Published May 27, 2015, doi:10.4049/jimmunol.1500369 The Journal of Immunology

Specific Contributions of CSF-1 and GM-CSF to the Dynamics of the Mononuclear Phagocyte System Cynthia Louis,* Andrew D. Cook,* Derek Lacey,†,‡ Andrew J. Fleetwood,* Ross Vlahos,x,{,‖ Gary P. Anderson,x and John A. Hamilton* M-CSF (or CSF-1) and GM-CSF can regulate the development and function of the mononuclear phagocyte system (MPS). To address some of the outstanding and sometimes conflicting issues surrounding this biology, we undertook a comparative analysis of the effects of neutralizing mAbs to these CSFs on murine MPS populations in the steady-state and during acute inflammatory reactions. CSF-1 neutralization, but not of GM-CSF, in normal mice rapidly reduced the numbers of more mature Ly6C2 monocytes in blood and bone marrow, without any effect on proliferating precursors, and also the numbers of the resident peritoneal macrophages, observations consistent with CSF-1 signaling being essential only at a relatively late state in steadystate MPS development; in contrast, GM-CSF neutralization had no effect on the numbers of these particular populations. In Aginduced peritonitis (AIP), thioglycolate-induced peritonitis, and LPS-induced lung inflammation, CSF-1 neutralization lowered inflammatory macrophage number; in the AIP model, this reduced number was not due to suppressed proliferation. More detailed studies with the convenient AIP model indicated that CSF-1 neutralization led to a relatively uniform reduction in all inflammatory cell populations; GM-CSF neutralization, in contrast, was more selective, resulting in the preferential loss among the MPS populations of a cycling, monocyte-derived inflammatory dendritic cell population. Some mechanistic options for the specific CSF-dependent biologies enumerated are discussed. The Journal of Immunology, 2015, 195: 000–000.

M

onocytes and macrophages are highly plastic, mononuclear phagocyte system (MPS) populations and, depending on their cytokine environment, can generate heterogenous effector populations, contributing to various aspects of inflammation (1). M-CSF or CSF-1 and GM-CSF are key cytokines regulating the development and function of cells of the MPS, namely, monocytes, macrophages, and dendritic cells (DCs), both in the steady-state and during inflammation (2, 3). How they regulate the MPS cell number and/or their phenotypes have become important issues especially given that clinical trials targeting their action in chronic inflammation/autoimmunity are already under way. Studies on the biology of CSF-1 have become somewhat more complex with the discovery that there is another ligand, IL-34, that shares the CSF-1 receptor (CSF-1R; c-fms;

*Department of Medicine, The University of Melbourne, Royal Melbourne Hospital, Parkville, Victoria 3050, Australia; †Walter and Eliza Hall Institute of Medical Research, Parkville, Victoria 3052, Australia; ‡Department of Medical Biology, The University of Melbourne, Melbourne, Victoria 3052, Australia; xLung Health Research Centre, Department of Pharmacology and Therapeutics, The University of Melbourne, Parkville, Victoria 3010, Australia; {School of Health Sciences, Royal Melbourne Institute of Technology University, Bundoora, Victoria 3083, Australia; and ‖Health Innovations Research Institute, Royal Melbourne Institute of Technology University, Bundoora, Victoria 3083, Australia Received for publication February 19, 2015. Accepted for publication April 27, 2015. This work was supported by Project Grants and a Senior Principal Research Fellowship (to J.A.H.) from the National Health and Medical Research Council of Australia, and a University of Melbourne graduate research scholarship (to C.L.). Address correspondence and reprint requests to Prof. John A. Hamilton, The University of Melbourne, Department of Medicine, Royal Melbourne Hospital, Parkville, VIC 3050, Australia. E-mail: [email protected] The online version of this article contains supplemental material. Abbreviations used in this article: ADCC, Ab-dependent cell-mediated cytotoxicity; AIP, Ag-induced peritonitis; cDC, conventional DC; CSF-1R, CSF-1 receptor; DC, dendritic cell; i.n., intranasally; mBSA, methylated BSA; Mo-DC, monocytederived DC; MPS, mononuclear phagocyte system; PEC, peritoneal exudate cell; SSc, side-scatter. Copyright Ó 2015 by The American Association of Immunologists, Inc. 0022-1767/15/$25.00 www.jimmunol.org/cgi/doi/10.4049/jimmunol.1500369

CD115) (4–6). Even though CSF-1 and GM-CSF were first defined by their ability to generate in vitro macrophage and granulocyte colonies from progenitor cells (7–9), it is apparent that their cellular functions are much broader, with numerous changes brought about on mature MPS populations with relevance also to inflammatory reactions (2, 10, 11). One confounding factor in defining the specific roles of these CSFs in regulating cells in the MPS is the disagreement around the definition of macrophages and DCs (3, 12–14). A unified nomenclature has recently been proposed to attempt to circumvent this fundamental problem in which MPS cells can be classified primarily by their ontogeny and secondarily by their location, function, and phenotype in the steady-state and in inflammatory conditions (13). Even though a number of articles using sophisticated in vivo technology in mice have recently appeared addressing the origins of tissue macrophages during development, there is still debate around the fundamental question of the contribution, if any, of blood monocytes, including their subpopulations, to tissue macrophages in the adult at steady-state, and also to what extent local macrophage proliferation contributes to their homeostatic control or to that during inflammatory reactions (3, 15–24). Regarding the role of CSF-1 in the regulation of macrophage lineage development in vivo, there is disagreement as to how early a nonredundant function for CSF-1 is operating in the differentiation pathway (25, 26), and also how quickly, or even whether, blockade of its signaling can lead to depletion of certain monocyte/ macrophage populations in the steady-state (27–30); there is in addition disagreement about whether such a blockade can lower inflammatory macrophage numbers (27–30). It has been suggested that one reason for such discrepancies is the different mode of action of two different anti–CSF-1R mAbs (27–29, 31). Likewise, for the role of GM-CSF in inflammation, there is disagreement whether it can regulate the development of inflammatory DCs (monocytederived DCs [Mo-DCs]) (30, 32–34).

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CSF-1, GM-CSF, AND MONONUCLEAR PHAGOCYTE SYSTEM DYNAMICS

In our view, if we are to resolve some of these issues and to understand the respective biologies of CSF-1 and GM-CSF, it is important to make direct comparisons in the same experiments. Also, we consider that a specific mAb neutralization approach in a number of in vivo models is a worthwhile strategy to dissect these biologies. In this article, we monitored MPS population numbers and phenotypes using mainly such mAb neutralization in the steady-state and in a range of murine inflammation models. Two major conclusions below for the MPS are that CSF-1 signaling appears to be essential only at a relatively late stage in monocyte/ macrophage lineage development, most likely as a survival/growth factor and/or tissue retention factor, whereas GM-CSF depletion results in the preferential loss of a monocyte-derived, inflammatory DC population in certain contexts.

CD11c (clone HL3), CD115 (clone AFS98), CD117 (clone 2B8), Ly6C (clone AL-21), Ly6G (clone 1A8), F4/80 (clone BM8), CD103 (clone 2E7), CD64 (clone X54-5/7.1), BrdU (clone B44), and the corresponding isotype controls, from either BD Biosciences or Biolegend. Cell viability before sorting was assessed using 7-amino-actinomycin D (BD Pharmingen). Cells were analyzed using a CyAn ADP Analyzer (Beckman Coulter) or LSRFortessa (BD Biosciences), or sorted using a FACSAria III instrument (BD Biosciences).

