Homologous peptide of connective tissue growth factor ameliorates epithelial to mesenchymal transition of tubular epithelial cells

www.elsevier.com/locate/issn/10434666 Cytokine 36 (2006) 35–44

Homologous peptide of connective tissue growth factor ameliorates epithelial to mesenchymal transition of tubular epithelial cells Yujun Shi, Zhidan Tu, Wei Wang, Qing Li, Feng Ye, Jinjing Wang, Jing Qiu, Li Zhang, Hong Bu *, Youping Li Key Laboratory of Transplant Engineering and Immunology, Ministry of Health, West China Hospital, Sichuan University, Chengdu 610041, PR China Received 13 March 2006; received in revised form 18 October 2006; accepted 26 October 2006

Abstract The hallmark of failing renal transplants is tubular atrophy and interstitial fibrosis. The cytokine connective tissue growth factor (CTGF or CCN2) plays an important role in epithelial–mesenchymal transition (EMT) of tubular epithelial cells (TECs). A unique domain within CTGF (IRTPKISKPIKFELSG) which binds to its potential receptor integrin avb3 has been identified. This study was carried out to further characterize a synthetic hexadeca-peptide (P2) homologous to this domain and to determine its effect on CTGF-mediated solid phase cell adhesion, EMT induction and fibrogenesis in rat renal NRK-52E cells. Results showed that both P2 and recombinant CTGF bound to NRK-52E cells. Unlike CTGF, P2 had little effect on EMT induction including cytoskeleton remodeling and expression of a-smooth muscle actin (a-SMA) and E-cadherin, nor did it have effect on fibrogenic induction including alternation of extracellular matrix (ECM) proteins, collagen type I and IV at gene and protein levels. All data showed that P2 bound preferably on the surface of NRK-52E cells and inhibited the effect of CTGF on EMT induction and cell fibrogenesis, probably by occupying the binding sites of CTGF within its potential receptors. Therefore, P2 may be used as a potential anti-fibrotic agent.  2006 Elsevier Ltd. All rights reserved. Keywords: Connective tissue growth factor; Fibrosis; Peptide; Epithelial–mesenchymal transition; Tubular epithelial cells

1. Introduction Renal interstitial fibrosis is a common pathological change in most end-stage renal diseases [1–4], including chronic renal allograft dysfunction which remains as a leading cause of renal allograft failure. The typical morphologic changes in the end-stage renal allograft include tubular atrophy, interstitial fibrosis, glomerulopathy and occlusive changes [5–8]. Many factors contribute to chronic allograft dysfunction, making it difficult to identify the pathogenesis of the kidney failure [9–16]. Prevention of the interstitial fibrotic lesion may offer a promising strategy to improve the end-stage function of the renal allograft [6,17].

*

Corresponding author. E-mail address: [email protected] (H. Bu).

1043-4666/$ - see front matter  2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.cyto.2006.10.009

The downstream signal mediator of transforming growth factor beta (TGF-b), known as connective tissue growth factor (CTGF, also known as CCN2), is a matricellular protein which plays an important role in pathological fibrosis. Recent data have revealed that elevated CTGF is detectable in fibrotic lesion, and the durative overexpression of CTGF is essential to scarring formation [18–22]. Both in vivo and in vitro studies confirmed that CTGF expresses in renal tubular epithelial cells (TECs), which strongly suggest that CTGF is one of the key cytokines in renal fibrosis and plays a crucial role in epithelial–mesenchymal transition (EMT) of TECs [23–25]. The interaction between CTGF and its target cells and the specific CTGF receptors in TECs are still unknown. Recently, CTGF has emerged as a novel ligand for different integrin subtypes to elicit specific biological effects [26,27]. Gao, et al reported that a motif containing 16 residues in C-terminal portion of CTGF is a unique and essential

