Progressive tubulointerstitial fibrosis is the common end point leading to end-stage renal disease in experimental and clinical settings. Since the peptide hormone leptin is involved not only in the regulation of obesity but also in the regulation of inflammation and fibrosis, we tested the hypothesis whether leptin deficiency has an impact on tubulointerstitial fibrosis in mice. Leptin-deficient (ob/ob) and leptin receptor-deficient mice (db/db) were exposed to 14 days of unilateral ureteral obstruction (UUO). The degree of fibrosis and inflammation was compared with that in sham-operated mice by performing immunohistochemistry, quantitative PCR, and Western blotting. We found that tubulointerstitial fibrosis was significantly reduced in the obstructed kidneys of ob/ob compared with db/db mice or control mice. Detailed analysis of infiltrating inflammatory cells by immunohistochemistry revealed a significant reduction of CD4+ cells at 14 days after UUO in both ob/ob and db/db mice. In contrast, we could not detect significant differences in CD8+ cells and macrophage content. Transforming growth factor (TGF)-β mRNA levels, TGF-β-induced Smad-2/3 activation, and the upregulation of downstream target genes were significantly reduced in ob/ob mice. In addition, we demonstrated that leptin could enhance TGF-β signaling in normal rat kidney fibroblasts in vitro. We conclude that leptin can serve as a cofactor of TGF-β activation and thus plays an important role in renal tubulointerstitial fibrosis. Therefore, selective blockade of the leptin axis might provide a therapeutic possibility to prevent or delay fibrotic kidney disease.
- tubulointerstitial fibrosis
- transforming growth factor-β
tubulointerstitial fibrosis characterizes the devastating final common pathway in most progressive renal diseases, regardless of their etiology (20, 24). Several cellular pathways, including fibroblast activation as well as tubular epithelial-mesenchymal transition (EMT), have been identified as the major avenues for the generation of the matrix-producing cells in diseased conditions. Among the many fibrogenic factors that regulate renal fibrosis, transforming growth factor-β (TGF-β) plays an essential role in a variety of kidney diseases, including diabetic nephropathy (19, 24, 37). After binding of TGF-β to serine/threonine kinase receptors of both type II and type I, the activated type I receptor transmits intracellular signals through the phosphorylation of the receptor-associated R-Smads (Smad2/3). The activated R-Smads form complexes with the common pathway C-Smad (Smad4). Only this active complex can translocate to the nucleus, associate with specific DNA-binding partners, and initiate transcriptional responses of specific target genes (37).
Interstitial fibrosis can be induced in vivo in rodents by unilateral ureteral obstruction (UUO). This disease model is characterized by early macrophage infiltration, interstitial fibrosis, and tubular injury mainly induced by activation of local TGF-β1 (2, 19). The increase of TGF-β facilitates matrix accumulation by simultaneous activation of renal fibroblasts and tubular epithelial cells, which both contribute to exaggerated matrix protein synthesis (2).
Leptin, a peptide hormone of the long-chain helical cytokine family, is predominantly produced by white adipose cells (46). Leptin plays a crucial role in body weight regulation by inhibiting food intake via hypothalamic effects and stimulating energy expenditure in skeletal muscle cells (12). Maffei et al. (27) were the first to report a strong positive correlation between leptin levels and the body mass index in humans and rodents. However, the ubiquitous distribution of the functional long form of the leptin receptor Ob-Rb in almost all tissues underlies the pleiotropism of leptin (13, 36). Several authors have suggested an essential fibrogenic role for leptin in experimental liver fibrosis (18, 35, 39). In addition, recent studies in renal fibroblasts and glomerular endothelial cells have indicated that leptin might induce cellular proliferation and matrix expression through synergistic activation of the TGF-β1 signaling system (21, 42). The significance of intact leptin signaling in renal fibrosis has not been studied so far.
Leptin-deficient (ob/ob) and leptin receptor-deficient (db/db) mice share the same phenotype. Both strains lack hypothalamic inhibition of appetite, leading to obesity, insulin resistance, and type II diabetes (9, 46). Both ob/ob and, most commonly, db/db mice have been intensively investigated as experimental models of diabetic nephropathy (38). Furthermore, ob/ob mice display impaired cell-mediated immunity and a propensity to develop Th2-dominant immune responses (25). Ob/ob mice are protected from inflammation and fibrosis in different disease models, including autoimmune encephalomyelitis, nephrotoxic nephritis, and liver fibrosis (18, 29, 41). Consistently, leptin replacement restores the susceptibility to experimental damage in rodents and humans (43).
