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(Journal of Leukocyte Biology. 2002;72:530-537.)
© 2002 by Society for Leukocyte Biology

Macrophage accumulation at a site of renal inflammation is dependent on the M-CSF/c-fms pathway

Yannick Le Meur*, Gregory H. Tesch*,{dagger}, Prudence A. Hill{ddagger}, Wei Mu*, Rita Foti*, David J. Nikolic-Paterson*,{dagger} and Robert C. Atkins*,{dagger}

* Department of Nephrology and
{dagger} Monash University Department of Medicine, Monash Medical Centre, Clayton, Victoria, Australia; and
{ddagger} Department of Anatomical Pathology, St. Vincent’s Hospital, Fitzroy, Victoria, Australia

Correspondence: Dr. Greg Tesch, Department of Nephrology, Monash Medical Centre, 246 Clayton Road, Clayton, Victoria 3168, Australia. E-mail: gtesch{at}hotmail.com


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Production of macrophage-colony stimulating factor (M-CSF), the major macrophage growth factor, is increased in tissues during inflammation. Therefore, we determined whether M-CSF, acting through its receptor c-fms, contributes to macrophage accumulation at a site of tissue injury. Daily treatment with anti-c-fms or control antibody was given to mice with renal inflammation resulting from unilateral ureteric obstruction (UUO). Following UUO, kidney M-CSF mRNA increased in association with macrophage accumulation (days 1, 5, and 10) and local macrophage proliferation (days 5 and 10). Anti-c-fms treatment caused a minor inhibition of monocyte recruitment at day 1, reduced macrophage accumulation by 75% at day 10, but did not affect blood monocyte counts or the CD4 and CD8 lymphocytic infiltrate. Prevention of macrophage accumulation by anti-c-fms treatment was associated with a 90% reduction in local macrophage proliferation at days 5 and 10 without evidence of increased macrophage apoptosis. Therefore, M-CSF/c-fms signaling plays a key role in macrophage accumulation during tissue injury.

Key Words: monocyte • proliferation • cytokine • mouse


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Monocyte/macrophages are key mediators of wound repair, tissue remodeling, and inflammation. The number of macrophages appearing at a site of inflammation is dependent on the nature of the inflammatory stimulus, the severity of injury, and the tissue involved [1 ]. In many diseases, macrophage accumulation has been mostly attributed to the recruitment of circulating blood monocytes and to some extent to prolonged cell survival. This concept is based on the findings that resident tissue macrophages have little proliferative capacity and that macrophage accumulation in response to stimuli that induce a phagocytic response (e.g., heat-killed bacteria, glass, newborn calf serum) occurs through recruitment of blood monocytes with little or no local proliferation [1 ]. It is now well established that recruited monocytes can proliferate locally in some inflammatory diseases such as glomerulonephritis, allograft rejection, pancreatitis, and arthritis [2 3 4 5 6 ]. These studies, based on immunohistochemistry findings, suggest that local macrophage proliferation may be at least as important as recruitment of circulating monocytes in macrophage accumulation during inflammation. Monocyte proliferation outside the bone marrow has also been demonstrated by in vitro studies using monocytes extracted from peripheral blood and glomeruli [7 , 8 ]. However, the relative importance of local proliferation to macrophage accumulation during inflammation and the mechanism of local macrophage proliferation remain to be established.

The mechanisms driving macrophage proliferation at sites of injury are not well defined. Macrophage-colony stimulating factor (M-CSF), also called colony stimulating factor-1 (CSF-1), is likely to be involved since it is the principal factor in the survival and differentiation of cells of the monocyte/macrophage lineage [9 , 10 ]. In vitro, macrophage proliferation is M-CSF-dependent, and it has been shown that macrophages require M-CSF for most of the G1 phase and entry into the S phase of proliferation [11 ]. The hypothesis of in vivo M-CSF-dependent macrophage proliferation is supported by colocalization of M-CSF expression and local macrophage proliferation in human and experimental glomerulonephritis [12 , 13 ]. Evidence that M-CSF-dependent macrophage proliferation is important for the maintenance and accumulation of tissue macrophages is suggested by studies showing that systemically administered M-CSF can restore macrophages to the tissues of CSF-1-deficient op/op mice [14 ], and implantation of M-CSF-producing cells under the kidney capsule incites kidney macrophage accumulation in autoimmune lupus mice [15 ]. This evidence suggests that strategies that prevent the action of M-CSF on macrophages may be valuable for determining the contribution of local proliferation to macrophage accumulation during injury.