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binding domain for its potential receptors, integrin avb3, in hepatic stellate cells (HSC) [27]. Since both the integrin av and b3 subunits are also detected in TECs [28,29], it is possible that blockage of the binding between CTGF and integrin avb3 may interrupt the biological effects of CTGF, specifically on cell transdifferentation. In this study, we showed that P2, the hexadeca-peptide homologous to the binding motif of CTGF, can competitively bound to the renal TEC NRK-52E cells without activating the downstream molecules involved in the cell transition to myofibroblasts (MFB). This peptide, therefore, may be used as a competitive inhibitor of CTGF and hence provides a promising strategy to treat renal interstitial fibrosis. 2. Materials and methods 2.1. Antibodies and reagents Hamster monoclonal anti-integrin av subunit, rabbit anti-mouse E-cadherin, goat polyclonal anti-collagen type I, rabbit polyclonal anti-collagen type IV were purchased from Santa Cruz Biotechnology (USA). Hamster monoclonal anti-integrin b3 was from BD Biosciences. Mouse antismooth muscle actin (a-SMA) monoclonal antibody was obtained from DakoCytomation (Denmark). Mouse monoclonal anti-b-actin was from Abcam Ltd. (UK). All fluorescein isothiocyanate (FITC)-labeled secondary antibodies were products of DakoCytomation. Horseradish peroxidase (HRP)-labeled secondary antibodies were obtained from Chemicon International, Inc (USA). SV Total RNA Isolation kit was from Promega Corporation (USA). One step RNA PCR kit for reverse transcriptionPCR was from Takara Biotechnology Co., Ltd. (Dalian, China). The CyQUANT cell proliferation assay kit was from Molecular Probes (USA). M-PER Mammalian Protein Extraction Reagent and SuperSignal West Pico Chemiluminescent Substrate for Detection of HRP were from Pierce Biotechnology (USA).

2.2. CTGF and peptide Recombinant human CTGF was a product of BioVision, Ltd. (USA). CTGF proteins used in this study was of a lower molecular weight isoform containing the 98 amino acid residues of the C-terminal portion of the full-length CTGF protein, which exerted heparin binding, cell adhesion, and mitogenic activity. The rhCTGF was highly conserved between rat and human. A part of the rhCTGF was labeled with an EZ-Label Rhodamine Protein Labeling Kit (Pierce Biotechology). In brief, 20 lg lyophilized CTGF was dissolve in 200 ll BupH borate buffer, and then the mixture was transferred to a reaction tube. One microliters of rhodamine dye was added in the reaction tube and mixed well and incubated at room temperature for 1 h. The sample was added to the Slide-A-Lyzer Mini Dialysis Unit to float in a container with 100 ml PBS for 1 h to remove the excess fluorescent dye. The peptide IRTPKISKPIKFELSG, according to Gao et al. [27] named P2 was synthesized using a CS036 peptide synthesizer. The final product was dissolved in 50% aqueous acetonitrile and lyophilized to yield a white solid. The crude peptide was examined by reverse phase high performance liquid chromatography (HPLC) for purity, and the correct molecular weight was confirmed by using a Kratos Axima CFR plus V2.3.2a electrospray mass spectrometer [30] as shown in Fig. 1. The major peak was purified with >95% purity. The synthetic P2 was diluted in dimethyl sulfoxide (DMSO) and stored at 70 C prior to further use. At the same time, the antisense peptide of P2, which sequence was GSLEFKIPKSIKPTRI named P2c was synthesized as a control peptide of P2. 2.3. Cell culture Rat renal TEC NRK-52E cells were originally obtained from American Type Culture Collection (ATCC). The cells

Fig. 1. Electrospray mass chromatogram and reverse phase high performance liquid chromatogram of synthesized P2. The mass chromatogram showed that the molecular weight of P2 is 1814.47 and the HPLC showed that the purity is 95%.

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were cultured in Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal bovine serum in saturated humidity and 5% CO2 and were passaged about every 2 days. 2.4. Detection of integrin av and b3 expression Immunocytochemistry staining was used to detect the expression of integrin av and b3 in cell membrane. NRK52E cells were grown on coverslips; hamster monoclonal anti-rat integrin av and b3 subunits antibodies were used to detect the expression of integrin subunits av and b3, both antibody were diluted in 1:200. HRP-labeled secondary antibody and DAB (3, 3 0 -diaminobenzidine) were used to visualize the positive expression. To further determine the expression of integrin av and b3 subunits in cell membrane, flow cytometric (FCM) analysis was used. After trypsination and washing with PBS for 3 times, 106 cells were resuspended in PBS with 1 lg/ml anti-av or anti-b3 antibodies for 1 h, respectively. Then cells were washed and 1 lg/ml FITC-labeled secondary antibodies were added and incubated for 1 h. The fluorescence of FITC was detected by flow cytometry. 2.5. Cell morphological and cytoskeleton remodeling The cell morphological changes were detected by invert phase-contrast microscopy. BODIPY FL staining was used to detect the F-actin remodeling after the cells were treated with 104 lmol/L CTGF or P2 for 48 h, and 4 0 , 6-diamidino-2-phenylindole dihydrochloride hydrate (DAPI, Sigma) was used to mark the nucleus. BODIPY FL stained F-actin was observed under a laser scanning confocal microscope (LSCM) at an excitation of 512 nm and an emission of 566 nm. The blue fluorescence of DAPI stained nucleus was observed at an excitation of 488 nm and an emission of 525 nm. 2.6. Binding assay To test the binding ability of P2 and CTGF to the NRK52E cells, solid phase adhesion assay was applied [27]. After trypsination, NRK-52E cells were washed with PBS three times and were resuspended in serum-free DMEM. CTGF

