. 20
  20. Sconocchia, G., Keyvanfar, K., El, O. F., Grube, M., Rezvani, K., Fujiwara, H., McCoy, J. P., Jr, Hensel, N., Barrett, A. J. (2005) Phenotype and function of a CD56+ peripheral blood monocyte Leukemia 19,69-76[Medline]
    21. 21
  21. Grip, O., Bredberg, A., Lindgren, S., Henriksson, G. (2007) Increased subpopulations of CD16(+) and CD56(+) blood monocytes in patients with active Crohn’s disease Inflamm. Bowel. Dis. 13,566-572[CrossRef][Medline]
    22. 22
  22. Qu, C., Edwards, E. W., Tacke, F., Angeli, V., Llodra, J., Sanchez-Schmitz, G., Garin, A., Haque, N. S., Peters, W., van Rooijen, N., Sanchez-Torres, C., Bromberg, J., Charo, I. F., Jung, S., Lira, S. A., Randolph, G. J. (2004) Role of CCR8 and other chemokine pathways in the migration of monocyte-derived dendritic cells to lymph nodes J. Exp. Med. 200,1231-1241[Abstract/Free Full Text]
    23. 23
  23. Fingerle, G., Pforte, A., Passlick, B., Blumenstein, M., Strobel, M., Ziegler-Heitbrock, H. W. (1993) The novel subset of CD14+/CD16+ blood monocytes is expanded in sepsis patients Blood 82,3170-3176[Abstract/Free Full Text]
    24. 24
  24. Fernandez, G. C., Ramos, M. V., Gomez, S. A., Dran, G. I., Exeni, R., Alduncin, M., Grimoldi, I., Vallejo, G., Elias-Costa, C., Isturiz, M. A., Palermo, M. S. (2005) Differential expression of function-related antigens on blood monocytes in children with hemolytic uremic syndrome J. Leukoc. Biol. 78,853-861[Abstract/Free Full Text]
    25. 25
  25. Allen, J. B., Wong, H. L., Guyre, P. M., Simon, G. L., Wahl, S. M. (1991) Association of circulating receptor Fc {gamma} RIII-positive monocytes in AIDS patients with elevated levels of transforming growth factor-β J. Clin. Invest. 87,1773-1779[Medline]
    26. 26
  26. Thieblemont, N., Weiss, L., Sadeghi, H. M., Estcourt, C., Haeffner-Cavaillon, N. (1995) CD14lowCD16high: a cytokine-producing monocyte subset which expands during human immunodeficiency virus infection Eur. J. Immunol. 25,3418-3424[Medline]
    27. 27
  27. Otani, I., Akari, H., Nam, K. H., Mori, K., Suzuki, E., Shibata, H., Doi, K., Terao, K., Yosikawa, Y. (1998) Phenotypic changes in peripheral blood monocytes of cynomolgus monkeys acutely infected with simian immunodeficiency virus AIDS Res. Hum. Retroviruses 14,1181-1186[Medline]
    28. 28
  28. Kim, W. K., Corey, S., Alvarez, X., Williams, K. (2003) Monocyte/macrophage traffic in HIV and SIV encephalitis J. Leukoc. Biol. 74,650-656[Abstract/Free Full Text]
    29. 29
  29. Soares, G., Barral, A., Costa, J. M., Barral-Netto, M., Van Weyenbergh, J. (2006) CD16+ monocytes in human cutaneous leishmaniasis: increased ex vivo levels and correlation with clinical data J. Leukoc. Biol. 79,36-39[Abstract/Free Full Text]
    30. 30
  30. Welch, G. R., Wong, H. L., Wahl, S. M. (1990) Selective induction of Fc {gamma} RIII on human monocytes by transforming growth factor-β J. Immunol. 144,3444-3448[Abstract]
    31. 31
  31. Karp, C. L., el-Safi, S. H., Wynn, T. A., Satti, M. M., Kordofani, A. M., Hashim, F. A., Hag-Ali, M., Neva, F. A., Nutman, T. B., Sacks, D. L. (1993) In vivo cytokine profiles in patients with kala-azar. Marked elevation of both interleukin-10 and interferon-{gamma} J. Clin. Invest. 91,1644-1648[Medline]
    32. 32
  32. Ghalib, H. W., Piuvezam, M. R., Skeiky, Y. A., Siddig, M., Hashim, F. A., el-Hassan, A. M., Russo, D. M., Reed, S. G. (1993) Interleukin 10 production correlates with pathology in human Leishmania donovani infections J. Clin. Invest. 92,324-329[Medline]
    33. 33
  33. Calzada-Wack, J. C., Frankenberger, M., Ziegler-Heitbrock, H. W. (1996) Interleukin-10 drives human monocytes to CD16 positive macrophages J. Inflamm. 46,78-85[Medline]
    34. 34
  34. Horelt, A., Belge, K. U., Steppich, B., Prinz, J., Ziegler-Heitbrock, L. (2002) The CD14+CD16+ monocytes in erysipelas are expanded and show reduced cytokine production Eur. J. Immunol. 32,1319-1327[CrossRef][Medline]
    35. 35