M-CSF stimulates macrophages by binding to a single class of the high affinity transmembrane receptor encoded by the c-fms proto-oncogene [16 , 17 ]. Antibodies that bind to c-fms and prevent its interaction with M-CSF are capable of blocking the activation of macrophages by M-CSF [18 ] and may be effective at suppressing the local proliferation of macrophages during inflammation. Unilateral ureteric obstruction (UUO) is a model of renal inflammation, which involves rapid macrophage accumulation associated with increased kidney M-CSF production and local macrophage proliferation [12 , 19 ]. In the current study, we investigated the consequences of blocking the M-CSF/c-fms pathway with an anti-c-fms antibody in the UUO model and determined the effect on local macrophage proliferation and accumulation.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Antibodies
Monoclonal antibodies (mAb) used in this study were: AFS98, rat anti-mouse c-fms [immunoglobulin G (IgG)2a], which neutralizes M-CSF signaling [18 ]; M1/9.3.4, rat anti-mouse CD45 (IgG2a); 5C6, rat anti-mouse Mac1 (IgG2b); GK1.5, rat anti-mouse CD4 (IgG2b); YTS169.4, rat anti-mouse CD8 (IgG2b); and 7/4, rat anti-mouse neutrophils (IgG2a, Serotec, Oxford, UK). The ASF98 hybridoma was kindly supplied by Dr. Shin-Ichi Nishikawa (Graduate School of Medicine, Kyoto University Medical School, Japan). All other hybridomas were purchased from the American Type Culture Collection (Manassas, VA). Antibodies were produced by culturing hybridomas in serum-free medium in i-MAb bags (Diagnostic Chemicals, Charlettetown, Canada). Antibodies for injection were further purified on a Detoxi-gel column (Pierce, Rockford, IL) to remove endotoxin. Some antibodies were biotinylated for use in immunohistochemistry. Immunofluorescence flow cytometry analysis of blood monocytes was performed with fluorescein isothiocyanate (FITC)-conjugated rat anti-mouse CD68 (FA-11, IgG2a, Serotec). Isotype-matched, negative control antibodies were used in all immunostaining experiments.

Analysis of in vivo antibody administration
Groups of four normal male C57/BL6 mice (20–25 g) were given daily intraperitoneal (i.p.) injections of saline, control mAb (M1/9, 50 mg/kg), or anti-c-fms mAb (AFS98, 50 mg/kg) for a period of 5 days. Heparinized blood was then collected from each mouse by cardiac puncture and was analyzed for total white blood cells (WBC) by an automated cell counter (Cell-Dyn 3700, Abbott Laboratories, Abbott Park, IL). The remaining blood was treated with ammonium chloride to remove red blood cells (RBC). The WBC were fixed in 2% paraformaldehyde for 20 min, washed with 0.1% saponin/phosphate-buffered saline (PBS), and then incubated with 5 µg/mL rat anti-mouse CD68-FITC or negative-control mAb-FITC in 0.1% saponin/1% fetal calf serum (FCS)/PBS for 30 min. The percentage of monocytes in each sample was then determined by immunofluorescence flow cytometry. These values were compared with the total WBC of each sample to calculate the amount of monocytes in each sample.

Bone marrow cells extracted from these mice were treated with ammonium chloride to lyse the RBC and then were incubated at 5 x 105 cells/well for 24 h in 24-well plates in the presence of 3H-thymidine (2.5 µCi/well) and media alone (10% FCS/Dulbecco’s modified Eagle’s medium) or media containing 20 ng/mL M-CSF. After incubation, the bone marrow cells were washed, trypsinized, and collected with a cell harvester and the 3H-thymidine uptake was measured by scintillation counting.

