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Augments Acute Macrophage-Mediated Renal Injury Via a Glucocorticoid-Sensitive Mechanism
*Department of Nephrology and Monash University Department of Medicine, Monash Medical Centre, Clayton, Victoria, Australia.
Correspondence to Dr. David J. Nikolic-Paterson, Department of Nephrology, Monash Medical Centre, Clayton Road, Clayton, Victoria 3168, Australia. Phone: 61-3-9594 3535; Fax: 61-3-9594 6530;
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Abstract |
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(IFN-) or dexamethasone (Dex), and then macrophage-mediated renal injury determined in vivo. In this model, rats were made leukopenic by administration of cyclophosphamide (CyPh). Two days later (day 0), animals were injected with sheep anti-GBM serum followed by a single injection of rat NR8383 macrophages on day 1 and then killed 3 or 24 h after cell transfer. NR8383 macrophages were incubated IFN- and/or Dex before adoptive transfer into animals. Induction of proteinuria and glomerular cell proliferation (PCNA+ cells) in this model was dependent on transfer of NR8383 macrophages. Exposure of macrophages to IFN- for 18 h (but not 3 h) before transfer caused a twofold increase in the degree of proteinuria and glomerular cell proliferation compared with unstimulated cells (Nil versus IFN-; P < 0.001). This was due to an increase in the number of transferred macrophages within the glomerulus and a significant increase in degree of renal injury per transferred glomerular macrophage. IFN- increased iNOS and PDGF-B gene expression and upregulated adhesion molecule expression in NR8383 macrophages. In contrast, exposure of NR8383 cells to Dex for 18 h (but not 1 h) abrogated renal injury due to a failure of transferred macrophages to accumulate within the glomerulus. In addition, Dex abrogated renal injury caused by IFN-–stimulated macrophages. In conclusion, activation of macrophages by IFN-, independent of any effect on other leukocytes or renal cells, can substantially augment macrophage-mediated renal injury. This IFN- augmentation of renal injury is sensitive to the action of glucocorticoids, which act directly on macrophages to prevent their recruitment to the inflamed glomerulus. This study provides the first evidence that it is possible to directly modulate macrophage-mediated renal injury. E-mail: David.Nikolic-Paterson@med.monash.edu.au |
Introduction |
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Many macrophage functions require the cell to be "primed" or "activated," such as interferon- (IFN-) priming for maximal nitric oxide production in response to bacterial challenge. However, we know very little of how activation affects the ability of macrophages to cause renal injury. A number of macrophage activating cytokines, such as IFN-, macrophage migration inhibitory factor, interleukin-1, and tumor necrosis factor-
, are produced within the inflamed kidney, and blockade of these cytokines inhibits macrophage accumulation and renal injury in experimental glomerulonephritis (8–14 ).
These studies suggest that modulation of macrophage activation may be an important therapeutic goal in the treatment of glomerulonephritis. However, systemic cytokine blockade cannot distinguish between direct effects on macrophage activation and indirect effects on other infiltrating cells (T cells, neutrophils) or intrinsic renal cells (endothelium, mesangial cells, podocytes, tubular epithelial cells). Therefore, to investigate the importance of macrophage activation in the mediation of renal injury, it is necessary to manipulate macrophage activation independently of all other cells within the kidney. We addressed this question by using an adoptive transfer model of macrophage-mediated renal injury. This makes it possible to manipulate macrophages in vitro before transfer into recipient animals, thus enabling an assessment of their capacity to cause renal injury. IFN- was used to activate macrophages, while dexamethasone (Dex) was used to inhibit macrophage activation.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Macrophages were incubated with 30 µg/ml of bromodeoxyuridine (BrdU) during the last 24 h of culture, washed three times in PBS, and then transferred to recipient animals. Immunochemical staining of cell spots showed that BrdU was incorporated into DNA in approximately 70% of the cells, and this label was used to identify cells after transfer into recipient animals.
Analysis of Nitric Oxide (NO) Production
NR8383 macrophages were cultured in media alone or with recombinant rat IFN- (PeproTech Inc., Rocky Hill, NJ), Dex (David Bull Laboratories, Warwick, UK), or IFN- plus Dex at a range of concentrations for between 1 and 18 h. Cells then were washed three times in PBS, and 1 x 106 cells cultured in 24-well plates in 0.5 ml of media for 24 h with or without IFN- stimulation.