Adoptive cell transfer Bone marrow was flushed from the tibias and femurs of donor MacGreen mice. After RBC lysis using ACK lysis buffer, CD115+ cells were MACSenriched using CD115-Biotin Ab and anti-Biotin microbeads (Miltenyi Biotec) according to manufacturer’s instructions. Monocyte purity after enrichment was consistently .90%. A total of 1.0 3 106 enriched monocytes were transferred i.v. into mBSA-challenged AIP mice on day 1.

BrdU pulsing

Materials and Methods Reagents The following mAbs were gifts: neutralizing anti-mouse CSF-1R (AFS98 [IgG2a]: S-I Nishikawa; RIKEN Center for Developmental Biology, Kobe, Japan), neutralizing anti-mouse GM-CSF (22E9.11; IgG2a): J. Abrams; DNAX, Palo Alto, CA), and neutralizing anti-mouse CSF-1 and IgG2a isotype control (F. Dodeller; MorphoSys, Munich, Germany) (35).

Mice C57BL/6 mice were obtained from WEHI, Kew (VIC, Australia). GMCSF2/2 (29) and Csf1r-EGFP (MacGreen) mice (36), backcrossed onto the C57BL/6 background, are bred in our on-site animal facility. BALB/c mice were obtained from the Animal Resource Centre (Perth, Australia). Mice were fed standard rodent chow and water ad libitum, and were housed in sawdust-lined cages in groups of five. Mice 8–12 wk old were used in all experiments, which were approved by the Animal Experimentation Ethics Committee of The University of Melbourne and conducted in compliance with the guidelines of the National Health and Medical Research Council of Australia on animal experimentation.

Experimental models Peritonitis models. Ag-induced peritonitis (AIP) was as described previously (37) except that more challenge Ag was administered. C57BL/6 mice were immunized intradermally at the base of the tail with 100 mg methylated BSA (mBSA; Sigma-Aldrich, St. Louis, MO), emulsified in an equal volume of CFA containing 5 mg/ml heat-killed Mycobacterium tuberculosis (H37Ra; Difco, Detroit, MI); 14 d later, the primary immunization protocol was repeated as a boost. Seven days later, mice were injected i.p. with 200 mg mBSA to induce peritonitis. Peritoneal exudate cells (PECs) were also elicited by i.p. injection of 1 ml Brewer’s thioglycolate medium (Difco) as previously described (37). LPS-induced inflammation. For the lung inflammation model, BALB/c mice were injected intranasally (i.n.) with 10 mg LPS (Escherichia coli serotype 026:B6; Sigma-Aldrich) (29). For the spleen inflammation model, BALB/c mice were injected i.v. with 10 mg LPS (E. coli serotype 026:B6; SigmaAldrich) (30).

In steady-state BrdU kinetics experiments, mice were given three separate i.p. injections of 2 mg BrdU (Sigma-Aldrich) 2 h apart, and BrdU labeling was monitored from days 1 to 5. For the short-term BrdU pulse, mice were injected with 1 mg BrdU i.p. 2–3 h before analysis. To assess BrdU incorporation, we stained, fixed, and permeabilized cell suspensions using Cytofix/Cytoperm and Perm/Wash buffer (BD Pharmingen) according to the manufacturer’s instructions. Cells were incubated at 37˚C for 60 min in 30 mg DNase (Sigma-Aldrich), followed by staining with anti–BrdU-FITC for 30 min, washed, and analyzed by flow cytometry.

Isolation and in vitro culture of MPS populations For sorting blood monocyte subsets, blood was collected by cardiac puncture into heparin solution, RBCs lysed using ACK lysis buffer, and cells stained for Ly6G, CD115, CD11b, and Ly6C. Monocytes were sorted as 7amino-actinomycin D2 Ly6G2 CD115+ CD11b+ into Ly6C+ and Ly6C2 subsets. Monocyte purity after sorting was consistently .98%. For sorting AIP MPS populations, cells were stained for Ly6G, CD115, CD11c, and MHCII. Sorted cells were cultured in 96-well ultra-low attachment plates (Costar) for 24 or 96 h for sorted AIP MPS populations as indicated, or for 48 h for blood monocyte subsets, in RPMI 1640 supplemented with 10% heat-inactivated FCS, 2 mM L-glutamine, and 100 U penicillin and streptomycin. GM-CSF (10 ng/ml) or CSF-1 (50 ng/ml) was added to some wells. Cultured blood monocytes were analyzed for viability and the expression of Ly6C, F4/80, CD11c, and MHCII. Cultured AIP MPS subsets were analyzed for cell viability or gene expression.

Gene expression analysis AIP MPS subsets were FACS sorted as R1, R2 + R3, and R4 populations (see later for nomenclature) from wild-type C57BL/6 mice as indicated, and total RNA was isolated using RNeasy kits (QIAGEN). RNA was reverse transcribed into cDNA (SuperScript II; Invitrogen), and quantitative real-time PCR was performed on a 7900 HT instrument (Applied Biosystems) (38). Predeveloped FAM-labeled primers (Applied Biosystems) were used for IL-6, IL-1b, IL-12p35, IL-23p19, CSF-1R, FLT3, ZBTB46, CCR7, and CSF2Ra. Custom-designed prime sequences were used for TGM2, MERTK, and UBC using SYBR green. Gene expression was analyzed using UBC as an endogenous control.

mAb treatment

Statistical analysis

For each mAb administration, mice were given 150 mg anti–GM-CSF (22E9), 250 mg anti–CSF-1R (AFS98), 150 mg anti–CSF-1, or their respective isotype control mAb. In steady-state experiments, mice were i.p. injected on days 0 and 2, and tissues were analyzed on day 4. In steadystate BrdU kinetics experiments, mice were given mAbs i.p. starting on day 21, day 0 being the day of BrdU pulsing, and booster mAb injections were given every other day until day 4. For peritonitis experiments, mAbs were given i.p. on days 1 and 2, day 0 being the day of peritonitis induction, and cells were analyzed on day 3 or 4 as indicated. For the LPSinduced lung inflammation model, mAbs were given i.n. 3 h before LPS challenge, and bronchoalveolar lavage was examined 3 d after. For LPSinduced splenic inflammation, mAbs were given i.p. on days 21 and 3 h before LPS i.v. challenge, and the spleen was analyzed 24 h after.

Data are expressed as mean 6 SEM. Statistical differences were assessed using unpaired Student t test, paired Student t test, or one-way ANOVA Prism 4 software (GraphPad Software) as indicated. A p value #0.05 was considered statistically significant.