Obstructive nephropathy
UUO was performed on male C57/BL6 mice (20–25 g) obtained from Monash University Animal Services (Victoria, Australia). Mice underwent proximal ureteric ligation on the left kidney via a midline abdominal incision. Groups of eight untreated mice were killed at days 1, 5, or 10 after surgery (experiment 1). In addition, groups of eight mice received daily i.p. injections of control mAb (M1/9, 50 mg/kg) or anti-c-fms mAb (AFS98, 50 mg/kg) starting immediately after UUO surgery (day 0) and were killed at days 1, 5, or 10 (experiment 2). Two hours before killing, mice were given an i.p. injection of 50 mg/kg bromodeoxyuridine (Sigma Chemical Co., St. Louis, MO) to label proliferating cells.

Probes
A 720 base-pair cDNA fragment of mature rat M-CSF and a 358 base-pair cDNA fragment of rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were amplified by reverse transcriptase-polymerase chain reaction and were cloned into the pMOSBlue vector (Amersham Pharmacia Biotech, Sydney, Australia). Sense and antisense riboprobes for rat M-CSF and GAPDH were labeled with digoxigenin-uridine triphosphate (DIG-UTP) using a T7 RNA polymerase kit (Roche Biochemicals, Mannheim, Germany).

Northern blotting
Total cellular RNA was extracted from whole kidneys using Trizol (Gibco-BRL, Grand Island, NY). RNA samples (15 µg) were denatured with glyoxal and dimethylsulfoxide, size-fractionated on 1.2% agarose gels, and capillary-blotted onto positively charged nylon membranes (Roche Biochemicals). Membranes were hybridized overnight with DIG-labeled cRNA probes at 68°C in DIG Easy Hyb solution (Roche Biochemicals). After hybridization, membranes were washed and incubated with sheep anti-DIG Ab (Fab) conjugated with alkaline phosphatase. Chemiluminescence substrate (CPD-star, Roche Biochemicals) was then incubated with the membrane, and emissions were captured on Kodak XAR film. The exposed film was analyzed by densitometry using the Gel-Pro Analyzer program (Media Cybernetics, Silver Spring, MD).

In situ hybridization
In situ hybridization to detect M-CSF was performed on formalin-fixed tissue sections using DIG-labeled sense or antisense M-CSF cRNA probe as previously described [20 ]. The hybridized probe was detected using alkaline phosphatase-conjugated sheep anti-DIG IgG and color development with nitroblue tetrazolium (NBT)/5-bromo-4-chloro-3-indolyl phosphate (BCIP; Roche Biochemicals).

Immunohistochemistry staining
Tissue sections (4 µm) from kidneys fixed in 2% paraformaldehyde-lysine-periodate were incubated for 20 min with 0.6% hydrogen peroxide followed by avidin and biotin block (Vector Laboratories, Burlingame, CA) and 10% normal rabbit serum plus 10% normal rat serum to prevent nonspecific detection. Sections were then incubated overnight with 1 µg/ml biotin-conjugated rat anti-mouse Ab, recognizing Mac1, neutrophils, CD4, CD8, or CD45. After washing in PBS, sections were incubated with ABC solution (ABC Kit, Vector Laboratories) for 1 h and were developed with 3,3-diaminobenzidine (DAB; Sigma Chemical Co.) to produce a brown color.

For evaluating macrophage proliferation, tissue sections were immunostained for Mac1 and then microwave-treated at 800W for 12 min in 10 mM sodium citrate (pH 6) to retrieve nuclear antigens, prevent antibody cross-reactivity, and inactivate endogenous alkaline phosphatase [21 ]. Following microwave treatment, sections were maintained at 4°C during incubations with 10% bovine serum albumin (20 min) and primary Ab (overnight) consisting of fluorescein-conjugated antiproliferating cell nuclear antigen (PCNA; 1:100, Roche Biochemicals) or fluorescein-conjugated antibromodeoxyuridine (anti-BrdU; 1:100, Dako, Carpinteria, CA). After washing, sections were incubated at room temperature with alkaline phosphatase-conjugated sheep antifluorescein Fab fragments (1:300, Roche Biochemicals) for 1 h and were developed with Fast Blue BB salt (Sigma Chemical Co.).

