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Division of Cellular and Molecular Pathology, Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania.
Correspondence to Dr. Youhua Liu, Department of Pathology, University of Pittsburgh School of Medicine, S-405 Biomedical Science Tower, 200 Lothrop Street, Pittsburgh, PA 15261. Phone: 412-648-8253; Fax: 412-648-1916; E-mail: liuy{at}msx.upmc.edu
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Abstract |
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-smooth muscle actin–positive myofibroblast cells is a key event in the progression of chronic renal diseases that leads to end-stage renal failure. Although the origin of these myofibroblasts in the kidney remains uncertain, emerging evidence suggests that renal myofibroblasts may derive from tubular epithelial cells by a process of epithelial to mesenchymal transition. It was demonstrated that hepatocyte growth factor (HGF) exhibited a remarkable ability to block this phenotypic transition both in vitro and in vivo. HGF abrogated the -smooth muscle actin expression and E-cadherin depression triggered by transforming growth factor-ß1 in tubular epithelial cells in a dose-dependent manner. HGF also blocked morphologic transformation of tubular epithelial cells and inhibited the expression and extracellular deposition of fibronectin. In a mouse model of renal fibrosis disease induced by unilateral ureteral obstruction, transforming growth factor-ß type I receptor expression was specifically increased in renal tubules, and myofibroblastically phenotypic transition of the tubules was evident in vivo. Remarkably, injections of exogenous HGF blocked myofibroblast activation and drastically prevented renal interstitial fibrosis in the obstructed kidneys. These results suggest that tubular epithelial to myofibroblast conversion may play an important role in the pathogenesis of renal fibrosis and that blocking this phenotypic transition could provide a novel therapeutic strategy for the treatment of fibrotic diseases. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-smooth muscle actin (SMA)-positive myofibroblast cells is believed to be a central event that plays a key role in the progression of chronic renal fibrosis. Indeed, myofibroblast activation has been identified as a predictor of renal disease progression both in experimental animal models and in patients (4,5). Clearly, a possible key to an effective therapy for progressive renal fibrosis is to find a strategy that prevents the activation and accumulation of renal myofibroblasts in the diseased kidneys.
Although the fundamental role of myofibroblasts in progressive renal fibrosis is well established, the origin of these SMA-positive cells in the kidney under pathologic conditions remains poorly defined. They are often presumed to derive from local activation of renal interstitial fibroblasts and/or perivascular smooth muscle cells (6). Previous studies from our laboratory showed that blockage of hepatocyte growth factor (HGF) signaling in vivo by a neutralizing antibody markedly induces de novo expression of SMA in renal tubular epithelium in the remnant kidney model of chronic renal disease (7). Some SMA-positive renal tubular epithelial cells are found to become elongated and sometimes separated from neighboring cells. This finding is consistent with the hypothesis that, under pathologic conditions, tubular epithelial cells may transdifferentiate into myofibroblasts, as marked by the expression of SMA (8,9). Furthermore, these observations suggest that endogenous HGF signaling is essential for preserving and maintaining the tubular epithelial cell phenotype by blocking this epithelial to myofibroblastic conversion in vivo.
Here we provide in vitro and in vivo evidence to support that renal myofibroblasts may derive from tubular epithelial cells by an epithelial to myofibroblast transition process under chronically pathologic conditions and that HGF exhibits a remarkable ability to block this epithelial to myofibroblast transition both in vitro and in vivo.
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Materials and Methods |
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Animals and HGF Treatment
Male CD-1 mice that weighed approximately 18 g to 20 g were obtained from Harlan Sprague-Dawley (Indianapolis, IN). Eighteen mice were randomly assigned to three groups with 6 mice each. Unilateral ureteral obstruction (UUO) was performed using an established procedure (12,13). Briefly, under general anesthesia, complete ureteral obstruction was performed by double-ligating the left ureter using 4-0 silk after a midline abdominal incision. Sham-operated mice had their ureters exposed and manipulated but not ligated. Starting on the day of surgery, mice were administrated recombinant human HGF through tail-vein injections at a dose of 200 µmg/kg body wt every 12 h for 6 d. Control mice received an injection of the same volume of vehicle (0.9% saline solution). Mice were killed 7 d after surgery, and the kidneys were removed. One part of the kidneys was fixed in 10% phosphate-buffered formalin for histologic and immunohistochemical studies after paraffin embedding. Another part was immediately frozen in OCT compound for cryosection. The remaining kidneys were snap-frozen in liquid nitrogen and stored at -80°C for protein extractions.