Cell isolation and FACS analysis Cells suspensions were prepared as previously described (29) and analyzed by flow cytometry. For FACS analysis or sorting, cells were stained with Fc block anti-CD16/32 (clone 2.4G2) and fluorochrome-conjugated mAbs specific for mouse MHCII (clone M5/114.15.2), CD11b (clone M1/70),

Results CSF-1 is required for homeostatic maintenance of murine Ly6C2 monocytes Previously, CSF-1R neutralization using an IgG2a mAb isotype (clone AFS98) led to a rapid preferential depletion of murine Ly6C2 blood monocytes (28–30), whereas an IgG1 mAb isotype (clone M279) gave a slower rate of depletion (27). It was argued that this difference was due to an Ab-dependent cell-mediated cytotoxicity (ADCC) mechanism associated with the IgG2a isotype of the AFS98 mAb (27). To test whether this rapid depletion after CSF-1R neutralization with AFS98 might be caused by such

The Journal of Immunology an ADCC mechanism and also to exclude a role for the alternative CSF-1R ligand, namely, IL-34, we monitored the monocyte populations in the bone marrow, spleen, and blood (for gating, see Supplemental Fig. 1A) of mice treated with anti–CSF-1 mAb. Similar to the effect of AFS98 (28, 29), anti–CSF-1 rapidly depleted the Ly6C2 monocytes from each of these sites, whereas the Ly6C+ monocytes were only reduced, albeit slightly, in the bone marrow (Fig. 1A). Thus, the rapid Ly6C2 blood monocyte depletion seen earlier in anti–CSF-1R (AFS98)–treated animals (28, 29) is likely to be due to neutralization of CSF-1–specific signaling. This preferential loss of Ly6C2 monocytes from the sites in question would not appear to be due to inhibition of their proliferation because they were not cycling to any appreciable extent, as monitored by a 2-h BrdU pulse (Supplemental Fig. 1B). CSF-1 neutralization does not prevent early monopoiesis or the loss of surface Ly6C from Ly6C+ monocytes The selective depletion of the blood Ly6C2 monocytes after anti– CSF-1/CSF-1R neutralization would not appear to be due to higher surface CSF-1R expression because such expression has been reported to be similar on blood Ly6C2 and Ly6C+ monocytes (29, 39). Another possible mechanism is that CSF-1 is required for the Ly6C+ monocyte conversion (maturation) into Ly6C2 monocytes (27, 28, 40). Such a requirement might be expected to lead to an accumulation of Ly6C+ monocytes as a result of AFS98 (CSF1R) blockade, which, however, was not observed (28, 29) possibly because of the relatively short t1/2 (18 h) of this population in the circulation (41) (see Discussion). To examine further the role of CSF-1 in governing the turnover rate and the precursor-product relationship of the monocyte subpopulations, we again administered anti–CSF-1 mAb, but in

FIGURE 1. Requirement for CSF-1 for the homeostatic maintenance of monocytes. (A) Number of steady-state monocyte subpopulations in C57BL/6 mice at day 4 after isotype or anti–CSF-1 mAb administration at days 0 and 2. (B and C) BrdU labeling kinetics of monocyte subpopulations in bone marrow (left) and blood (right) of C57BL/6 mice treated three separate times with BrdU on day 0. Mice also received isotype or anti– CSF-1 mAb starting on day 21 and every other day until day 4. (B) Ly6C+ monocytes; (C) Ly6C2 monocytes. Data are mean 6 SEM of three experiments; n = 5 mice/group. Unpaired Student t test; isotype versus anti–CSF-1. *p , 0.05, ***p , 0.005.

3 combination with a BrdU tracing protocol. Mice were given three i.p. injections of BrdU (2 h apart), and the BrdU labeling was monitored in the bone marrow and blood monocyte compartments over a 5-d period (41). With this method, all Ly6C+ monocytes in the bone marrow and blood rapidly acquired BrdU within 1 d (41), and this BrdU uptake and kinetics were not affected by CSF-1 neutralization (Fig. 1B, Supplemental Fig. 2A, 2B). It has been shown previously that BrdU-labeled blood Ly6C+ monocytes derive from the cycling bone marrow Ly6C+ monocytes (41); the relative delay in reaching their maximum incorporation (Fig. 1B) is consistent with this concept. The data in Fig. 1B also indicate that the proliferating Ly6C+ monocytes in the bone marrow were still capable of dividing even upon CSF-1 neutralization. Supporting this finding, the recently characterized less mature and proliferating (CD115+ CD1172 [c-Kit] Ly6C+ CD11b2) monocyte population in the bone marrow (42) (Supplemental Fig. 1A) also exhibited similar BrdU labeling kinetics in anti–CSF-1–treated mice as compared with their isotype-treated cohort (Supplemental Fig. 2C). In line with their relatively low proliferation (42) (Supplemental Fig. 1B), bone marrow Ly6C2 monocytes, unlike their Ly6C+ counterparts (Fig. 1B, Supplemental Fig. 1B), were not immediately labeled with BrdU but rather underwent progressive BrdU incorporation presumably as a result of subsequent loss of Ly6C expression by BrdU-labeled Ly6C+ monocytes (Fig. 1C, Supplemental Fig. 2A) (41). Reflecting this progression, despite the rapid depletion of Ly6C2 monocytes after CSF-1 neutralization (Fig. 1A), the proportion of BrdU+ cells among the remaining Ly6C2 bone marrow and blood monocytes actually increased as compared with isotype-treated mice (Fig. 1C, Supplemental Fig. 2A, 2B). This increase was presumably due to CSF-1 neutralization preferentially depleting the BrdU2 Ly6C2 population (Supplemental Fig. 2A, 2B), that is, the ones generated before the BrdU pulsing. The fact that bone marrow and blood BrdU+ Ly6C2 monocytes still developed concurrently from BrdU + Ly6C+ monocytes even in the presence of anti–CSF-1 mAb (Supplemental Fig. 2A, 2B) suggests, perhaps surprisingly, that CSF-1 is not involved in the loss of surface Ly6C expression. To extend the earlier observations as to the role of CSF-1 in governing the precursor-product relationship and the turnover rate of the blood monocyte subpopulations, we turned to an in vitro system, in which blood monocyte subsets were sorted and cultured in the absence or presence of CSF-1. As shown in Supplemental Fig. 2D, over 2 d Ly6C+ untreated monocytes underwent spontaneous loss of Ly6C expression, and this pattern was comparable with that noted in CSF-1–treated cells, thus corroborating the in vivo observations in Supplemental Fig. 2B; even after 5 d in the presence of CSF-1, Ly6C was still detected (data not shown). Even though we could not show that CSF-1 removes surface Ly6C from the Ly6C+ monocytes in vitro, and even though CSF-1 depletion had no effect on their numbers in vivo, this population, but not the Ly6C2 monocytes, could still respond to CSF-1 in vitro by upregulating surface F4/80 (Supplemental Fig. 2E); this response could reflect differentiation to a macrophage phenotype, and there was some specificity because neither surface CD11c nor MHCII were upregulated by CSF-1 in the Ly6C+ monocytes (nor in the Ly6C2 monocytes). By way of comparison, GM-CSF, in contrast, preferentially upregulated CD11c and MHCII in the Ly6C+ monocytes, but not F4/80 (Supplemental Fig. 2E); we also found that it lowered monocyte Ly6C expression in vitro (Supplemental Fig. 2D) even though its deletion in vivo did not alter the steady-state blood monocyte subpopulation numbers or ratio (29).