For double labeling of c-fms and Mac1, c-fms immunostaining was first performed by sequential incubation with 5 µg/mL ASF98 mAb, 0.6% hydrogen peroxide, avidin and biotin block (Vector Laboratories), biotin-conjugated rabbit anti-rat IgG (1:100, Vector Laboratories), biotin-conjugated goat anti-rabbit IgG (1:100, Vector Laboratories), and ABC complex (Vector Laboratories). After brown color development of c-fms labeling with DAB, tissue sections were treated with 0.6% hydrogen peroxide and then 20% normal rat serum. Tissue sections were then incubated with fluorescein-conjugated Mac1 mAb, followed by horseradish peroxidase-conjugated sheep antifluorescein Fab fragments (1:300, Roche Biochemicals). Mac1 labeling was observed by blue color development with Vector SG (Vector Laboratories).

Quantitation of immunohistochemistry
Tissue macrophage proliferation was determined by two distinct immunostaining methods, double-labeling of Mac1+BrdU+ cells and Mac1+PCNA+ cells. BrdU labels proliferating cells in the S phase of cell cycle, whereas PCNA labels proliferating cells through the late G1-M phase of the cell cycle. The number of macrophages (Mac1+), proliferating macrophages (Mac1+PCNA+, Mac1+BrdU+), T cells (CD4+, CD8+), and total leukocytes (CD45+) in the kidney cortex was assessed in 25 consecutive, high power (x400) cortical fields (representing 30–40% of kidney cortex in the cross-section) by means of a 0.02 mm2 graticule fitted in the eyepiece of the microscope and expressed as cells/mm2 [2 , 3 ]. All scoring was performed on blinded slides.

Apoptosis: deoxy (d)-UTP nick-end labeling (TUNEL) method
An in situ terminal deoxyribonucleotide transferase (TdT)-mediated TUNEL method was used to identify apoptotic cells within kidney tissue sections [22 ]. Formalin-fixed tissue sections (4 µm) were digested with 20 µg/ml proteinase-K for 30 min at 37°C and incubated with TdT and fluorescein-dUTP (Roche Biochemicals) for 1 h at 37°C. Sections were then incubated with 10% normal sheep serum and 10% FCS for 20 min to prevent nonspecific antibody detection. Labeled DNA strands were detected by incubating with alkaline phosphatase-conjugated sheep antifluorescein Fab fragments (1:300, Roche Biochemicals) for 1 h and developing with NBT/BCIP (Roche Biochemicals).

Statistical analysis
Statistical differences between two groups were analyzed by the unpaired Student’s t-test (parametric data) or the Mann Whitney U-test (nonparametric data), and differences among multiple groups of data were assessed by one-way ANOVA. Data were recorded as the mean ± SD, and values of P < 0.05 were considered significant. All analyses were performed using the statistical software in Statview 5.0 (SAS Institute Inc., Cary, NC).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Anti-c-fms mAb does not reduce blood monocytes in vivo
In comparison with saline-treated mice, mouse peripheral blood monocytes (Fig. 1 a ) and total WBC (Fig. 1b) were not different in mice treated for 5 days with neutralizing anti-c-fms mAb or anti-CD45 mAb. Basal proliferation of bone marrow-derived cells was also not different between treatment groups (Fig. 2 ); however, bone marrow cells from mice treated with anti-c-fms antibody had a 22% reduced proliferative response to exogenous M-CSF (Fig. 2) . In addition, the 5-day treatment with anti-c-fms mAb did not affect the number of resident kidney Mac1+ cells (22±4/mm2) compared with treatment with saline (22±7/mm2) or anti-CD45 mAb (25±7/mm2).