The aim of our study was to investigate whether deletion of leptin signaling leads to reduced tubulointerstitial fibrosis. Since functional short-form leptin receptors (Ob-Ra) have been demonstrated in the kidney (42), we performed UUO in both ob/ob and db/db mice compared with appropriate controls.
MATERIALS AND METHODS
We studied ob/ob mice at 8 wk and db/db mice at 7 wk of age, before the onset of diabetic glomerulopathy (9, 46). Male wild-type (WT), ob/ob, db/db, and heterozygous db/m mice (n = 5 per group) were purchased from the Charles River Laboratories (www.criver.com). The animals received a standard diet with water ad libitum. Serum creatinine levels were identical in the mice used compared with the appropriate control strains. However, weight was already notably different between ob/ob and C57Bl/6 wild-type mice (21.1 ± 1.2 vs. 50.6 ± 4.3 g) as well as db/m and db/db mice (22.1 ± 0.8 vs. 38.8 ± 1.9 g). Blood glucose level measurements revealed the beginning of hyperglycemia in the db/db group at the end of the experiment (at 9 wk of age: average 23.0 ± 3.3 mM). UUO was performed as follows: after induction of general anesthesia by application of Avertin (2.5%), the abdominal cavity was exposed via a midline incision and the left ureter was ligated at two points with 4-0 silk and dissected in between. Successful ureteral obstruction was later confirmed by observation of dilation of the renal pelvis and proximal ureter. Sham-operated mice of each strain (n = 5 per group) were used as controls. After 14 days of UUO, mice were euthanized. After anesthesia with Avertin (2.5%), a laparotomy was performed and urine was collected by puncturing the bladder with a 23-gauge needle. The abdominal aorta was then cannulated with a 23-gauge needle, and the organs were perfused with ice-cold lactated Ringer solution. Both kidneys were removed, cut in thirds, and then fixed for 20 h in 3.75% paraformaldehyde in Soerensen's phosphate buffer and embedded in paraffin for histological examination, snap frozen in isopentane (−40°C) for cryostat sectioning, or frozen in liquid nitrogen and stored at −80°C for protein chemistry and TaqMan PCR analysis.
Blood glucose levels were measured with the Glucometer Elite (Bayer, Leverkusen, Germany). Creatinine levels were measured using an automated method (Beckman analyzer; Beckman, Munich, Germany). All procedures were carried out according to guidelines from the American Physiological Society and were approved by local authorities.
Primary antibodies used for Western blotting and immunhistochemical studies were rabbit anti-phospho-Smad2 (Cell Signaling Technology, Beverly, MA), anti-mouse T lymphocytes (CD4 and CD8; BD Pharmingen, Heidelberg, Germany), rat anti-mouse monocytes/macrophages (F4/80; Serotec, Oxford, UK), rabbit anti-mouse fibronectin (Poesel&Lorei, Hanau, Germany), mouse anti-α-smooth muscle actin (α-SMA; DAKO, Hamburg, Germany), and rabbit anti-mouse β-tubulin (Santa Cruz Biotechnology, Santa Cruz, CA). Horseradish peroxidase (HRP)-conjugated goat anti-rabbit as secondary antibodies for Western blotting were purchased from Santa Cruz Biotechnology.
Histological analysis was carried out on 2-μm paraffin sections cut on a rotation microtome. Following a standard deparaffinization and staining protocol, periodic acid-Schiff (PAS)-, Trichrome-, and Sirius red-stained renal cortex sections were assessed with a digital camera (Axiocam; Zeiss, Jena, Germany) connected to a light microscope (Axioplan-2; Zeiss) by two independent investigators blinded to the study. Analysis of inflammation was done by semiquantitative scoring of the infiltrating cells in 10 randomly chosen, nonoverlapping fields of cortex and outer medulla in PAS-stained sections (×200 magnification) as follows: 0 = none, 1 = weak, 2 = moderate, 3 = high, and 4 = very high numbers of infiltrating cells. Fibrosis was examined in Sirius red-stained sections by fluorescence microscopy, using the Cy3 channel, and scored as follows: 0 = normal, thin tubular basement membrane; 1 = moderately thickened basement membrane; 2 = severely thickened basement membrane; 3 = severely thickened basement membrane plus a few interpositioned collagen fibrils; and 4 = severely thickened basement membrane plus numerous interpositioned collagen fibrils. Data are expressed as the mean score of 10 randomly chosen, nonoverlapping fields of cortex and outer medulla per section (×400 magnification). Fibrosis was additionally assessed in Trichrome-stained sections and scored as follows: 0 = none, 1 = weak, 2 = moderate, 3 = severe, and 4 = very severe fibrosis per section (×200 magnification).