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CSF-1, GM-CSF, AND MONONUCLEAR PHAGOCYTE SYSTEM DYNAMICS

CSF-1 can maintain both Ly6C+ and Ly6C2 monocytes in vitro An additional possible mechanism to explain the specific loss of Ly6C2 monocytes from the blood upon CSF-1 neutralization is their preferential requirement for CSF-1 to maintain their survival. However, consistent with their similar CSF-1R expression (29, 39), at least in vitro, CSF-1 can maintain the numbers of both Ly6C+ and Ly6C2 blood monocyte populations over a 48-h period (Supplemental Fig. 2F) (see Discussion). Again, the data for GMCSF are included by way of comparison and show that GM-CSF has a mitogenic action on the Ly6C+ blood monocytes (see later). CSF-1 is required for the homeostatic maintenance of resident peritoneal macrophages Because we wanted to extend our analysis of the CSF-mediated control of the MPS to that occurring in tissues, we used the peritoneal cavity as a convenient and sterile nonlymphoid tissue. We defined the MPS in the naive peritoneal cavity using a CD115based gating strategy as proposed before (43) and, notwithstanding the challenges faced in defining categorically MPS populations (3, 12–14), the following widely adopted surface marker nomenclature was mainly used later for this purpose. As shown in Fig. 2A, we defined macrophages as CD115+ CD11b+; these comprised the major MHCII2 (F4/80hi) and the minor MHCII+ (F4/80int) macrophages (data not shown) (44–46). In the CD1152 gate, a very small but distinct CD11c+ MHCII+ population was evident; we refer to these as conventional DCs (cDCs) and, as for cDCs described in other tissues (47, 48), these consisted of a major CD11b+ (CD1032) and a minor CD11b2 (CD103+) subset (Fig. 2A). We also monitored the BrdU labeling pattern of the macrophages and the putative cDC population. A single 2-h BrdU pulse labeled a small proportion of the macrophages (Fig. 2B). In addition, using the BrdU tracing protocol as described earlier to give time for cycling precursors to be labeled (three i.p. injections, 2 h apart), by day 4 only ∼4% of the CD115+ macrophages were BrdU+ (Fig. 2C), indicative of a slow turnover rate typically noted before for macrophages in sterile tissues (21, 46); in contrast, ∼35% of CD1152CD11c+MHCII+ cells were BrdU+ (Fig. 2C), indicative of a fast turnover rate characteristic of cDCs and presumably via recruitment from rapidly dividing bone marrow precursors (49, 50). We have shown previously that anti–CSF-1R mAb (AFS98) rapidly depleted resident peritoneal macrophages as judged by cell morphology (29). As shown in Fig. 2D, mice treated with anti– CSF-1R or anti–CSF-1 mAb had similarly reduced numbers of resident peritoneal macrophages compared with isotype control mAb-treated mice, again indicating control by CSF-1 signaling. Consistent with their lack of CSF-1R (CD115) expression, the cDCs were not affected by CSF-1R/CSF-1 blockade (Fig. 2D). The rapid macrophage depletion after CSF-1 neutralization would not appear to be due to suppression of peritoneal macrophage proliferation given their low steady-state cycling status (Fig. 2B) (see Discussion). As described before (29), anti–GM-CSF–treated or GM-CSF2/2 mice had comparable numbers of resident peritoneal macrophages (Fig. 2D), indicating a GM-CSF–independent homeostasis; unlike DCs in barrier tissues, such as the gut (30), steady-state peritoneal cDC numbers were also GM-CSF independent (Fig. 2D). In AIP, CSF-1 is required for the accumulation of all populations, whereas for the MPS, GM-CSF preferentially controls the Mo-DC population MPS populations in AIP. Because we next aimed to explore the CSF-mediated control over tissue MPS development and function during an inflammatory response, we decided to use an AIP model

FIGURE 2. CSF-1, but not GM-CSF, controls steady-state peritoneal macrophage numbers. (A) Representative FACS plots showing the gating strategy to define resident peritoneal macrophage and DC subsets. After pregating on CD115+ CD11b+ cells, peritoneal macrophages were subdivided into two populations based on MHCII expression (44): in the CD1152 gate, a cDC population was identified as CD11c+ MHCII+, which was further subdivided into CD11b2 CD103+ and CD11b+ CD1032 (47). (B) BrdU labeling of steady-state peritoneal macrophages and cDCs after a 2-h pulse. (C) BrdU labeling of steady-state peritoneal macrophages and cDCs at day 4 after treated three separate times with BrdU on day 0. (B and C) Numbers indicate the frequency of cells within the indicated gates. (D) Number of steady-state peritoneal macrophages and cDCs in C57BL/6 mice at day 4 after isotype, anti–CSF-1R, anti–CSF-1, or anti–GM-CSF mAb administration at days 0 and 2 (top), or in wild-type and GM-CSF2/2 mice. Data are mean 6 SEM of two experiments; n = 5 mice/group. Unpaired Student t test; isotype versus anti–CSF-1R, anti–CSF-1, or anti– GM-CSF. ***p , 0.005.

that, unlike the widely used thioglycolate-induced peritonitis (37) (see later), we had shown before to be GM-CSF dependent as judged at least by exudate numbers (29, 51, 52). In this model, C57BL/6 mice are immunized intradermally with Ag (mBSA) in CFA, followed later by an i.p. Ag challenge to induce cellular exudates (29, 37, 51). For the present studies, we used a higher amount of challenge Ag than previously (200 versus 100 mg) to generate a stronger and more prolonged inflammatory reaction. Before exploring the roles of CSF-1 and GM-CSF in this model, we first defined the MPS in the inflamed peritoneal cavity again

The Journal of Immunology

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using our CD115-based gating strategy. At 4 d after AIP induction by Ag challenge, exudate CD115+ CD11b+ cells could be divided into three subpopulations (R1–R3; Fig. 3A). The CD115+ CD11c+ MHCII+ subset (R1) was designated as Mo-DCs (Fig. 3A) (53– 55). These R1 cells displayed a DC-like morphology (Fig. 3B). In contrast with CD11c+ MHCII+ Mo-DCs reported elsewhere to be Ly6C+ (30, 34, 52), but consistent with Mo-DCs described in other models (53, 56), AIP Mo-DCs were mostly Ly6C2 (Fig. 3B). A second subset of CD115+ CD11b+ cells (R2) was designated as macrophages, constituting the majority of the CD115+ cells and expressing MHCII, but not CD11c (Figs. 3A and 4A) (56). These R2 cells had a macrophage appearance and displayed heterogeneous Ly6C expression (Fig. 3B), consistent with their being of blood monocyte origin and their maturation over time into Ly6C2 cells as the inflammatory reaction persists (16, 29, 43, 55, 56) (see later). A third subset of CD115+ CD11b+ cells (R3) was designated as monocytes because they expressed neither CD11c nor MHCII (Fig. 3A) but were mainly Ly6C+ (Fig. 3B); their morphology was consistent with this (Fig. 3B). In the CD1152 gate, we observed a CD11c+ MHCII+ (R4) cDC population (Fig. 3A), with morphological characteristics consistent with this nomenFIGURE 4. CSF-1 and GM-CSF dependence of MPS population numbers in AIP. (A) PECs were harvested at day 4 after AIP induction from C57BL/6 mice treated with isotype, anti–CSF-1R, anti–CSF-1, or anti– GM-CSF mAb. (B) FACS plots, gated on CD115+, depict the frequencies of the R1–R3 subpopulations. Data are mean 6 SEM from at least three experiments; n = 5 mice/group. One-way ANOVA; isotype versus anti– CSF-1R, anti–CSF-1, or anti–GM-CSF. *p , 0.05, **p , 0.01.