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  		Figure 1. Treatment of normal mice with anti-c-fms mAb does not affect circulating levels of monocytes or total WBC. Flow cytometry analysis was used to determine the numbers of CD68+ monocytes (a) and total WBC (b) in normal mice after 5 days of daily injections with saline or 50 mg/kg/day anti-CD45 mAb or anti-c-fms mAb. Data = mean ± SD, n = 4.

 


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  		Figure 2. Anti-c-fms mAb treatment of normal mice reduces M-CSF-induced but not basal proliferation of bone marrow-derived cells. Bone marrow cells were extracted from normal mice after 5 days of daily injections with saline or 50 mg/kg/day anti-CD45 mAb or anti-c-fms mAb. These cells were immediately cultured in the presence or absence of M-CSF in media containing 3H-thymidine, and the cell proliferation was measured after incubation for 24 h. Data = mean ± SD. Data were obtained from the mean of six replicates from each of three mice. *, P < 0.05 versus control treatment.

 
Pathological findings in obstructed kidneys
Following ureter ligation, there was rapid dilatation of the ureter and progressive pathological changes in the tubulointerstitial compartment including tubular atrophy and dilatation, apoptosis and necrosis of tubular cells, accumulation of interstitial monocytes and proliferating cells, and interstitial fibrosis.

M-CSF expression increases with macrophage accumulation in the injured kidney
Constitutive expression of M-CSF was detected in the normal kidney by Northern blotting (Fig. 3 ). Within the normal kidney, in situ hybridization showed that M-CSF mRNA was expressed in only a few glomerular cells (1–2%) and in a small proportion of kidney cortical tubules (15–20%; Fig. 4 a ). Compared with the nonobstructed contralateral kidney, which is unaffected, M-CSF mRNA was significantly increased in the obstructed kidneys of untreated mice at days 1 and 5 (Fig. 3) . In situ analysis detected M-CSF mRNA in 50% of cortical tubules at day 5 and in 80–100% of cortical tubules at day 10 within untreated, obstructed kidneys (Fig. 4b) . The glomerular expression of M-CSF mRNA did not alter during disease compared with normal.



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  		Figure 3. M-CSF mRNA is increased in obstructed mouse kidneys. (a) Northern blot showing M-CSF mRNA in normal mouse kidneys, contralateral (cont), nonobstructed kidneys from UUO mice (days 1 and 5), and obstructed kidneys from UUO mice (days 1 and 5). (b) Graph indicating the ratio of M-CSF to GAPDH mRNA. Data = mean ± SD, n = 4, **, P < 0.0001 versus normal.

 


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  		Figure 4. M-CSF-dependent macrophage proliferation and accumulation in obstructed mouse kidneys are reduced by anti-c-fms antibody treatment. In situ hybridization showing M-CSF mRNA (purple) in normal mouse kidney tubules (arrows; a) and increased expression in an obstructed mouse kidney 10 days after ureter ligation (b). Immunoperoxidase staining detected many peritubular Mac1+ macrophages (brown) in obstructed kidneys at day 10 of disease in mice treated with control antibody (c), which were largely reduced by treatment with anti-c-fms antibody (d). Double immunostaining showed that c-fms (brown) expression was restricted to macrophages (blue-gray) in obstructed kidneys (e). TUNEL staining identified apoptotic cells in the tubules (arrow) and the interstitium (arrowheads) of obstructed kidneys at day 5 (f). Proliferating macrophages (arrows) were readily detected in the obstructed kidneys of control antibody-treated mice at day 10 by double immunostaining for macrophages (brown) and cell proliferation (blue nuclei) identified by expression of PCNA (g) or BrdU (h). Proliferating cells were also detected in tubules (arrowheads, g and h). Original magnification: a and b, x100; c–f, x200; g and h, x400.