The accumulation of nitrite, a stable metabolite of nitric oxide, was measured in cell culture supernatants by the Griess assay. Briefly, 50 µl of supernatant was combined with an equal volume of Griess agent and incubated at room temperature for 10 min. The colored reaction product was spectrophotometry at 550 nm with a reference calibration curve of known concentrations of sodium nitrite. Data is expressed as nanomoles of nitrate secreted by 106 cells in a 24-h period.
Adoptive Transfer Model of Macrophage-Mediated Renal Injury
A modified version of rat anti-GBM disease in inbred male Sprague-Dawley rats (150 to 180 g body wt) was used as described previously (7). Animals were made leukopenic (
98% reduction in white blood cell numbers) by a single intraperitoneal injection of 165 mg/kg cyclophosphamide (CyPh) 60 h before intravenous injection of sheep anti-rat GBM serum (day 0). Transfer of NR8383 macrophages (6 x 107 cells per animal) was given by tail vein injection on day 1. Animals were either killed 3 h after macrophage transfer (to assess glomerular macrophage recruitment) or placed in metabolic cages for a 24-h urine collection and then killed (to assess renal injury).
The NR8383 macrophages were incubated with medium alone, 1000 U/ml IFN-, 1 µM Dex, or IFN- plus Dex, and then washed three times before transfer into recipient animals. The same amount of NR8383 cells was transferred into each animal irrespective of treatment. Washing of cells prevented any possible carryover of IFN- or Dex into the recipient animals. The effect of IFN- and Dex on activation of NR8383 macrophages was confirmed in all experiments by incubating 1 x 106 of the treated cells in 0.5 ml of culture medium and measuring nitric oxide production over the next 24 h.
Biochemical Analyses
Protein excretion in 24-h urine collections was determined using the benzethonium chloride method. Whole blood cell counts were performed on a Cell-Dyn 3500 automated cell counter (Abbott Laboratories, Abbott Park, IL) using heparinized blood collected from tail veins. All analyses were performed by the Department of Biochemistry, Monash Medical Center.
Antibodies
The following monoclonal antibodies (mAb) were used in this study: ED1, anti-rat CD68 labels monocytes and macrophages (16); PC-10, anti-proliferating cell nuclear antigen (PCNA) labels cells in the G1, S, and G2 phase of the cell cycle (Dako Ltd, Glostrup, Denmark); M744 mAb, anti-BrdU (Dako Ltd); OX-7, anti-Thy-1 antigen (CD90), which labels rat thymocytes and glomerular mesangial cells (17,18 ); OX-42, anti-rat CD11b/c (iC3b) (19); 1A29, anti-rat ICAM-1 (CD54) (Serotec, Oxford, UK) (20); WT.3, anti-rat CD18 (LFA-1 
-chain) (Serotec); and 73.5, mouse monoclonal antibody against human leukocytes that does not react with rat tissues, was used as negative control. Peroxidase- and alkaline phosphatase-conjugated goat anti-mouse IgG, peroxidase-conjugated mouse anti-peroxidase complexes (PAP), and alkaline phosphatase-conjugated mouse anti-alkaline phosphatase complexes (APAAP) were purchased from Dako Ltd. FITC-conjugated sheep anti-mouse IgG F(ab')2 fragment (AMRAD Biotec., Boronia, Victoria, Australia) was used in flow cytometry.
Immunohisotochemistry
Immunohistochemical staining to detect ED1+ macrophages was performed on paraffin sections (4 µm) of methylcarn-fixed tissue using a three-layer peroxidase-based detection method, as described previously (21). In brief, sections were dewaxed, hydrated in PBS, heated for 10 min in 0.1 M sodium citrate, pH 6.0, in a microwave oven, washed in PBS, blocked with 10% FCS and 10% normal sheep serum in PBS for 30 min, drained, incubated with the ED1 mAb in 10% normal rat serum (NRS), 1% BSA in PBS overnight at 4°C, washed in PBS, and then endogenous peroxidase–inactivated by incubation in 0.3% H2O2 in methanol. Sections then were washed in PBS, incubated with peroxidase-conjugated goat anti-mouse IgG, washed in PBS, incubated with mouse PAP complexes, and developed with 3,3-diaminobenzidine to produce a brown color. Immunostaining with the PC-10 (PCNA) antibody used the same protocol, except that the PAP complex amplification step was omitted.