FIGURE 3. MPS in AIP. (A) Representative FACS plots showing the gating strategy to define peritoneal exudate macrophage and DC subsets in the day 4 AIP cavity (see Materials and Methods). After pregating on CD115+ CD11b+ cells, peritoneal cells were subdivided into R1 Mo-DCs, R2 macrophages, and R3 monocytes, based on their CD11c and MHCII expression. The CD1152 gate included cDCs (R4, CD11c+ MHCII+), neutrophils (R5, Ly6Ghi CD11b+), and eosinophils (R6, CD11bint SSchi). (B) Sorted MPS R1–R4 populations; (top) Giemsa staining; (bottom) Ly6C and CD11b expression profile. Numbers indicate the frequency of cells within the indicated gates. (C) Mean fluorescence intensity of surface F4/ 80 and CD64 expression by R1–R6 populations described in (A). Data are representative of more than three experiments; n = 5 mice/group.

clature (Fig. 3B), which were Ly6C2 and again either CD11b+ or CD11b2 (Fig. 3B), as described in the steady-state peritoneum (Fig. 2A) and in other nonlymphoid tissues (30, 47, 48). The CD1152 gate also contained Ly6Ghi CD11b+ neutrophils (R5), Ly6G2 CD11bint SSchi eosinophils (R6; Fig. 3A), and lymphocytes (not depicted). In support of the nomenclature, the neutrophils and cDCs, but not the lymphocytes, expressed surface GM-CSF receptor a-chain (data not shown). F4/80 and CD64 expression are often used to distinguish macrophages and Mo-DCs from cDCs (53, 57, 58). Overall, all CD115+ subpopulations (R1– R3) had higher expression of both F4/80 and CD64 than the CD1152 exudate cells (R4–R6; Fig. 3C), although the eosinophils expressed reasonable levels of the former marker. F4/80 and CD64 were lower in the R3 monocytes than in the R2 macrophages consistent with a less mature state. By way of additional definition of the MPS populations in the AIP model, we examined from sorted R1, R2 + R3, and R4 populations the gene expression profiles of several hallmark lineage-defining transcripts, namely, CSF-1R, MerTK, TGM2 for macrophages, and FLT3, ZBTB46, and CCR7 for cDCs, as well as GM-CSFRa itself (32, 47, 50, 53, 59–61). As depicted in Supplemental Fig. 3A, higher CSF-1R, MerTK, and TGM2 expression were evident in the CD115+ R1 and R2 + R3 populations, whereas FLT3, ZBTB46, and CCR7 expression were found only in the R4 cDC population, observations consistent with the surface marker phenotyping; GM-CSFRa mRNA was distributed uniformly. Monocyte origin of the MPS populations in AIP. To gain support for the Ly6C+ monocyte origin of the R1–R3 populations in the AIP model, we injected i.v. Ly6C+ GFP+ bone marrow monocytes from the Csf1r-EGFP (Macgreen) mouse (36) into syngeneic C57BL/6 recipients 24 h after AIP induction, that is, around the time Ly6C+ mononuclear cells appear in the cavity in this model (29). Seventy-two hours after this i.v. transfer, GFP+ cells could be

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CSF-1, GM-CSF, AND MONONUCLEAR PHAGOCYTE SYSTEM DYNAMICS

readily identified within the CD115+ CD11b+ gate, but not among the CD1152 exudate cells (Supplemental Fig. 3B). In addition, the GFP+ cells had mixed expression of CD11c and MHCII as for the R1–R3 subpopulations, in line with a monocytic origin of these populations, but not of the R4 cDCs. CSFs and MPS numbers in AIP. CSF-1R or GM-CSF neutralization diminished the accumulation of AIP exudate macrophages during AIP development, although a detailed phenotypic analysis was not carried out previously (29, 51). As for the steady-state peritoneum (Fig. 2D), we compared the effect of anti–CSF-1R and anti–CSF-1 administration on the AIP MPS populations to determine whether it is CSF-1 signaling that is in fact important and whether there is selectivity in cell population control. There was a reduction in total exudate cell number, which was uniform in all CD115+ R1–R3 subpopulations by both mAbs (Fig. 4A, 4B). Interestingly, all CD1152 cell populations were also reduced by anti–CSF-1 mAb and, apart from eosinophils where there was a trend, also by anti– CSF-1R (Supplemental Fig. 3C) (see Discussion). In contrast with CSF-1R/CSF-1 neutralization, at the dose of 2 3 150 mg anti–GM-CSF mAb, among the CD115+ cells, only the Mo-DC population was reduced both numerically (Fig. 4A) and proportionally (Fig. 4B); this anti–GM-CSF mAb protocol resulted in only a trend in the reduction of inflammatory macrophage and monocyte numbers (Fig. 4A). CD1152 populations that expressed the GM-CSF receptor, namely, PMN, cDCs, and eosinophils, were also lowered, but not the CD1152 lymphocyte population (Supplemental Fig. 3C). At the higher dose of anti–GM-CSF mAb (2 3 375 mg), in addition to MoDCs, inflammatory macrophages and monocytes were also significantly reduced, but again not the lymphocytes (Supplemental Fig. 3D). Thus, in AIP, CSF-1 neutralization around the time of Ag challenge indicates that CSF-1 at this time point controls the accumulation of all MPS and other exudate populations; GM-CSF neutralization, in contrast, preferentially controls among the MPS populations the number of Mo-DCs (see Discussion). CSFs and MPS cycling and gene expression in AIP There has been more interest recently in the contribution of local proliferation of inflammatory macrophage/DC populations to their increased numbers (17, 19, 20, 62). To explore whether reduced cell cycling in the inflamed cavity itself might be contributing to the lower numbers of AIP MPS populations observed earlier upon CSF neutralization, we pulse-labeled mAb-treated AIP mice with BrdU 2 h before termination at day 3. Macrophages and monocytes were labeled only to a small extent (#2%), whereas, interestingly, ∼11% of the Mo-DC population (Fig. 5A) and cDCs (data not shown) were in S phase after this short pulse. Administration of the anti–GM-CSF mAb, but not the anti–CSF-1 mAb, reduced the percentage of BrdU+ Mo-DCs (Fig. 5B), suggesting perhaps that the accumulation of AIP Mo-DCs could be partially dependent on local proliferation in a GM-CSF–dependent manner. The cycling status of the cDCs was not diminished by GM-CSF or CSF-1 neutralization (data not shown). We showed earlier (Supplemental Fig. 2E, 2F) that when blood Ly6C+ monocytes were cultured in the presence of GM-CSF, in addition to becoming CD11c+ MHCII+ Ly6C2, they increased in number at the end of the culture period, indicating a mitogenic effect of GM-CSF. We found that the AIP Mo-DCs also proliferated in response to GM-CSF in vitro with CSF-1 at least being able to maintain their number over a 4-d period (Fig. 5C), that is, responses similar to the blood Ly6C+ monocytes (Supplemental Fig. 2F). Comparable in vitro data for the AIP macrophages are also provided.