  35. Vanham, G., Edmonds, K., Qing, L., Hom, D., Toossi, Z., Jones, B., Daley, C. L., Huebner, B., Kestens, L., Gigase, P., Ellner, J. J. (1996) Generalized immune activation in pulmonary tuberculosis: co-activation with HIV infection Clin. Exp. Immunol. 103,30-34[CrossRef][Medline]
    36. 36
  36. Drevets, D. A., Dillon, M. J., Schawang, J. S., Van Rooijen, N., Ehrchen, J., Sunderkotter, C., Leenen, P. J. (2004) The Ly-6Chigh monocyte subpopulation transports Listeria monocytogenes into the brain during systemic infection of mice J. Immunol. 172,4418-4424[Abstract/Free Full Text]
    37. 37
  37. Sacks, D., Noben-Trauth, N. (2002) The immunology of susceptibility and resistance to Leishmania major in mice Nat. Rev. Immunol. 2,845-858[CrossRef][Medline]
    38. 38
  38. Tacke, F., Ginhoux, F., Jakubzick, C., van Rooijen, N., Merad, M., Randolph, G. J. (2006) Immature monocytes acquire antigens from other cells in the bone marrow and present them to T cells after maturing in the periphery J. Exp. Med. 203,583-597[Abstract/Free Full Text]
    39. 39
  39. Belkaid, Y., Kamhawi, S., Modi, G., Valenzuela, J., Noben-Trauth, N., Rowton, E., Ribeiro, J., Sacks, D. L. (1998) Development of a natural model of cutaneous leishmaniasis: powerful effects of vector saliva and saliva preexposure on the long-term outcome of Leishmania major infection in the mouse ear dermis J. Exp. Med. 188,1941-1953[Abstract/Free Full Text]
    40. 40
  40. Fogg, D. K., Sibon, C., Miled, C., Jung, S., Aucouturier, P., Littman, D. R., Cumano, A., Geissmann, F. (2006) A clonogenic bone marrow progenitor specific for macrophages and dendritic cells Science 311,83-87[Abstract/Free Full Text]
    41. 41
  41. Varol, C., Landsman, L., Fogg, D. K., Greenshtein, L., Gildor, B., Margalit, R., Kalchenko, V., Geissmann, F., Jung, S. (2007) Monocytes give rise to mucosal, but not splenic, conventional dendritic cells J. Exp. Med. 204,171-180[Abstract/Free Full Text]
    42. 42
  42. Vazquez-Torres, A., Jones-Carson, J., Baumler, A. J., Falkow, S., Valdivia, R., Brown, W., Le, M., Berggren, R., Parks, W. T., Fang, F. C. (1999) Extraintestinal dissemination of Salmonella by CD18-expressing phagocytes Nature 401,804-808[CrossRef][Medline]
    43. 43
  43. Join-Lambert, O. F., Ezine, S., Le, M. A., Jaubert, F., Okabe, M., Berche, P., Kayal, S. (2005) Listeria monocytogenes-infected bone marrow myeloid cells promote bacterial invasion of the central nervous system Cell. Microbiol. 7,167-180[Medline]
    44. 44
  44. Serbina, N. V., Pamer, E. G. (2006) Monocyte emigration from bone marrow during bacterial infection requires signals mediated by chemokine receptor CCR2 Nat. Immunol. 7,311-317[CrossRef][Medline]
    45. 45
  45. Sawyer, R. T., Strausbauch, P. H., Volkman, A. (1982) Resident macrophage proliferation in mice depleted of blood monocytes by strontium-89 Lab. Invest. 46,165-170[Medline]
    46. 46
  46. Melnicoff, M. J., Horan, P. K., Breslin, E. W., Morahan, P. S. (1988) Maintenance of peritoneal macrophages in the steady state J. Leukoc. Biol. 44,367-375[Abstract]
    47. 47
  47. Van Furth, R., Diesselhoff-den Dulk, M. C., Mattie, H. (1973) Quantitative study on the production and kinetics of mononuclear phagocytes during an acute inflammatory reaction J. Exp. Med. 138,1314-1330[Abstract]
    48. 48
  48. Mordue, D. G., Sibley, L. D. (2003) A novel population of Gr-1+-activated macrophages induced during acute toxoplasmosis J. Leukoc. Biol. 74,1015-1025[Abstract/Free Full Text]
    49. 49
  49. Robben, P. M., LaRegina, M., Kuziel, W. A., Sibley, L. D. (2005) Recruitment of Gr-1+ monocytes is essential for control of acute toxoplasmosis J. Exp. Med. 201,1761-1769[Abstract/Free Full Text]
    50. 50
  50. Van Furth, R., Diesselhoff-den Dulk, M. M. (1984) Dual origin of mouse spleen macrophages J. Exp. Med. 160,1273-1283[Abstract/Free Full Text]
    51. 51
  51. van Rooijen, N., Kors, N., Kraal, G. (1989) Macrophage subset repopulation in the spleen: differential kinetics after liposome-mediated elimination J. Leukoc. Biol. 45,97-104[Abstract]