Transferred macrophages were detected by two-color immunostaining (ED1+BrdU+ cells) in paraffin sections of formalin-fixed tissue. Staining for BrdU using the M744 mAb was performed as described for the ED1 mAb above and were then given a second round of microwave oven heating to block antibody crossreactivity, inactivate endogenous alkaline phosphatase, and enhance detection of the CD68 antigen (21). After blocking as above, sections were incubated with the ED1 mAb, incubated sequentially with alkaline phosphatase-conjugated goat anti-mouse IgG and mouse APAAP complexes, and developed with Fast Blue BB Salt (Ajax Chemicals, Melbourne, Australia). Slides were counterstained by PAS.
Proliferating mesangial cells (OX-7+PCNA+ cells) were detected in 4-µm sections of methylcarn-fixed paraffin tissues by two-color immunostaining with the OX-7 mAb followed by the PC-10 mAb using the method as described above for BrdU/ED1 staining, except that microwave oven heating was not used before OX-7 staining.
The number of ED1+ macrophages or PCNA+ proliferating cells was counted in at least 50 glomerular cross-sections (gcs) per animal. All scoring was performed on blinded slides.
Flow Cytometry
NR8383 macrophages were incubated in the presence or absence of 1000 U/ml IFN-, 1 µM Dex, or IFN- plus Dex for 18 h. Cells were harvested using 0.02% EDTA/PBS and then resuspended in ice-cold PBS containing 1% FCS and 0.02% NaN3. To detect adhesion molecule expression, cells were incubated with primary antibodies for 60 min, washed three times, and then incubated with FITC-conjugated sheep anti-mouse IgG diluted in 1% FCS, 10% heat-inactivated NRS, 0.02% NaN3 in PBS. The 73.5 mAb was used as an isotype-matched negative control. Cells were analyzed using a FACScan flow cytometer (Cytomation, Fort Collins, CO) equipped with Cylops SUMMIT software.
Macrophage Apoptosis
Macrophage apoptosis was assessed by flow cytometry using FITC-conjugated annexin V and propidium iodide as described previously (22). NR8383 macrophages cultured with IFN-, Dex, IFN- + Dex, or LPS were washed and resuspended in 100 µl of annexin V binding buffer (Clontech laboratories, Inc., Palo Alto, CA). Cells then were incubated with FITC-conjugated annexin V (final concentration, 0.5 µg/ml) (ClonTech) and 5 µg/ml propidium iodide (PI, Sigma) for 15 min in the dark. Samples then were diluted with 400 µl of binding buffer and analyzed by flow cytometry. The analysis discriminates between viable cells (Annexin V- PI-), apoptotic cells (Annexin V+ PI-), and dead cells (PI+).
Reverse-Transcription (RT)-PCR
NR8383 macrophages (1 x 106 cells/well) were incubated in medium alone or with 1000 U/ml IFN-, 1 µM Dex, or IFN- plus Dex for 18 h. Total RNA then was extracted using the TRIZOL reagent (Life Technologies BRL, Gaithersburg, MD), and cDNA was reverse transcribed from 5 µg of RNA using SuperScript First-Strand Synthesis System (Life Technologies BRL).
Amplification was carried out using the GeneAmp PCR system 2400 (Perkin Elmer, Burwood, Victoria, Australia) through 18 to 40 cycles of denaturation at 94°C for 30 s, annealing at individual temperatures for 30 s, and extension at 72°C for 1 min. A point approximately halfway through the logarithmic amplification curve was selected for semiquantitative analysis. Amplification of the housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), was used as the control for RNA extraction and reverse transcription.
Primers used were: iNOS sense, 5'-TGA AAG AAC TCG GGC ATA CC-3'; iNOS anti-sense, 5'-AAC AAT CCA CAA CTC GCT CC-3'; RT1B (rat MHC Ia) sense, 5' -TAG AGA ACA GAG ATG CCG CT-3'; RT1B anti-sense, 5'-ACC CTT ACC TTC TTT CCC AG-3'; IFN- sense, 5'-GAT TCT TCG GAC TCT CTG AC-3'; IFN- anti-sense, 5'-TAT TGG CAC ACT CTC ATC CC-3': PDGF-B sense, 5'-GAA GCC AGT CTT CAA GAA GGC CAC-3'; PDGF-B anti-sense, 5'-AAC GGT CAC CCG AGT TTG AGG TGT-3'; IL-1 sense, 5'-GCT ACC TAT GTC TTG CCC GT-3'; IL-1 anti-sense, 5'-GCC GTC TTT CAT CAC ACA GG-3'; GAPDH sense, 5'- AAA GGG TCA TCA TCT CCG CC-3'; GAPDH anti-sense, 5'-CCT GCT TCA CCA CCT TCT TG-3'.