FIGURE 5. CSF-1 and GM-CSF dependence of MPS population cycling and gene expression. (A) Representative FACS plots showing BrdU labeling in day 3 AIP R1–R3 MPS populations after a short i.p. pulse (2 h) of BrdU. Numbers indicate the frequency of cells within the indicated gates. Data are representative of three experiments; n = 4 mice per group. (B) Frequency of BrdU labeling in day 3 AIP R1 Mo-DC population from C57BL/6 mice treated with isotype, anti–CSF-1, or anti–GM-CSF mAb after a short pulse of BrdU (2 h). Data are mean 6 SEM of three experiments; n = 4 mice/group. One-way ANOVA; isotype versus anti– CSF-1 or anti–GM-CSF. *p , 0.05. (C) Number of sorted AIP R1 MoDCs and R2 macrophages after in vitro culture for 4 d in the absence or presence of CSF-1 or GM-CSF; t0 denotes starting number of cells. Data are mean 6 SEM of triplicate culture from one of two experiments. Paired Student t test; t0 versus untreated, CSF-1, or GM-CSF. *p , 0.05, **p , 0.01, ***p , 0.005. (D) Inflammatory cytokine mRNA expression in FACS-sorted (day 3) AIP R1 Mo-DCs and R2 macrophages after subsequent culture (24 h), in the absence or presence of CSF-1 or GMCSF. Data are mean 6 SEM from one of two experiments.

GM-CSF and CSF-1 can behave as proinflammatory cytokines possibly in part because they can activate MPS cells (2). We have already shown that AIP macrophages from GM-CSF2/2 produce lower levels of proinflammatory cytokines ex vivo than their wildtype counterparts (51). We therefore measured also the CSF effects on some inflammatory cytokine gene expression by AIP macrophages and Mo-DCs in vitro. As shown in Fig. 5D, GM-CSF, but not CSF-1, upregulated IL-6, IL-1b, IL-12p35, and IL-23p19 mRNA expression particularly in the Mo-DCs. Adherent AIP exudate cells gave similar results to the sorted macrophages (data not shown). CSF control of MPS populations in other inflammation models We next determined whether the CSF modulation of the MPS found earlier in AIP occurred more widely in other models of inflammation.

The Journal of Immunology Thioglycolate-induced peritonitis. Another, and the most common, murine peritonitis model is that elicited by thioglycolate. The same phenotyping strategy as earlier indicated that at day 4 the PECs were mainly CD115hi inflammatory macrophages and CD115int SSchi eosinophils (Supplemental Fig. 4A) with only a few neutrophils (data not shown) (45). As in Gautier et al. (45), and unlike the AIP CD11c2 MHCII+ macrophages (Fig. 3A), the thioglycolate-elicited macrophages were mainly CD11c2 MHCII2 (Supplemental Fig. 4A). Even given the CD11c expression in a small proportion of CD115+ cells, a macrophage phenotype in this model has been supported by transcriptomic analysis (45). Unlike in AIP, a distinct population of CD115+ CD11c+ MHCII+ cells (Mo-DCs) was not observed. Similar to the steady-state and AIP peritoneal cavities (Figs. 2A and 3A, respectively), CD11c+ MHCII+ Ly6C2 cDCs could be readily identified in the CD1152 SSc low gate and were also subdivided based on CD11b expression (Supplemental Fig. 4A). Thus, despite both peritonitis models resulting in the recruitment of Ly6C+ monocytes, the developmental fate of these recruited monocytes differed between the two models: the CD115+ CD11c+ MHCII+ MoDC population was only found in the AIP, but not in the thioglycolate-induced peritonitis, whereas inflammatory macrophages were found in both models but differed in their MHCII expression. We reported previously that the exudate macrophage numbers in thioglycolate-induced peritonitis were not altered in GM-CSF2/2 mice or in wild type mice treated with anti–GM-CSF mAb (29, 51). GM-CSF blockade again did not alter the number (Fig. 6A) or surface phenotype (data not shown) of the thioglycolate-induced exudate macrophages. In contrast, we now show that exudate macrophage numbers were reduced by both anti–CSF-1R and anti–CSF-1 mAb treatment, consistent again with CSF-1 signaling being responsible (Fig. 6A). None of the mAbs affected the cDC subsets (data not shown). The lack of involvement of GM-CSF in regulating exudate cell number in thioglycolate-induced peritonitis is consistent with the absence of a GM-CSF–dependent MoDC population in this model noted earlier. LPS-induced lung inflammation. We have shown before that both anti–GM-CSF and anti–CSF-1R mAbs, as judged only by morphology, reduced the number of bronchoalveolar exudate macrophages appearing after i.n. LPS administration, the former mAb also lowering the number of infiltrating neutrophils (29, 63). In this study, we extended these observations by analyzing the population changes in more detail and also tested the effect of anti–CSF-1 mAb administration. The bronchoalveolar exudate cells were divided into three populations based on the literature (23), namely, a Ly6G2 autofluorescent/FITC+ CD11bint CD11chi alveolar macrophage population, a Ly6G2 autofluorescent/FITC+ CD11bhi CD11cint population, mostly containing inflammatory monocyte-derived cells, and Ly6G+ CD11b+ neutrophils (Supplemental Fig. 4B). GM-CSF neutralization led to markedly lower numbers of alveolar macrophages and inflammatory monocyte-derived cells (Fig. 6B), as well as neutrophils (data not shown). Both anti–CSF-1R and anti– CSF-1 mAbs reduced the number of alveolar macrophages and monocyte-derived cells to a similar extent (Fig. 6B), but not neutrophils (data not shown). These results suggest, as discussed earlier, that they are due neither to cell depletion via a CSF-1R–dependent mechanism nor to IL-34 signaling, but it is in fact CSF-1 signaling that is regulating the number of inflammatory macrophages and monocyte-derived cells in this lung inflammation model. LPS-induced splenic inflammation. According to Greter et al. (30), inflammatory Mo-DCs (CD11b+ Ly6C+ F4/80+ CD11c+ MHCII+ Ly6G2) accumulate in the spleen 24 h after i.v. E.coli LPS, but

7

FIGURE 6. CSF-1 and GM-CSF dependence of inflammatory MPS population numbers is context dependent. (A) Number of CD115+ macrophages in day 4 thioglycolate-elicited PECs of C57BL/6 mice treated with isotype, anti–CSF-1R, anti–CSF-1, or anti–GM-CSF mAbs. Data are mean 6 SEM from two experiments; n = 5 mice/group. (B) Number of CD11bint CD11chi alveolar macrophages and CD11bhi CD11cint monocyte-derived cells in bronchoalveolar lavage of BALB/c mice that were given isotype, anti–CSF-1R, anti–CSF-1, or anti–GM-CSF mAb, and challenged with 10 mg LPS i.n. 3 d before analysis. Data are mean 6 SEM from a single experiment; n = 6 mice/group. (C) Number of splenic myeloid populations in naive or i.v. LPS-challenged BALB/c mice (24 h) that were also given i.p. isotype, anti–CSF-1R, anti–CSF-1, or anti–GM-CSF mAb. Data are mean 6 SEM of two experiments; n = 5 mice/group. Unpaired Student t test: isotype versus anti–CSF-1R, anti–CSF-1, or anti– GM-CSF; naive versus LPS. *p , 0.05, **p , 0.01.