Immunohistochemistry was performed on cryosections and paraffin sections. Nonspecific binding sites were blocked with 10% normal donkey serum (Jackson ImmunoResearch Laboratory, West Grove, PA) for 30 min. Sections were then incubated with the primary antibody for 1 h. All incubations were performed in a humidified chamber at room temperature. For fluorescent visualization of bound primary antibodies, cryostat sections were further incubated with Cy3-conjugated secondary antibodies (Jackson ImmunoResearch Laboratory) for 1 h. On paraffin-embedded sections, a secondary goat anti-mouse antibody (Zytomed, Berlin, Germany) was visualized using diaminobenzidine (Zytomed). Sections were analyzed using a Zeiss Axioplan-2 imaging microscope with the computer program AxioVision 3.0 (Zeiss). Quantitative analysis of CD4-, CD8-, and F4/80-positive cells was done by counting the cell numbers in 10 randomly chosen, nonoverlapping fields (expressed as a mean) of cortex and outer medulla per section (×400 magnification). Semiquantitative analysis was scored as follows: 0 = none, 1 = weak, 2 = moderate, 3 = high, and 4 = very high expression. Semiquantitative analysis of α-SMA and fibronectin expression was done in 10 randomly chosen, nonoverlapping fields by scoring as follows: 0 = none, 1 = weak, 2 = moderate, 3 = high, and 4 = very high expression (×200 magnification).
For protein isolation, frozen kidney tissue from each animal was homogenized in RIPA buffer (50 mM Tris, pH 7.5, 150 mM NaCl, 0.5% sodium deoxycholate, 1% Nonidet P-40, and 0.1% SDS) containing protease inhibitor (Complete Mini; Roche, Mannheim, Germany), 1 mM sodium orthovanadate, 50 mM NaF, and 200 μg/l okadaic acid. The samples were centrifuged for 15 min (14,000 g). The supernatant was aliquoted on the basis of the protein concentration measured using a Bradford protein assay (Pierce, Rockford, IL) and stored at −80°C. Fifty micrograms of protein per lane were separated by SDS-PAGE and transferred to polyvinylidene difluoride membrane (Immobilon-P; Millipore, Bedford, MA). After probing with primary antibodies, antigen-antibody complexes were detected with HRP-labeled anti-rabbit and anti-mouse antibodies, respectively, and visualized using enhanced chemiluminescence reagents (Pierce) according to the manufacturer's protocol. Quantification was done by measuring relative density compared with β-tubulin (Quantify One; Bio-Rad).
RNA extraction and real-time quantitative PCR.
Total mRNA was extracted using Trizol reagent (Invitrogen). For quantitative real-time PCR (qPCR), 1 μg of DNase-treated total RNA was reverse transcribed using SuperScript II reverse transcriptase (Invitrogen), and qPCR was performed on an SDS 7700 system (Applied Biosystems, Darmstadt, Germany) using 10 ng of reverse-transcribed mRNA as a template, Rox dye as internal control (Invitrogen), FastStart Taq polymerase (Roche Diagnostics, Mannheim, Germany), and gene-specific primers in combination with SYBR green chemistry (Molecular Probes, Eugene, OR). PCR amplification was carried out for 10 min at 96°C and 40 cycles of 10 s at 95°C and 1 min at 60°C. β-Actin served as internal reference gene for normalization. Primer sequences are available on request. Quantification was carried out using QGene software (30).
Normal rat kidney (NRK) fibroblasts (catalog no. 6509; American Type Culture Collection, Rockville, MD; kindly provided by Prof. L. Schäfer, University of Frankfurt, Frankfurt/Main, Germany) were grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% heat-inactivated fetal calf serum, 100 U/ml penicillin, and 100 μg/ml streptomycin for 8–15 passages. To obtain quiescent NRK fibroblasts, cells were maintained in serum-starved DMEM supplemented with 0.5% heat-inactivated fetal calf serum for 24 h before the addition of recombinant mouse (rm)leptin (R&D Systems, Minneapolis, MN) or recombinant human (rh)TGF-β1 (Cell Sciences, Canton, MA).