Products were run on 1.2% agarose gels containing ethidium bromide and bands recorded using the Gel Doc 2000 system (Bio-Rad Laboratories, North Ryde, New South Wales, Australia). Densitometric analysis of the DNA bands used the GelPro 3.0 analyzer program (Media Cybernetics, Silver Spring, MD). Results were expressed as a ratio to the GAPDH band.
Statistical Analyses
Statistical analyses were performed using GraphPad Prism 3.0 (GraphPad Software, San Diego, CA/). Comparison of three groups or more used one-way ANOVA, and individual group means were compared post test with the Tukey Multiple Comparison Test. Correlation between variables was determined with the Pearson single correlation coefficient. All values are expressed as mean ± SD.
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IFN- Activates NR8383 Macrophages In Vitro
The addition of rat recombinant IFN- caused a dose-dependent activation of NR8383 macrophages as shown by secretion of nitric oxide (Figure 3a). A dose of 1000 U/ml, which gave maximal macrophage activation, was selected for subsequent studies. To evaluate the time required for IFN- to activate macrophages, cells were incubated with IFN- for different periods and washed, and then nitric oxide production measured over the next 24 h. A 3-h stimulation with IFN- caused significant macrophage activation, with maximal activation seen with an 18-h stimulation (Figure 3b). Activation of NR8383 macrophages in response to an 18-h stimulation with IFN- was also characterized by upregulation of: iNOS mRNA, RT1B mRNA, cell surface expression of leukocyte adhesion molecules, and PDGF-B mRNA (Table 1, Figure 4).
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Augments Macrophage-Mediated Renal Injury activation of NR8383 macrophages for 3 h before transfer had no effect on glomerular macrophage accumulation or the degree of renal injury compared with macrophages cultured in medium alone (Figure 1, a through c). However, IFN- activation of NR8383 macrophages for 18 h before transfer resulted in a twofold increase in the degree of proteinuria and glomerular cell proliferation (Figure 1, b and c). There was an excellent correlation between the number of transferred IFN- activated macrophages and the degree of proteinuria (r2 = 0.82, P = 0.0003) and glomerular cell proliferation (r2 = 0.83, P = 0.0003).
The 18-h IFN- activation caused a significant increase in glomerular macrophage accumulation 24 h after transfer (Figure 1a). We also examined the initial glomerular macrophage recruitment at 3 h after transfer and found a small, but significant increase in the accumulation of IFN-–activated macrophages relative to unstimulated cells (Figure 5). Glomerular macrophage accumulation in rat anti-GBM disease is dependent on leukocyte adhesion molecule expression (23). Therefore, we examined cell-surface adhesion molecule expression in NR8383 macrophages by flow cytometry. IFN- activation resulted in a significant increase in expression of ICAM-1 and CD11b/c but had no effect on CD18 (Table 1).
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–activated macrophages induced a significantly higher level of proteinuria and glomerular cell proliferation per transferred macrophage (Figure 6). Next, we examined gene expression of molecules that have been shown to play a pathogenic role in experimental glomerulonephritis (13,24–26 ). Semiquantitative PCR demonstrated that 18 h of IFN- stimulation of NR8383 macrophages caused a significant increase in mRNA expression for iNOS and PDGF-B but did not change IL-1 mRNA levels (Figure 4).
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activation of NR8383 macrophages in dose-dependent fashion (Figure 3c). We selected a 1 µM dose to given optimal inhibition of macrophage activation in subsequent experiments. We also found that Dex needs to be added within 6 h of IFN- stimulation to substantially inhibit macrophage activation, with maximal effect being seen when Dex was added together with IFN- (Figure 3d).