Western Blot Analysis
HKC cells and cytokine-treated cells were lysed with sodium dodecyl sulfate (SDS) sample buffer (62.5 mM Tris-HCl [pH 6.8], 2% SDS, 10% glycerol, 50 mM DTT, and 0.1% bromophenol blue). Kidney tissues were homogenized by a polytron homogenizer (Brinkmann Instruments, Westbury, NY) in RIPA lysis buffer (1% NP40, 0.1% SDS, 100 µg/ml phenylmethylsulfonyl fluoride, 0.5% sodium deoxycholate, 1 mM sodium orthovanadate, 2 µg/ml aprotinin, 2 µg/ml antipain, and 2 µg/ml leupeptin in PBS) on ice. The supernatants were collected after centrifugation at 13,000 x g at 4°C for 20 min. Protein concentration was determined using a BCA protein assay kit (Sigma), and whole-tissue lysates were mixed with an equal amount 2x SDS loading buffer (125 mM Tris-HCl, 4% SDS, 20% glycerol, 100 mM DTT, and 0.2% bromophenol blue), as described previously (14). Samples were heated at 100°C for approximately 5 to 10 min before loading and were separated on precasted 10% or 5% SDS-polyacrylamide gels (Bio-Rad, Hercules, CA). The proteins were electrotransferred to a nitrocellulose membrane (Amersham, Arlington Heights, IL) in transfer buffer containing 48 mM Tris-HCl, 39 mM glycine, 0.037% SDS, and 20% methanol at 4°C for 1 h. Nonspecific binding to the membrane was blocked for 1 h at room temperature with 5% nonfat milk in TBS buffer (20 mM Tris-HCl, 150 mM NaCl, and 0.1% Tween 20). The membranes were then incubated for 16 h at 4°C with various primary antibodies in blocking buffer containing 5% milk at the dilutions specified by the manufacturers. The mouse monoclonal anti-SMA antibody (clone 1A4) was purchased from Sigma. The monoclonal antibodies for E-cadherin (clone 36) and fibronectin (clone 10) were obtained from Transduction Laboratories (Lexington, KY). The goat polyclonal anti–type I collagen antibody was obtained from Southern Biotechnology Associates (Birmingham, AL). The anti–TGF-ß type I receptor (sc-398) and anti-actin (sc-1616) antibodies were purchased from Santa Cruz Biochemicals (Santa Cruz, CA). After extensive washing in TBS buffer, the membranes were then incubated with horseradish peroxidase–conjugated secondary antibody (Bio-Rad) for 1 h at room temperature in 1% nonfat milk dissolved in TBS. Membranes were then washed with TBS buffer, and the signals were visualized using the enhanced chemiluminescence system (ECL, Amersham).
Immunofluorescence Staining
Indirect immunofluorescence staining was performed using an established procedure (15). Briefly, control or cytokine-treated HKC cells cultured on coverslips were washed with cold PBS twice and fixed with cold methanol:acetone (1:1) for 10 min on ice. After extensive washing with PBS containing 0.5% bovine serum albumin three times, the cells were blocked with 20% normal donkey serum in PBS buffer for 30 min at room temperature and then incubated with the specific primary antibodies described above, except for the mouse monoclonal antivimentin (clone V9) and rat monoclonal anti–E-cadherin (clone DECMA-1), which were obtained from Sigma. To visualize the primary antibodies, cells were stained with FITC-conjugated secondary antibodies (Sigma). After washing, cells were double-stained with 4',6-diamidino-2-phenylindole, HCl to visualize the nuclei. For visualizing F-actin, cells were stained with tetramethylrhodamine isothiocyanate (TRITC)-conjugated phalloidin. Stained cells were mounted with antifate mounting medium (Vector Laboratories, Burlingame, CA) and viewed with a Nikon Eclipse E600 Epi-fluorescence microscope (Melville, NY).
For co-localization of SMA and the proximal tubular marker in kidney sections using immunofluorescence staining, frozen sections at 5 µm thickness were cut using a cryostat, mounted onto poly L-lysine–coated slides, and fixed in 4% paraformaldehyde in PBS for 30 min. The slides were stained for SMA using the Vector M.O.M. immunodetection kit by the protocol specified by the manufacturer (Vector Laboratories). The slides were then stained with fluorescein-conjugated lectin from Tetragonolobus purpureas (Sigma). Stained slides were viewed and photographed as described above.