independently of a requirement for GM-CSF. We therefore assessed whether the effects of CSF neutralization on the MPS populations in the LPS-induced splenic inflammation would differ from those in LPS-induced lung inflammation, that is, would be tissue specific. In the normal spleen, red pulp macrophages and monocytes, which both reside in the CD64+ Ly6G2 gate, can be distinguished by their differential F4/80 and CD11b profile with the former being CD11bint F4/80hi and the latter CD11bhi F4/80int (28). Splenic monocytes can be divided into a major Ly6C+ and a minor Ly6C2 subset, and the cDCs can be identified as CD642 CD11chi MHCIIhi cells (Supplemental Fig. 4C) (28). After i.v. E. coli LPS administration, splenic red pulp macrophage numbers were unaltered, with anti–GM-CSF blockade making no difference (Fig. 6C) and, consistent with the findings of others in the steady-state, were depleted by anti–CSF-1R (28) (Fig. 6C), with anti–CSF-1 treatment showing a similar effect. Splenic Ly6C+ and Ly6C2 monocyte numbers were also not modulated by i.v. LPS and, as for the steady-state bone marrow and blood, neither population was reduced by anti–GM-CSF blockade, but the Ly6C2 subpopulation was again depleted by anti–CSF-1R and anti–CSF-1 blockade (Fig. 6C, Supplemental Fig. 4D). In the naive spleen, both monocyte subpopulations were MHCII2, but with the Ly6C2 subpopulation expressing more CD11c (Supplemental Fig. 4D);

8

CSF-1, GM-CSF, AND MONONUCLEAR PHAGOCYTE SYSTEM DYNAMICS

upon LPS challenge, both monocyte subpopulations remained MHCII2 , but CD11c increased proportionally on the Ly6C+ subpopulation (Supplemental Fig. 4D). GM-CSF blockade did not alter this change in CD11c expression. Using the current gating approach (Supplemental Fig. 4C) and one previously described (30) (data not shown), we failed to observe the induction of a distinct CD11b+ Ly6C+ CD11c+ MHCII+ Ly6G2 Mo-DC population in the spleen after i.v. LPS challenge (Supplemental Fig. 4D); therefore, unlike in AIP, we could not examine the role of GM-CSF in the development of the population in this model. The CD642 CD11chi MHCIIhi cDCs were reduced after LPS challenge (Fig. 6C), with no difference after treatment with anti–GM-CSF, anti–CSF-1R, or anti–CSF-1 (Fig. 6C, Supplemental Fig. 4D). An LPS-induced neutrophilic expansion was not influenced by any of the mAbs (Fig. 6C).

Discussion Seeing that a major function of CSF-1 and GM-CSF in the MPS is likely to be as prosurvival/growth factors, a major focus of our mAb neutralization approach was on the effect on cell population numbers both in the steady-state and during inflammation. This method also eliminates the inherent developmental defects seen in gene knockout mice and enabled us to explore the distinctive roles of these key cytokines in the adult murine MPS populations. The neutralization of the actions of both CSFs in the same experiments enabled us to compare directly their respective biologies and to address successfully some fundamental issues regarding their functions, as well as of the MPS system, in the steady-state and during inflammation. One conclusion that is apparent is that neutralization of the activity of the respective CSFs leads to different outcomes indicating different roles. In the steady-state, using an anti–CSF-1 mAb by way of comparison, we were able to confirm and extend our prior findings on the rapid monocyte subpopulation and tissue macrophage depletion with the AFS98 (anti–CSF-1R) mAb (29), suggesting, as a result of the similarities found, that the latter mAb was likely to be neutralizing CSF-1 signaling. These data provide an important clarification because it has been concluded by others that the rapid monocyte/macrophage loss caused by AFS98 is more likely to reflect cellular depletion by toxicity or cell clearance via an ADCC mechanism rather than the biology of CSF-1R signaling (27, 31); the anti–CSF-1 mAb data are also important because they suggest that the AFS98 effects observed before (28, 29) and earlier in this article are not due to blockade of the other CSF-1R ligand, IL-34. We suggest that anti–CSF-1 Abs be more widely used than currently done so that CSF-1 and IL-34 biologies can be more clearly defined; this approach also eliminates the risk for nonspecific effects when inhibitors of CSF-1R kinase activity are used (3). Inhibition of CSF-1 action in the steady-state clearly and rapidly depleted peritoneal macrophages and the Ly6C2 monocyte subpopulations in blood, bone marrow, and spleen. These target populations are not cycling to any appreciable extent. The cell disappearance could be due to loss of cell viability and/or residence time due to altered trafficking in the blood or tissue. Interestingly, neither the number of the cycling bone marrow Ly6C+ monocyte and proliferating monocyte populations nor their cycling status was suppressed. We provided at least in vitro evidence for CSF-1 to be a prosurvival factor for both Ly6C+ and Ly6C2 blood monocyte subpopulations. We propose that the lack of depletion of the former subpopulation could be due, in contrast with its relatively long-lasting Ly6C2 counterpart, to its relatively short t1/2 in the circulation (∼18 h) (41), which would not allow sufficient time for any loss of cell viability to be manifested. Even though administered CSF-1 can instruct hemopoietic stem cells to