    52. 52
  52. Taylor, P. R., Martinez-Pomares, L., Stacey, M., Lin, H. H., Brown, G. D., Gordon, S. (2005) Macrophage receptors and immune recognition Annu. Rev. Immunol. 23,901-944[CrossRef][Medline]
    53. 53
  53. Wijffels, J. F., de Rover, Z., Beelen, R. H., Kraal, G., van Rooijen, N. (1994) Macrophage subpopulations in the mouse spleen renewed by local proliferation Immunobiology 191,52-64[Medline]
    54. 54
  54. Naik, S. H., Metcalf, D., van Nienwenhuijze, A., Wicks, I., Wu, L., O’Keeffe, M., Shortman, K. (2006) Intrasplenic steady-state dendritic cell precursors that are distinct from monocytes Nat. Immunol. 7,663-671[CrossRef][Medline]
    55. 55
  55. Serbina, N. V., Salazar-Mather, T. P., Biron, C. A., Kuziel, W. A., Pamer, E. G. (2003) TNF/iNOS-producing dendritic cells mediate innate immune defense against bacterial infection Immunity 19,59-70[CrossRef][Medline]
    56. 56
  56. Merad, M., Manz, M. G., Karsunky, H., Wagers, A., Peters, W., Charo, I., Weissman, I. L., Cyster, J. G., Engleman, E. G. (2002) Langerhans cells renew in the skin throughout life under steady-state conditions Nat. Immunol. 3,1135-1141[CrossRef][Medline]
    57. 57
  57. Bogunovic, M., Ginhoux, F., Wagers, A., Loubeau, M., Isola, L. M., Lubrano, L., Najfeld, V., Phelps, R. G., Grosskreutz, C., Scigliano, E., Frenette, P. S., Merad, M. (2006) Identification of a radio-resistant and cycling dermal dendritic cell population in mice and men J. Exp. Med. 203,2627-2638[Abstract/Free Full Text]
    58. 58
  58. Ginhoux, F., Tacke, F., Angeli, V., Bogunovic, M., Loubeau, M., Dai, X. M., Stanley, E. R., Randolph, G. J., Merad, M. (2006) Langerhans cells arise from monocytes in vivo Nat. Immunol. 7,265-273[CrossRef][Medline]
    59. 59
  59. Schaerli, P., Willimann, K., Ebert, L. M., Walz, A., Moser, B. (2005) Cutaneous CXCL14 targets blood precursors to epidermal niches for Langerhans cell differentiation Immunity 23,331-342[CrossRef][Medline]
    60. 60
  60. Conrad, S. M., Strauss-Ayali, D., Field, A. E., Mack, M., Mosser, D. M. (2007) Leishmania-derived murine monocyte chemoattractant protein 1 enhances the recruitment of a restrictive population of CC chemokine receptor 2-positive macrophages Infect. Immun. 75,653-665[Abstract/Free Full Text]
    61. 61
  61. Zaph, C., Scott, P. (2003) Interleukin-12 regulates chemokine gene expression during the early immune response to Leishmania major Infect. Immun. 71,1587-1589[Abstract/Free Full Text]
    62. 62
  62. Holt, P. G. (2000) Antigen presentation in the lung Am. J. Respir. Crit. Care Med. 162,S151-S156[Abstract/Free Full Text]
    63. 63
  63. Fels, A. O., Cohn, Z. A. (1986) The alveolar macrophage J. Appl. Physiol. 60,353-369[Abstract/Free Full Text]
    64. 64
  64. Tarling, J. D., Lin, H. S., Hsu, S. (1987) Self-renewal of pulmonary alveolar macrophages: evidence from radiation chimera studies J. Leukoc. Biol. 42,443-446[Abstract]
    65. 65
  65. Landsman, L., Varol, C., Jung, S. (2007) Distinct differentiation potential of blood monocyte subsets in the lung J. Immunol. 178,2000-2007[Abstract/Free Full Text]
    66. 66
  66. Kirby, A. C., Raynes, J. G., Kaye, P. M. (2006) CD11b regulates recruitment of alveolar macrophages but not pulmonary dendritic cells after pneumococcal challenge J. Infect. Dis. 193,205-213[CrossRef][Medline]
    67. 67
  67. Gonzalez-Juarrero, M., Shim, T. S., Kipnis, A., Junqueira-Kipnis, A. P., Orme, I. M. (2003) Dynamics of macrophage cell populations during murine pulmonary tuberculosis J. Immunol. 171,3128-3135[Abstract/Free Full Text]
    68. 68
  68. Stout, R. D., Jiang, C., Matta, B., Tietzel, I., Watkins, S. K., Suttles, J. (2005) Macrophages sequentially change their functional phenotype in response to changes in microenvironmental influences J. Immunol. 175,342-349[Abstract/Free Full Text]



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