Morphological and Immunohistochemical Studies
Kidney sections from paraffin-embedded tissues were prepared at 4-µm thickness using a routine procedure (7). Sections were stained with hematoxylin/eosin for routine histology. Another set of sections was stained with Masson-Trichrome method for identifying interstitial collagen by blue color (7,13). Immunohistochemical localization was performed using the Vector M.O.M. immunodetection kit. The primary antibodies used were antifibronectin (Transduction Laboratories), antivimentin and anti–E-cadherin (Sigma), polyclonal anti–TGF-ß1 (sc-146), anti–TGF-ß type I receptor, and mouse monoclonal anti–proliferating cell nuclear antigen (sc-56; Santa Cruz Biochemical). As a negative control, the primary antibody was replaced with either nonimmune mouse or rabbit IgG, corresponding to species of the primary antibodies. Tubular E-cadherin and vimentin expression was semiquantitatively determined by counting positive tubules in at least 10 randomly chosen nonoverlapping high-power (x400) fields for each mouse. Injury score, collagen deposition, and fibronectin expression were scored on a scale from 0 to 3, as previously reported (7,16): 0, absent; 1, mild; 2, moderate; and 3, severe. The overall mean scores were calculated based on individual values, which were determined on at least 10 fields per mouse, six mice per group.
Determination of Tissue TGF-ß1 Levels by Enzyme-Linked Immunosorbent Assay
For measurement of tissue TGF-ß1 level, kidneys were homogenized in the extraction buffer containing 20 mM Tris-HCl (pH 7.5), 2 M NaCl, 0.1% Tween-80, 1 mM ethylenediaminetetraacetate, and 1 mM phenylmethylsulfonyl fluoride, and the supernatant was recovered after centrifugation at 19,000 x g for 20 min at 4°C. Kidney tissue TGF-ß1 level was determined by using a commercial Quantikine TGF-ß1 enzyme-linked immunosorbent assay kit in accordance with the protocol specified by the manufacturer (R & D Systems). The concentration of active TGF-ß1 and total TGF-ß1 (acid-activated) in kidneys was expressed as picograms per milligram of total protein.
Statistical Analyses
Animals were randomly assigned to control and treatment groups. All data examined were expressed as mean ± SEM. For Western blot analysis, quantitation was performed by scanning and determination of the intensity of the hybridization signals. Statistical analysis of the data were performed by Student-Newman-Kuels test using SigmaStat software (Jandel Scientific, San Rafael, CA). Values of P < 0.05 were considered to be statistically significant.
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Results |
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SMA In VitroSMA, the phenotypic marker of myofibroblasts. Among the agents tested, TGF-ß1, a well-characterized profibrogenic cytokine, markedly induced de novo SMA expression in HKC cells (Figure 1a). Dose-response studies revealed that TGF-ß1 induced SMA expression at a concentration as low as 0.1 ng/ml. Remarkably, HGF dramatically abrogated TGF-ß1–induced SMA expression in HKC cells in a dose-dependent manner (Figure 1, b and c). At a concentration of 20 ng/ml, HGF completely blocked SMA expression induced by 2 ng/ml TGF-ß1. As the molecular weight of HGF (95 kD) is much larger than that of TGF-ß1 (12.5 kD), the molar concentrations of HGF (approximately 0.21 nM) and TGF-ß1 (approximately 0.16 nM) were comparable. Other cytokines, such as epidermal growth factor and insulin-like growth factor-I, did not significantly suppress TGF-ß1–induced SMA expression at the same molar concentration as HGF (0.2 nM) (Figure 1b). These results suggest that HGF is the most potent, if not unique, cytokine capable of blocking TGF-ß1–induced SMA expression in tubular epithelial cells in vitro. Using an indirect immunofluorescence staining, we also demonstrated that TGF-ß1 induced SMA expression and assembly in HKC cells, as shown by the presence of cytoplasmic SMA-positive microfilaments (Figure 1, d through g). HGF markedly blocked this induction of SMA in tubular epithelial cells. Of note, TGF-ß1 was unable to induce SMA expression and phenotypic conversion in mouse inner medullary collecting duct (mIMCD-3) cells that are developmentally derived from ureteric bud epithelium (data not shown).
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SMA in HKC cells but also prohibits the loss of the epithelial marker E-cadherin. The loss of E-cadherin expression in tubular epithelial cells after TGF-ß1 treatment was also independently confirmed by indirect immunofluorescence staining (Figure 2, b through e). In agreement with the Western blot data, treatment of TGF-ß1 resulted in total loss of E-cadherin staining in the plasma membrane of HKC cells and co-incubation with TGF-ß1 and HGF largely restored the E-cadherin protein staining.