Dexamethasone Inhibits Macrophage-Mediated Renal Injury
Exposure of NR8383 macrophages to Dex for 1 h had no effect on glomerular macrophage accumulation or the induction of renal injury (Figure 1, d through f). However, 18 h of exposure to Dex abolished macrophage-mediated renal injury (Figure 1, e and f). This was attributed to abrogation of glomerular macrophage accumulation assessed at 24 h after transfer (Figure 1d). Examination at 3 h after transfer showed a significant reduction in glomerular recruitment of Dex-treated macrophages compared with unstimulated cells, although a minor infiltrate of transferred cells was still evident (Figure 5). This inhibition of glomerular macrophage accumulation was not due to induction of macrophage cell death. Control NR8383 macrophages were 90.7 ± 2.1% viable as determined by flow cytometry using annexin V and propidium iodide staining. Macrophages remained 89 to 91% viable after 18 h of stimulation with Dex or IFN- plus Dex.
Dexamethasone Inhibits Renal Injury Induced by IFN- Activated Macrophages
Addition of Dex at the same time as IFN- inhibited macrophage activation in vitro as assessed by nitric oxide production (Figure 3d) and gene transcription of iNOS, PDGF-B, and IL-1 (Figure 4). Dex was also shown to abolish renal injury induced by IFN-–treated macrophages in vivo (Figure 1, e and f). This was attributed to inhibition of glomerular accumulation of transferred macrophages (Figure 1d), without any effect on macrophage viability. In addition, Dex abolished IFN- upregulation of macrophage expression of leukocyte adhesion molecules (Table 1).
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Discussion |
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and Dex on macrophage-mediated renal injury independently of any effects of these treatments on other immune cells or intrinsic renal cells within the inflamed kidney. We have demonstrated, for the first time, that modulation of macrophage activation can exert a major influence on the ability of macrophages to cause renal injury.
Macrophage stimulation with IFN- for an 18-h period significantly augmented macrophage-mediated renal injury. This effect operated through two mechanisms. First, there was an increase in the number of glomerular macrophages 24 h after transfer. This was due to both an increase in glomerular macrophage recruitment at 3 h after transfer and an increased retention of macrophages within the glomerulus up to 24 h after transfer. This may have been, in part, due to an increase in cell surface expression of leukocyte adhesion molecules on IFN- stimulated NR8383 macrophages at the time of transfer. It has also been suggested that autocrine production of IFN- by infiltrating macrophages is critical for the development of macrophage infiltration and development of lupus nephritis in IFN--/- MRL/lpr mice (27). Although, whether the IFN- producing cells seen in the kidney 4 mo after adoptive transfer of IFN-+/+ Mac-1+ spleen cells into IFN--/- mice were macrophages or some other cell type(s) arising from the transfer was not determined (27). IFN- production by NR8383 cells is not a mechanism for increased glomerular recruitment of transferred macrophages in our study since IFN- mRNA was detected by RT-PCR in endotoxin-stimulated NR8383 cells, but not in unstimulated or IFN-–stimulated cells (data not shown). Another possible reason for increased glomerular retention of transferred cells is inhibition of macrophage apoptosis. We have previously identified apoptosis of glomerular macrophages in rat anti-GBM disease, suggesting that this is an important mechanism for the removal of macrophages from the inflamed glomerulus (28), and IFN- has been shown to inhibit macrophage apoptosis (29). The second mechanism to account for IFN- augmentation of renal injury is that IFN- activation caused a significantly greater degree of renal injury (proteinuria and mesangial cell proliferation) per transferred glomerular macrophage compared with unstimulated macrophages. This provides the first evidence that it is possible to directly modulate macrophage-mediated renal injury.
A number of IFN- inducible macrophage products have been implicated in causing proteinuria (reactive oxygen species, nitric oxide, tumor necrosis factor-, IL-1, and proteases) (6). We showed that IFN- upregulated iNOS mRNA and nitric oxide production in NR8383 macrophages; however, a role for nitric oxide in macrophage-mediated renal injury needs to be tested by specific blockade. Interestingly, IFN- did not increase the constitutive levels of IL-1 mRNA in NR8383 cells. This cytokine, which is known to play a pathogenic role in the macrophage-mediated phase of rat anti-GBM disease (13), may be important for the induction of proteinuria by unstimulated NR8383 cells, although this postulate requires demonstration by specific blockade of this molecule in macrophages. IFN- activation of macrophages also resulted in increased mesangial cell proliferation. This may be due to the increased PDGF-B gene expression seen in IFN- stimulated NR8383 macrophages, because this growth factor plays an important role in mesangial cell proliferation in glomerulonephritis (24–26 ). Other possible explanations for the increased mesangial proliferation include macrophage production of other mesangial cell growth factors, such as fibroblast growth factor-2 (25), or macrophage-induced PDGF production by intrinsic glomerular cells.