 
Immunostaining for Mac1 allowed assessment of progressive, interstitial macrophage accumulation in obstructed kidneys, as neutrophils that also express Mac1 were present in small numbers in obstructed kidneys and did not increase in number during the development of renal injury (day 1: 42±6; day 5: 48±8; day 10: 52±11 neutrophils/mm2). Interstitial Mac1+ cells were present in small numbers in the cortex of normal kidneys (22±7 Mac1+ cells/mm2). In comparison, there was a marked increase in interstitial Mac1+ cells in the cortex of untreated, obstructed kidneys at day 1 (110±14 Mac1+ cells/mm2), which progressed rapidly at days 5 (590±131 Mac1+ cells/mm2) and 10 (1572±224 Mac1+ cells/mm2). Double immunostaining for Mac1 and c-fms in untreated, obstructed kidneys indicated that only macrophages expressed c-fms during disease progression (Fig. 4e) .

Anti-c-fms treatment reduces macrophages but not T lymphocyte accumulation in the injured kidney
Leukocyte accumulation in the obstructed kidneys of mice treated with control antibody was comparable with untreated mice. In contrast, treatment of mice with anti-c-fms antibody resulted in a minor inhibition of kidney monocyte recruitment at day 1 of obstruction and largely suppressed the progressive (4- to 12-fold) increase of total leukocytes and macrophages in obstructed kidneys at days 5 and 10 (Figs. 4d and 5, a and b). This inhibition of interstitial macrophage accumulation was verified by semiquantitative assessment of mononuclear cell infiltrate on histochemically stained sections at day 10 (infiltrate score: control Ab treatment, 1.8±0.8 vs. anti-c-fms Ab treatment, 0.8±0.5, P<0.05). In contrast, the CD4 and CD8 lymphocyte accumulation in obstructed kidneys of mice treated with anti-c-fms antibody was not different with mice treated with control antibody (Fig. 5c and 5d) .



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  		Figure 5. Macrophage but not T lymphocyte accumulation in obstructed kidneys is reduced by anti-c-fms Ab treatment. Immunostaining was used to detect the accumulation of (a) CD45+ total leukocytes, (b) Mac1+ macrophages, (c) CD4+ T lymphocytes, and (d) CD8+ T lymphocytes in the obstructed kidneys of control Ab and anti-c-fms Ab-treated mice during the progression of inflammation. Compared with control treatment, anti-c-fms Ab reduced the levels of CD45+ and Mac1+ cells but not CD4+ and CD8+ cells in obstructed kidneys at days 5 and 10 of injury. Data = mean ± SD, n = 8, **, P < 0.0001 versus control treatment.

 
Anti-c-fms treatment prevents macrophage proliferation without increasing macrophage apoptosis
At day 1 after obstruction, there were no proliferating macrophages detected within the obstructed or nonobstructed contralateral mouse kidneys. At days 5 and 10, many interstitial macrophages (Mac1+) were found to be proliferating (7–9% BrdU+, 13–14% PCNA+) in the obstructed kidneys of mice treated with anti-CD45 control antibody (Fig. 4g and 4h) , but macrophage proliferation was absent in the contralateral kidneys. In these control mice, macrophage accumulation in the obstructed kidneys strongly correlated with macrophage proliferation (Mac1 vs. Mac1+BrdU+, r=0.93, P<0.001, and Mac1 vs. Mac1+PCNA+, r=0.88, P<0.001). In comparison, mice treated with anti-c-fms antibody had a large reduction in macrophage proliferation in obstructed kidneys (Fig. 6a and b). Anti-c-fms treatment reduced Mac1+BrdU+ kidney cells by 89% at day 5 and 92% at day 10 and Mac1+PCNA+ kidney cells by 88% at day 5 and 90% at day 10 of disease.



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  		Figure 6. Macrophage proliferation in obstructed kidneys is prevented by anti-c-fms Ab treatment. Immunostaining was used to detect the accumulation of (a) Mac1+BrdU+ proliferating macrophages and (b) Mac1+PCNA+ proliferating macrophages during the progression of inflammation in obstructed kidneys. Macrophage proliferation was prominent at days 5 and 10 in the obstructed kidneys of control Ab-treated mice and was largely prevented by treatment with anti-c-fms Ab. Data = mean ± SD, n = 8, **, P < 0.0001 versus control treatment.