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SMA Expression and Prevents Myofibroblast Activation In VivoSMA abundance (Figure 5a). Strikingly, intravenous injections of exogenous HGF (200 µg/kg body wt every 12 h for 6 d) largely blocked renal SMA expression (Figure 5a). Quantitative studies revealed that the level of SMA protein in the obstructed kidneys after HGF administration was markedly inhibited compared with that given vehicle (Figure 5b), suggesting that HGF prevents renal myofibroblast activation in vivo.
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SMA-positive cells in the diseased kidney, we used a double immunofluorescence staining for both SMA and a proximal tubule marker, FITC-labeled lectin from Tetragonolobus purpureas (25,26). The SMA staining typically was limited to small arteries and arterioles in the kidneys of sham-operated mice. However, a large increase in SMA staining was observed in the widened interstitium and proximal tubular epithelium of the obstructed kidneys (Figure 5). Co-localization (yellow) of SMA (red) and the proximal tubular cell marker (green) indicates that the tubular epithelial cells acquired a phenotypic marker of myofibroblasts in the obstructed kidney (Figure 5), suggesting that there are cells at the transitional stage between the epithelial and myofibroblast phenotypes. Of note, the SMA-positive proximal tubular epithelial cells typically were disorganized, irregular in shape, and estranged from neighboring cells (Figure 5). After injection of exogenous HGF, renal SMA staining in the obstructed kidneys declined toward the baseline level (Figure 5), suggesting that, similar to the in vitro situation, HGF drastically blocks this phenotypic conversion in vivo.
Both TGF-ß1 and TGF-ß Type I Receptor Are Specifically Upregulated in Tubular Epithelium In Vivo
Because TGF-ß1 initiates tubular epithelial to myofibroblast conversion in vitro, we next examined the expression of TGF-ß1 and its type I receptor in the diseased kidneys. The obstructed kidneys displayed a marked increase in TGF-ß1 expression at 7 d after surgery (Figure 6). Quantitative determination by a specific enzyme-linked immunosorbent assay exhibited approximately 15-fold and 22-fold increases in active and total TGF-ß1 protein, respectively, in the obstructed kidneys at 7 d after UUO (Figure 6b). Importantly, immunohistochemical staining revealed that TGF-ß1 expression was largely upregulated in renal tubular epithelium in the diseased kidneys (Figure 6g). Injection of exogenous HGF markedly inhibited TGF-ß1 expression in the obstructed kidney (Figure 6).
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HGF Preserves Tubular Epithelial Phenotypes and Ameliorates Renal Fibrosis In Vivo
Because HGF blocks epithelial to myofibroblast conversion in vitro and myofibroblast activation in vivo, we reasoned that blockage of this phenotypic transition by HGF might provide a novel strategy for preventing renal fibrosis. To this end, we investigated the consequence of HGF injections in vivo by examining the expression of various cell-type–specific markers and renal fibrosis in the obstructed kidneys after UUO. As shown in Figure 7, epithelial cell marker E-cadherin was markedly decreased in the tubular epithelium from the obstructed kidneys of UUO mice, whereas de novo expression of the mesenchymal marker vimentin was evident in the renal tubules. These data essentially recapitulate our in vitro observations and suggest that cell phenotypic transdifferentiation may be of importance in the pathogenesis of renal fibrosis. Administration of exogenous HGF largely prevented this phenotypic conversion. Of interest, despite tubular atrophy as a characteristic feature of this model, increased cell proliferation, as demonstrated by proliferating cell nuclear antigen staining, was evident in the obstructed kidneys.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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SMA, display myofibroblastic morphology, lose epithelial E-cadherin, and produce interstitial matrix components such as fibronectin. Profibrogenic cytokine TGF-ß1 is a strong initiator of this conversion, and HGF dramatically blocks this process both in vitro and in vivo. These observations suggest that tubular epithelial cells actively play a role in the pathogenesis of chronic renal fibrosis, rather than being innocent bystanders, as previously thought. Our data also establish that tubular epithelial to myofibroblast transition perhaps is a new, novel target for therapeutic strategies designed to treat chronic renal fibrotic diseases.