Data are means ± SE. Unpaired Mann-Whitney U testing (2 sided) was used after the Kruskal-Wallis test had been applied to show significant differences between different UUO groups. P < 0.05 was considered statistically significant. Data analysis was performed using SPSS software (www.spss.com).
Leptin deficiency but not leptin receptor deficiency results in an attenuated profibrotic response in kidneys after UUO.
To study the effect of leptin deficiency on renal fibrosis, we performed UUO in WT mice, leptin-deficient ob/ob mice, leptin receptor-deficient db/db mice, and their littermates, db/m mice. Analysis was done 14 days after UUO. Examination of Sirius red- and Trichrome-stained renal cortex sections revealed dilated proximal tubules, interstitial expansion, ECM accumulation, and basement membrane thickening of both tubules and glomeruli in WT and ob/ob mice 14 days after UUO (Fig. 1). However, the degree of interstitial fibrosis in obstructed kidneys of WT, db/db, and db/m mice was much more pronounced compared with ob/ob mice, which exhibited significantly reduced basement membrane thickening, reduced interstitial expansion, and ECM deposition (Trichrome, P < 0.001; Sirius red, P < 0.05, respectively) (Fig. 1, k and l). No differences in tubular dilatation were evident among the groups. We also investigated the expression of α-SMA as a marker of activated myofibroblasts and that of fibronectin as a marker of ECM deposition by immunohistochemistry. As expected, expression of α-SMA was significantly increased in WT mice after UUO (Fig. 2 b). In contrast, expression of α-SMA was significantly attenuated in ob/ob mice (P < 0.005) (Fig. 2c). Analogous to the described histological findings, α-SMA expression after UUO was not attenuated in db/db mice (Fig. 2d). In addition, fibronectin deposition was solely reduced in ob/ob mice compared with WT, db/db, or db/m mice after 14 days of UUO (P < 0.005) (Fig. 2h).
In summary, histology and immunohistochemistry demonstrated markedly attenuated fibrosis, α-SMA, and fibronectin expression in ob/ob mice compared with db/db mice and respective controls. These data indicate that leptin deficiency but not deletion of the long-form leptin receptor Ob-Rb protects mice from matrix accumulation and tubulointerstitial fibrosis after UUO.
Effects of leptin signaling on inflammation and composition of inflammatory infiltrates in kidneys after UUO.
Since the UUO model directly links the inflammatory and profibrotic response, we examined whether leptin signaling has an influence on inflammatory cell infiltration in this model. Differences in leukocyte infiltration after UUO were semiquantitatively scored on PAS-stained sections. Obstructed kidneys of C57Bl/6, db/db, and db/m mice showed a homogenous inflammatory cell infiltration of the renal interstitium (Fig. 3). In contrast, we found significantly less inflammation in renal cortex sections of ob/ob mice after UUO (P < 0.005) (Fig. 3f).
To characterize the cellular composition of the infiltrates in this model, we performed immunohistochemistry with specific antibodies for monocytes/macrophages (F4/80) and lymphocyte subsets (CD4+ and CD8+). Interestingly, significant quantitative differences were detected among CD4+ lymphocytes (Fig. 3). CD4+ cells were twofold higher in controls compared with ob/ob and db/db mice after UUO (both P < 0.001). No significant differences were observed in CD8+ lymphocytes and monocytes/macrophages among groups (Fig. 3i). These data indicate that recruitment of CD4+ cells to sites of tissue inflammation requires leptin and the intact Ob-Rb.
TGF-β activation is reduced in leptin-deficient mice.
Since TGF-β is the central cytokine mediating UUO-induced tubulointerstitial fibrosis, we analyzed whether the advantageous response of leptin deficiency is based on a reduced activation of local TGF-β. When we measured TGF-β1 mRNA levels in whole kidney lysates after UUO, we detected reduced TGF-β1 expression levels solely in ob/ob mice (Fig. 4 a). Next, we performed Western blot analysis of whole kidney protein lysates. Obstructed kidneys of WT mice showed a strong and uniform phosphorylation response of Smad2/3. In contrast, Smad2/3 phosphorylation was significantly reduced in whole kidney lysates of ob/ob mice (Fig. 4b). The Smad2/3 phosphorylation response was similar in db/db mice compared with db/m mice (Fig. 4b), indicating a coactivating role for leptin in TGF-β activation in vivo. Analysis of mRNA expression levels of classic TGF-β downstream target genes in whole kidney tissue revealed significantly reduced expression levels of plasminogen activator inhibitor-1 (PAI-1), connective tissue growth factor (CTGF), matrix metalloproteinase-2 (MMP-2), and MMP-9 in ob/ob mice compared with all other groups after UUO (P < 0.005) (Fig. 5). Together, these data indicate a link between leptin and TGF-β in vivo.