generate macrophage lineage progeny (25, 26), our data, assuming adequate suppression of CSF-1 action by the anti–CSF-1/CSF-1R mAbs in our neutralization experiments, suggests again (27–29), and somewhat surprisingly, that endogenous CSF-1 is not necessary in the steady-state for the proliferation and differentiation along the macrophage lineage before the Ly6C+ monocyte stage; that is, it acts rather late in this cellular pathway. This conclusion is consistent with only the partial monocytopenia observed in some studies with Csf1op/op and Csf1r2/Csf1r2 mice (28, 29, 64, 65). By monitoring BrdU mean fluorescence intensity levels over time after pulsing, we were able to provide support for the evidence that in the steady-state Ly6C2 monocytes derive from Ly6C+ precursor monocytes in bone marrow and blood, an issue for which there has been some divergent opinion (27, 28, 42, 54, 66). Interestingly, in our hands, the loss of Ly6C from Ly6C+ monocytes in bone marrow and blood still occurred in the presence of anti–CSF-1 mAb, suggesting lack of CSF-1 involvement in this conversion, and this is consistent with our in vitro data with blood Ly6C+ monocytes. The lack of accumulation of the Ly6C+ monocytes in the blood upon CSF-1/CSF-1R neutralization noted earlier in this article and previously (28, 29) is also consistent with this possibility, although the short half-life of the blood Ly6C+ subpopulation mentioned earlier (41) and the high degree of cycling of the bone marrow counterparts may not allow any accumulation to be detected. The lack of effect of GM-CSF depletion, noted previously (29, 51) and earlier in this article, on steady-state monocyte subpopulation numbers in bone marrow and blood, and on resident peritoneal macrophages, does not exclude GM-CSF having a role in homeostasis in barrier tissues such as lung, gut, and skin where it is known to be expressed (30, 55, 67, 68). CSF-1 and GM-CSF neutralization around the time of the initiation of the inflammatory stimulus was chosen for the purpose of assessing the CSF regulation of MPS populations in different inflammation models. In the models where there were MPS exudates (AIP, thioglycolate-induced peritonitis, and LPS-induced lung inflammation), a common feature upon CSF-1/CSF-1R neutralization was reduced cell number across all MPS populations. Thus, these models indicate that CSF-1 neutralization can result in fewer inflammatory macrophages in a lesion, with perhaps significance for clinical approaches (11), but is in disagreement with the findings of others (27, 31). A number of studies have found that CSF-1R and CSF-1 neutralization/depletion can reduce inflammatory/autoimmune disease (2, 10, 29, 69–74), suggesting that targeting of this system may provide therapeutic options. One possible mechanism for the reduction in inflammatory MPS populations found in the earlier models after CSF-1 neutralization is that the removal of a resident macrophage population by such an approach lessens the signal(s) governing overall cellular infiltration; in support of this concept, in the AIP model, fewer CD1152 cells were also noted. Consistent with this proposal, depletion of resident peritoneal macrophages by clodronate-liposomes results in smaller peritoneal exudates (75). Perhaps the end result of anti– CSF-1/CSF-1R treatment on inflammatory cell infiltration is functionally equivalent, at least at this level, to this widely used tissue macrophage depletion strategy. In support of this proposal, resident tissue macrophages have been proposed to participate in the initiation of an inflammatory reaction via their ability to sense and produce proinflammatory cytokines and chemokines (76), and the depletion of perivascular macrophages impairs the recruitment of blood-derived inflammatory cells in a Staphylococcus aureus skin infection model (77). Alternatively, CSF-1 neutralization during an inflammatory reaction, as suggested earlier for the unstimulated peritoneal cavity, may lead to enhanced inflammatory

The Journal of Immunology macrophage apoptosis and/or migration out of the inflamed cavity leading to a faster resolution of the reaction; both mechanisms formulated for macrophages during the normal resolution of thioglycolate-induced peritonitis (45, 78). There has been recent focus on the significant contribution of local macrophage proliferation to their accumulation in inflamed tissues. However, unlike in other murine peritonitis models (17–20) and also, for example, unlike in atherosclerotic lesions (62), local proliferation of AIP macrophages does not appear to be contributing to their increased numbers at least over the short 3-d period, but rather monocyte recruitment is the principal mechanism for the macrophage accumulation; their reduced numbers upon CSF-1 neutralization cannot therefore be due to suppressed proliferation in this model. Besides being a model for other specific Ag-driven, GM-CSF– dependent inflammation models (51, 52) and allowing for the easy isolation of large numbers of purified subpopulations, including from adoptive transfer studies, the AIP model enabled us, in contrast with the widely used thioglycolate-induced peritonitis, to easily detect a putative, cycling inflammatory Mo-DC population. By deliberately selecting the AIP model for a detailed analysis, we were able to demonstrate by our mAb neutralization strategy that GM-CSF can increase the numbers and also modulate in vitro the function of this inflammatory Mo-DC population, at least in this model. Whether GM-CSF has such a role in inflammation/ autoimmunity is currently being debated, with some claiming that it is dispensable for the differentiation of inflammatory DCs (30, 33, 34, 79–81). The dose–response studies with anti–GMCSF mAb in the AIP model allowed us to show that the Mo-DCs appeared to be the most sensitive MPS population to GM-CSF depletion, that is, presumably requiring the highest remaining GM-CSF activity, possibly partly as a result of inhibition of their high degree of cycling (Fig. 5A). This GM-CSF– dependent, cycling AIP Mo-DC appears to be analogous to that noted for lamina propria Mo-DCs (55), and is perhaps a convenient model for the GM-CSF dependence of DC cycling, for example, in early atherosclerotic lesions (82). Interestingly, at least at 48 h under these nonadherent culture conditions, GM-CSF, rather than CSF-1, was mitogenic in vitro for the blood Ly6C+ monocyte subpopulation; CSF-1, however, eventually could expand this monocyte subpopulation in vitro at 5 d under the same conditions (data not shown). We suggest that the potent mitogenic effect of GM-CSF on blood monocytes in vitro and the concomitant modulation of certain differentiation markers, particularly for the Ly6C+ subpopulation, mimics what happens to some infiltrating monocytes when exposed to GM-CSF in the AIP cavity. The in vitro data showing GM-CSF–induced surface marker changes also support our finding that adoptive transfer i.v. of bone marrow monocytes into the AIP model can lead to cells with the Mo-DC phenotype. A high anti–GM-CSF mAb concentration, which is likely to neutralize more GM-CSF than a low mAb concentration, was required to reduce inflammatory macrophage and monocyte numbers in the AIP model, suggesting that, in contrast with the Mo-DCs mentioned earlier, only a low residual GM-CSF activity is needed to regulate their numbers. This observation is consistent with their lowering in GM-CSF2/2 mice (29, 51). It would be worth knowing whether these putative dose-dependent responses to GM-CSF in vivo are due in fact to direct effects on the target population(s) in question (30). We were also able to demonstrate in vitro that GM-CSF could activate AIP Mo-DCs and macrophages to express higher levels of certain proinflammatory genes; some of these GM-CSF–dependent gene products could be controlling the progression of the inflammatory reaction. Although

9 GM-CSF is thought to be important for the development of both CD11b+ and CD11b2 cDC subsets in the normal lamina propria, skin, and lung (30), we found that, unlike in the AIP cavity, GMCSF is dispensable for the presence of the minor peritoneal cDC population in the steady-state, indicating tissue-specific differences. This GM-CSF–independent cDC development in resting tissues was also reported elsewhere in the lung and kidney (83). In summary, our mAb neutralization approach in adult mice has led to the following conclusions/interpretations about the control of the MPS by CSF-1 and GM-CSF in the steady-state and in certain acute inflammation models, an area about which there is current debate. For CSF-1, a role in steady-state monopoiesis could not be demonstrated, it maintains steady-state mature blood monocytes and resident tissue macrophages by promoting their survival and/or altering their trafficking, it is required for the accumulation of maximum inflammatory macrophage numbers and not necessarily by their local proliferation, and it is required for nonspecific inflammatory cell influx possibly via a signal(s) from a resident macrophage population. For GM-CSF, in contrast, our data indicate that it does not control steady-state MPS development in the particular (sterile) locations studied, it can be shown in inflammation to control the development of a cycling population with features of Mo-DCs, it has mitogenic activity in vitro for blood Ly6C+ monocytes in particular, and in vitro it is a mitogen for and can activate Mo-DCs and inflammatory macrophages.

Acknowledgments We thank Jennifer Davis for assistance with the maintenance and care of the mice, Rifa Sallay for assistance with the manuscript, Huei Jiunn Seow and Jonathan Luke McQualter for help with LPS experiments and BD LSRFortessa, and members of Melbourne Brain Centre Parkville Flow Cytometry Facility for cell sorting.

Disclosures The authors have no financial conflicts of interest.

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