The identification of TGF-ß1 as a key player in renal epithelial to myofibroblast conversion allows us to reexamine its role and mechanisms in progressive renal fibrosis. It is widely accepted that TGF-ß1 is a key modulator of organ fibrosis after tissue injury (31,32). However, previous studies emphasized the role of TGF-ß1 in stimulating myofibroblasts to produce fibrogenic proteins. Little is known about the origin of renal myofibroblasts or the effects of TGF-ß1 on tubular epithelial cells in renal fibrogenesis. Paradoxically, it is the tubular epithelium where the TGF-ß type I receptor, which determines which cells respond to TGF-ß1, is specifically and almost exclusively upregulated in diseased kidneys (Figure 6) (33), suggesting that tubular epithelial cells are the in vivo targets of TGF-ß1 under pathologic conditions. Indeed, significant changes in cell phenotypes, such as loss of E-cadherin and de novo expression of SMA and vimentin, take place in renal tubules in the obstructed kidneys. Cells at the transitional stage with both SMA and tubular marker are abundant in vivo (Figure 5), which is consistent with a previous observation demonstrating the presence of cells still positive for epithelial markers in the widened interstitium of end-stage diseased kidney in patients (25). It is conceivable to speculate that once cells progress through this transitional stage, they will lose tubular marker and retain the SMA and myofibroblastic phenotypes. In view of the fact that TGF-ß1 initiates renal epithelial to myofibroblast transition in vitro and that tubule-specific elevation of TGF-ß1 and its receptor protein occurs in the diseased kidney, our results likely provide novel insights into the mechanisms of this potent profibrogenic cytokine in promoting chronic renal fibrosis.
Given that, except for the collecting duct cells, all renal epithelial cells are developmentally derived from the metanephrogenic mesenchyme via mesenchymal to epithelial conversion (19,34), it is not surprising to find that tubular epithelial cells possess the ability to transdifferentiate back into a mesenchymal phenotype under certain pathologic conditions. This process, in essence, is a reverse embryogenesis. In agreement with this view, we found that the renal collecting duct cell line mIMCD-3, which is developmentally derived from ureteric bud epithelium, cannot undergo phenotypic conversion after incubation with TGF-ß1, suggesting that renal epithelial to myofibroblast transition likely is limited to the tubular epithelia that developmentally derived from metanephrogenic mesenchyme. HGF is obviously a key regulator of renal cell transdifferentiation both during normal nephrogenesis and under pathologic circumstances. Previous studies showed that HGF and its specific receptor c-met are coexpressed in early metanephrogenic mesenchyme, leading to formation of an autocrine loop essential for promoting mesenchymal to epithelial transition during early nephrogenesis (35,36). Furthermore, coexpression of both HGF and c-met in vitro by transfection of fibroblasts induces a phenotypic conversion into epithelium (37), and HGF has been shown to be responsible for facilitating the conversion of metanephrogenic mesenchymal cells into epithelium (38). In light of the role of HGF in initiating/promoting mesenchymal to epithelial conversion during nephrogenesis, it is reasonable to speculate that HGF is also essential for preserving the tubular epithelial cell phenotype by blocking an epithelial to myofibroblast transition under pathologic conditions.
It should be noted that given the pleiotropic nature of its actions, HGF inhibition of renal fibrogenesis in the obstructed kidneys could be mediated by other mechanisms as well. Studies from our laboratory and others demonstrate that HGF protects tubular epithelial cells from apoptosis and promotes matrix degradation by upregulating proteinases (7,39–41). Therefore, inhibition of apoptosis and activation of matrix degradation by HGF could be two additional pathways that lead to the suppression of renal fibrogenesis in vivo. Of note, the presence of these potential multiple mechanisms by no means suggests that they are mutually exclusive. In fact, it is possible that the multiple pathways triggered by exogenous HGF may work in concert to lead to ultimate amelioration of renal fibrosis in the obstructed kidneys in vivo.
The observation that HGF blocks myofibroblast activation and fibrosis in animals after chronic renal injury may have significant implications for developing clinically relevant therapeutic strategies for renal diseases. It is tempting to propose that administration of HGF may provide a novel and effective treatment for chronic renal diseases by specifically inhibiting the activation of myofibroblasts, the principal cells responsible for the accumulation and deposition of extracellular matrix seen in the diseased kidney. Although the antifibrogenic effect of HGF in vivo remains to be verified in other animal models with different causes, delivery of exogenous HGF protein or its gene (42,43) seems to have the potential to block the progression of chronic renal fibrosis, one of the devastating diseases that is otherwise incurable.
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Acknowledgments |
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P < 0.01, UUO + vehicle versus UUO + HGF. (c through k) Double immunofluorescence staining shows co-localization of 
) and total (
) TGF-ß1 protein in the obstructed kidneys after UUO for 7 d. Administration of HGF inhibited renal TGF-ß1 expression. Data are presented as mean ± SEM from six animals per group (n = 6). *P < 0.01, sham versus UUO. 
). Data are presented as mean ± SEM from six animals per group (n = 6). **P < 0.01, sham versus UUO. 