Leptin augments TGF-β signaling in renal fibroblast in vitro.
Next, we examined the potential role of leptin as a cofactor for TGF-β activation in vitro. NRK fibroblasts were treated with a single dose of rmLeptin (100 ng/ml), rhTGF-β1 (5 ng/ml), or a combination of both. Stimulation with leptin alone resulted in a slight phosphorylation response of Smad2/3. Interestingly, combination treatment of leptin and TGF-β resulted in an enhanced Smad2/3 phosphorylation response compared with TGF-β alone (Fig. 6, a and b). These data indicate that leptin can serve as a cofactor of TGF-β activation in vitro.
Progressive fibrosis is the hallmark of end-organ failure in the kidney, since functional parenchymal cells are replaced by nonfunctional scar tissue. In the past, remarkable progress was made in slowing down the progression of chronic renal failure by such second-line therapeutic approaches as consequent blood pressure control. However, normalization of blood pressure alone is not efficient to prevent disease progression or reverse existing fibrosis. TGF-β is a powerful mediator of profibrotic responses in the kidney. TGF-β-induced tissue fibrosis is mediated by direct induction of matrix molecules, activation of resident fibroblasts, and EMT (2, 24). In the past, the blockade of TGF-β activation was proven to be a successful approach in preventing and even reversing matrix accumulation and fibrosis in animal models (5, 47). Especially in UUO and diabetic kidney disease, the role of TGF-β is very well defined as the key mediator of hypertrophic and prosclerotic changes (19, 33). However, TGF-β is a difficult target molecule for ablative therapy because of its pleiotropic effects on various tissues.
In addition to its central and peripheral metabolic effects on peripheral tissues, paracrine effects of leptin on TGF-β synthesis and synthesis of the TGF-β type II receptor have been described previously (15, 23, 42). Rat kidney interstitial fibroblasts respond to leptin treatments in vitro with increased mitogenesis and collagen expression (21). A similar effect was described in a model of hepatic fibrogenesis, where leptin induced collagen-I expression independently of TGF-β in human stellate cells (40). Likewise, leptin induced collagen-I expression in isolated mesangial cells of db/db mice. These direct effects of leptin are often potentiated by direct induction of the TGF-β type II receptor, leading to a more efficient TGF-β action as demonstrated in human stellate cells and mesangial cells (15, 39). Recently, Wolf et al. (42) demonstrated elevated TGF-β mRNA and subsequently increased matrix expression in glomerular endothelial cells after leptin treatment in vivo and in vitro.
In the present study we have demonstrated that leptin deficiency, but not leptin receptor deficiency, leads to significantly less inflammation, tissue damage, fibrotic changes, and matrix accumulation in ob/ob mice after UUO compared with db/db mice and respective controls. We also demonstrated markedly less TGF-β activation in leptin-deficient ob/ob compared with leptin receptor-deficient db/db mice after UUO. These results are well in line with our finding of markedly reduced mRNA levels of TGF-β, Smad2/3 phosphorylation (i.e., TGF-β activation) in ob/ob compared with db/db mice after UUO. Since db/db mice lack the functional long form of the leptin receptor (Ob-Rb), it is conceivable that either signaling via the short form of the receptor (Ob-Ra) or leptin per se as a cofactor of TGF-β activation might enhance interstitial fibrosis after UUO. We can demonstrate that leptin alone induces phosphorylation of Smad-2/3 in renal fibroblasts in vitro. In addition, leptin in combination with TGF-β1 leads to enhanced Smad phosphorylation.
We also have demonstrated that mRNA expression of typical downstream targets of TGF-β such as PAI-1 and CTGF accumulate in UUO but are significantly less induced in the obstructed ob/ob kidneys. PAI-1 promotes net proteolysis via inhibition of tissue-type plasminogen inhibitor (t-PA) and urokinase-type plasminogen activator (u-PA). Both t-PA and u-PA lead to activation of plasmin, which leads to fibrinolysis. Therefore, PAI-1 induction is a solid marker of ongoing tissue fibrosis. PAI-1 is upregulated in almost all known renal diseases, including UUO (10). PAI-1 knockout mice develop less tubulointerstitial fibrosis after UUO than their control littermates. Interestingly, this benefit results from less cellular infiltration as well as reduced mRNA levels for TGF-β and collagens but is not caused by an antiproteolytic effect via inhibition of t-PA or u-PA (32).