 
TUNEL staining was performed to identify whether the reduced number of kidney macrophages following anti-c-fms treatment could be attributed to increased macrophage apoptosis. Apoptosis was detected in 5 ± 2% of tubular epithelial cells in obstructed kidneys at days 5 and 10 and was not affected by anti-c-fms treatment. In contrast, interstitial, apoptotic cells were rare (2±1/mm2) in the obstructed kidneys and were not different between control antibody-treated (Fig. 4f) and anti-c-fms antibody-treated mice, suggesting that anti-c-fms treatment does not reduce macrophage accumulation in obstructed kidneys by inducing macrophage apoptosis.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The accumulation of macrophages within tissues during inflammation is known to be dependent on a dynamic balance of factors regulating the recruitment, proliferation, and removal of these cells [23 ]. However, the heirachy of importance of each of these components in the gathering of macrophages at an inflammatory site is poorly understood. Previous animal studies have suggested that recruitment of circulating blood monocytes into tissues is the major mechanism responsible for macrophage accumulation at inflammatory sites [1 ]. The current study now provides the first direct evidence that local macrophage proliferation can also play an important role in macrophage accumulation during tissue inflammation.

Blockade of the M-CSF/c-fms signaling pathway with an anti-c-fms antibody caused a major reduction in kidney macrophage accumulation during ureteric obstruction. Anti-c-fms antibody treatment did not reduce normal levels of blood monocytes or basal bone marrow cell proliferation and had only a minor inhibitory effect on bone marrow cell proliferation induced by exogenous M-CSF, indicating that the proliferative capacity of bone marrow monocytes is not adversely affected by anti-c-fms mAb treatment and suggesting that the inhibition of macrophage accumulation within the kidney is mostly a result of local suppression. Our analysis attributed the reduction in kidney macrophages to a 90% inhibition of macrophage proliferation observed at days 5 and 10 of injury. The importance of M-CSF-driven macrophage proliferation in macrophage accumulation in this model is emphasized by our findings that kidney M-CSF expression increases with progression of inflammation; infiltrating interstitial macrophages reside in close proximity to tubular cells, which are the major source of kidney M-CSF in this model [12 ]; the M-CSF receptor (c-fms) is expressed only on macrophages in the obstructed kidney; and blockade of M-CSF/c-fms signaling results in a 75% reduction of interstitial kidney macrophages at day 10 after ureteric obstruction.

The major action of M-CSF during ureteric obstruction appears to be the promotion of local macrophage proliferation; however, we also investigated the involvement of M-CSF in macrophage recruitment. Macrophage migration activity in obstructed kidneys has been shown to peak between 4 and 12 h after ureteric ligation and to decline thereafter [24 ]. During obstruction, early macrophage accumulation is associated with increased expression of several molecules that promote macrophage recruitment directly [monocyte chemoattractant protein-1 (MCP-1), osteopontin, intercellular adhesion molecule-1, vascular cell adhesion molecule-1] or indirectly (angiotensin II) [19 , 25 , 26 ]. Although in vitro studies have indicated that M-CSF may act as a chemoattractant [27 ] or may indirectly promote production of MCP-1 [28 , 29 ], M-CSF is generally considered a minor contributor to macrophage recruitment during inflammation. Our study revealed that M-CSF plays a small but significant role in early macrophage recruitment. At day 1 of obstruction, we detected a prominent macrophage infiltrate but no proliferating macrophages in the obstructed kidney. Anti-c-fms antibody treatment caused a minor (40%) inhibition of Mac1+ cells recruited into obstructed kidneys at day 1 compared with control antibody treatment. Therefore, M-CSF is involved in early macrophage recruitment during inflammation, presumably by promoting the chemoattraction or adhesion molecule interactions of infiltrating monocytes at the site of injury. However, macrophage accumulation in obstructed kidneys proceeds rapidly following the initial infiltration and correlates closely with macrophage proliferation, suggesting that the contribution of M-CSF to macrophage accumulation through recruitment is relatively small compared with its ability to promote macrophage accumulation through inducing local proliferation.