This study demonstrates that IFN- can significantly augment macrophage-mediated renal injury independently of any local or systemic effects of IFN-. Although the role of IFN- in macrophage activation is well described, the pathogenic role of this cytokine in experimental glomerulonephritis is somewhat controversial. There are conflicting reports as to whether anti-GBM disease is ameliorated or exacerbated in IFN- gene knockout mice (30,31 ). However, there is consensus that IFN- is required for the induction of autoantibody formation and nephritis in lupus-prone mice (32–34 ). Administration of IFN- in rat anti-Thy-1 mesangioproliferative disease increased glomerular macrophage accumulation and reduced mesangial cell proliferation (35), although the latter response was probably a direct effect of IFN- on mesangial cell proliferation (36).
Dexamethasone treatment for 18 h inhibited macrophage activation as shown by suppression of IFN-–induced nitric oxide production. Dex also abrogated renal injury caused by IFN-–activated macrophages. This was due to a decreased number of transferred macrophages accumulating within the glomerulus and was not due to dexamethasone-induced macrophage apoptosis. Thus, Dex–treated macrophages are unresponsive to the inflamed glomerulus.
Glucocorticoids are widely used as an immunosuppressive therapy in inflammatory kidney diseases. Glucocorticoids inhibit a variety of macrophage inflammatory responses. In particular, Dex has been shown to inhibit macrophage activation by IFN- in terms of nitric oxide production and upregulation of MHC class II expression (37,38 ), although not all studies concur that Dex can inhibit IFN-–induced upregulation of MHC class II expression (39,40 ). We confirmed the ability of Dex to inhibit IFN-–induced nitric oxide via suppression of iNOS gene transcription, although we did not observe any effect of Dex on IFN-–induced upregulation of RT1B gene expression. Indeed, a number of macrophage responses to IFN- are not inhibited by glucocorticoids. IFN- upregulation of Fc-R in human monocytes is enhanced by Dex (41,42 ), therapeutic concentrations of glucocorticoids have been shown to suppress the antimicrobial activity of human macrophages without impairing their responsiveness to IFN- (43,44 ), and IFN-–induced macrophage antibody-dependent cellular cytotoxicity is not inhibited by Dex (45). In our studies, Dex inhibited the ability of IFN-–activated macrophages to accumulate within the glomerulus and thus prevented renal injury. This finding may be related to Dex inhibition of IFN- upregulation of macrophage cell-surface adhesion molecules. These studies are the first in vivo demonstration that glucocorticoids act directly on macrophages to inhibit their recruitment to a site of inflammation. Previous studies have been limited to systemic glucocorticoid treatment in which there is significant inhibition of glomerular macrophage accumulation and renal injury (46,47 ); however, these studies do not distinguish between glucocorticoid effects on monocyte/macrophages and effects on other leukocytes and intrinsic renal cells.
An interesting aspect of this study was that long-term (18 h), but not short-term (1 to 3 h), stimulation of NR8383 macrophages was required to modulate macrophage-mediated renal injury. This suggests that a complex change in the pattern of gene transcription within the macrophage is required for these stimuli to modulate the macrophage response to the inflamed glomerulus. This finding is similar to the concept of macrophage "programming," in which exposure of macrophages to a specific stimulus determines how these cells will subsequently respond to a second stimulus (48,49 ). This programming concept is consistent with the inability of Dex to inhibit IFN-–induced nitric oxide production in vitro if the addition of Dex occurs more than 6 h after IFN- exposure.
In summary, this study has demonstrated that modulation of macrophage activation can profoundly affect macrophage-mediated renal injury in an adoptive transfer model. Importantly, we have shown that IFN- and Dex directly affect renal injury through their action on macrophages, independently of any effects on glomerular inflammation. Furthermore, the results support the notion that macrophages can be programmed to elicit a particular type of response when they subsequently encounter an inflammatory environment.
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