CTGF is mainly expressed in tubular cells in the kidney. It has been shown to induce EMT in tubuloepithelial cells, and its expression also can serve as a biomarker of ongoing kidney fibrosis (4, 7). Furthermore, CTGF antisense treatment has been shown to attenuate renal fibrosis after UUO (45). In addition, we detected less accumulation of MMP-2 and MMP-9 mRNA in ob/ob mice after UUO. This also could be explained by the lack of TGF-β activation, since MMPs are described target genes of TGF-β (1). The classic function of MMPs is the attenuation of fibrosis by degradation of matrix; however, recent studies have revealed some effects of MMP-2 and MMP-9 that might even lead to a more aggravated profibrotic response. Transgenic overexpression of MMP-2 in tubular cells leads to direct induction of renal injury and interstitial collagen (8). In addition, the activation of MMP-9 promotes degradation of the tubular basement membrane, which is a critical key step leading to kidney fibrosis via the induction of EMT (43, 44).
Ob/ob and db/db mice are widely accepted as classic disease models of type 2 diabetes. Both animal models display glomerular features similar to human diabetic glomerulopathy. However, tubulointerstital fibrosis, the key event leading to end-stage renal failure in human diabetic nephropathy, is rarely reported in db/db mice but virtually absent in ob/ob mice (3, 38). Based on our results, we hypothesize that the lack of leptin-mediated signaling responses is due to a missing cofactor necessary for local TGF-β activation that leads to fibrosis of the tubulointerstitial compartment in the latter model. Moreover, Gunduz et al. (14) reported an increase in glomerular TGF-β activation after leptin infusion, indicating a close regulatory connection of leptin and the TGF-β axis. However, the TGF-β activation in glomerular cells seems to occur independently of leptin, since local glomerular synthesis and activation of TGF-β and CTGF has been demonstrated in both model systems in the diabetic state (5, 34).
Despite the fact that we detected fewer inflammatory cells in the obstructed kidneys of ob/ob mice, we detected no difference in macrophage infiltration. Macrophages play a central role after tissue injury, since they produce numerous factors that promote activation of resident fibroblasts and synthesis of matrix. Macrophages of ob/ob mice display some phenotypic alterations (i.e., reduced phagocytosis capacity and increased IL-6 levels), but these have so far not directly been linked to fibrogenesis (22, 28). However, recently published evidence suggests that in the absence of active TGF-β, macrophages may be ineffective in promoting fibrosis (26, 31).
In our model we found decreased numbers of CD4+ cells in both ob/ob and db/db mice. This is in line with previous results showing that patients suffering from leptin deficiency had reduced numbers of circulating CD4+ cells and impaired T cell proliferation (11). However, since db/db mice are not protected from renal fibrosis, reduced numbers of CD4+ cells in ob/ob and db/db mice are probably not primary effectors in UUO. In general, the impact of TGF-β on immune cells and the immune-modulatory response is determined by the type of inflammatory microenvironment and the cell type. This is especially exemplified in the case of CD4+ cells. Naive cells and Th1 cells respond to TGF-β1 with a suppressed proliferative response, whereas TGF-β1 has little or no effect on the proliferation of Th2 cells (17). Moreover, TGF-β has direct immunomodulatory functions by influencing the conversion of naive infiltrating CD4+ cells to regulatory CD4+CD25+ cells (6). Thus the difference in CD4+ cell number in the leptin-deficient mice could be explained in part by a lack of local TGF-β activation. However, we can certainly speculate that tissue infiltration of CD4+ cells partially requires an intact leptin-leptin receptor axis. Our data are further supported by observations in experimental models of hepatic fibrosis demonstrating a reduced inflammatory and profibrotic response in the absence of leptin (16, 35).
In summary, we have provided evidence that leptin per se or leptin signaling via the short-form receptor (Ob-Ra) functions as an important coactivator of local TGF-β in the kidney. Targeting leptin signaling in inflammatory or profibrotic disease states of the kidney could be a novel approach to prevent tubulointerstital fibrosis.
This work was supported by an Emmy Noether Scholarship granted by Deutsche Forschungsgemeinschaft (Schi 587/2) to M. Schiffer.
We thank Melanie Paschy, Yvonne Nicolai, Herle Chlebusch, and Kerstin Bankes for excellent technical assistance.
↵* P. Kümpers and F. Gueler contributed equally to this work.
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