Several studies have shown that M-CSF/c-fms signaling is important for monocyte survival [9 , 10 ]. Addition of M-CSF to cultured blood monocytes promotes their proliferation and maturation and suppresses monocyte apoptosis [9 , 10 ]. Evidence that M-CSF signaling controls macrophage apoptosis is suggested by studies demonstrating that the Fc receptor for IgG cross-linking of IgG on monocytes protects against apoptosis by inducing M-CSF release [30 ], and incorporation of the Bcl-2 transgene into op/op mice replenishes tissue macrophages [31 ]. In previous work from our laboratory [22 ], we identified colocalization of apoptotic and proliferating macrophages within focal kidney lesions, which suggests that limiting M-CSF signaling may be an important control mechanism for proliferation-dependent macrophage accumulation. In contrast to our expectations, anti-c-fms treatment of obstructed kidneys did not result in an increase in the detection of apoptotic macrophages. This finding suggests that blockade of c-fms signaling is critical for the proliferation of tissue macrophages but not for the prevention of apoptosis. Within the local inflammatory site, other signals from cytokines or interactions with cell adhesion molecules or extracellular matrix may prevent or delay apoptosis by promoting macrophage differentiation [32 ] and expression of antiapoptotic factors [33 ], allowing the macrophages to exit into local draining lymph nodes before dying [34 ]. Alternatively, it is possible that macrophages undergoing apoptosis in focal lesions may be phagocytosed so rapidly by neighboring cells that it is impossible to detect increased macrophage apoptosis in these lesions by current methods. Therefore, a role for M-CSF signaling in the prevention of macrophage apoptosis within an inflammatory lesion, although not evident, cannot be excluded.

Conclusive proof of a role for M-CSF in inflammation has been difficult to obtain. M-CSF deficient op/op mice have low levels of circulating blood monocytes [35 ] and are a useful tool for examining the effects of macrophage depletion in models of tissue injury [29 , 36 , 37 ] but are not suitable for determining the requirement of locally produced M-CSF for macrophage accumulation at sites of inflammation. In addition, young op/op mice (3 weeks) have fewer tissue-specific macrophages compared with normals, but acquire similar numbers as they mature (4 months), which is associated with increased local interleukin-3 production [38 ]. Therefore, it appears that op/op mice can develop compensatory mechanisms to overcome their lack of M-CSF production. A better strategy for defining the role of M-CSF in inflammation is to prevent M-CSF signaling in mice with normal levels of blood monocytes. A recent study showed that a neutralizing anti-c-fms mAb could reduce the accumulation of macrophage-derived foam cells in the aortic root of apoE-deficient mice maintained on a high-fat diet [18 ]. Using the same mAb, our current study provides definitive proof that M-CSF signaling is responsible for local macrophage proliferation within an inflammatory lesion, supporting a role for locally produced M-CSF in promoting inflammation.

In summary, this study has demonstrated that prevention of local macrophage proliferation via blockade of the M-CSF/c-fms signaling pathway can result in a large reduction in macrophage accumulation during an inflammatory response, demonstrating that local proliferation can be a major contributor to macrophage accumulation during inflammation. The results also indicate that blockade of the M-CSF/c-fms signaling pathway is a highly effective and selective strategy for suppressing macrophage proliferation and accumulation in vivo without depleting levels of circulating blood monocytes. Consequently, therapeutic strategies targeting the M-CSF/c-fms signaling pathway may provide protection against macrophage-mediated injury in inflammatory diseases.


   ACKNOWLEDGEMENTS
 
This work was supported by a grant from the National Health and Medical Research Council of Australia and by a fellowship from the Fondation pour la Recherche Medicale, Paris, France (Y.L.).

Received May 6, 2002; accepted May 6, 2002.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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