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protein, P2, P2c or boiling denatured bovine serum albumin (BSA) was diluted to the same concentration of 5 · 103lmol/L and was used at 50 ll/well to pre-coat 96well plate at 4 C for 16 h. The wells were blocked with PBS containing 1% BSA for 1 h prior to the addition of 100 ll of NRK-52E cell suspension (2.5 · 105 cells/ well). Cells were incubated for 2 h to allow adhesion. The culture medium was discarded by inverting the microplate and blotting onto paper towels. Wells were washed four times gently with PBS pre-warmed to 37 C. 200 ll of CyQUANT GR dye/cell lysis buffer was added into each well and incubation in the dark at room temperature for 5 min. Adherent cells were counted by measuring the fluorescence intensity using a Bio-Tek Fluorescence Micro-plate Reader FLx800 at an excitation of 485 nm and an emission of 530 nm. All experiments were repeated 3 times. All values were expressed as mean ± SD. One-way analysis of variance (ANOVA) was used to determine statistical differences between groups using SPSS10.0 software. P values <0.05 were considered statistically significant. To determine whether P2 and CTGF competitively bind to NRK-52E cells, NRK-52E cells in 106/ml were resuspended in serum free medium to a total volume of 1 ml. All cell suspensions were simultaneously incubated with the same concentration of rhodamine-labeled CTGF at 104 lmol/L and various dosage of P2 or P2c at 0, 105, 104, 103 or 102 lmol/L, respectively. After 1 h incubation at room temperature on a rocker platform and three-time washes with PBS, all samples were analyzed by flow cytometric analyses. The fluorescence density of the samples represented the quantity of the bound rhodamine-labeled CTGF. 2.7. E-cadherin expression To detect the expression of E-cadherin in the cell membrane, immunocytochemistry staining was used. Cells were cultured on coverslips in DMEM with 104lmol/L CTGF or P2 for 48 h. Cells were fixed and incubated with rabbit anti-rat E-cadherin antibody (1:200) followed by staining with HRP-labeled secondary antibody and DAB. Further more, FCM was carried out to detect the expression of E-cadherin in cell membrane. Cells were

Fig. 2. Expression of integrin avb3 on the surface of NRK-52E cells. NRK-52E cells were cultured in DMEM, and immunohistochemistry staining was used to determine the expression of integrin av- and b3-subunits on the cell surface. Figures show that both integrin av and b3 subunits positively expressed on the cell surface. (DAB · 400).

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divided into 6 groups, group 1 was set as a blank control, neither CTGF nor P2 was added; in group 2, 104 lmol/L CTGF was added; in group 3, 104 lmol/L CTGF and 105 lmol/L P2; in group 4, 104 lmol/L CTGF and 104 lmol/L P2 were added; in group 5, 104 lmol/L CTGF and 102 lmol/L P2 were added; and in group 6, only 104 lmol/L P2 was added. Cells were cultured for 48 h, and then 106 cells were scraped and resuspended in PBS with 1 lg/ml anti-E-cadherin antibody for 1 h. Then cells were washed and 1 lg/ml FITC-labeled secondary antibodies were added and incubated for 1 h. The fluorescence of FITC was detected by flow cytometer.

Fig. 3. FCM analysis of integrin av and b3 subunits expression in NRK52E cell membrane. 106 cells were resuspended in PBS with 1 lg/ml antiav or anti-b3 antibodies for 1 h, respectively. Then cells were washed and 1 lg/ml FITC-labeled secondary antibodies were added and incubated for 1 h. The fluorescence of FITC was detected with FCM. The fluorescence intensity of FITC represented the expression of integrin av or b3 subunit. (A), the isotype control, (B) and (C), the expression of integrin av- and b3subunit, respectively.

2.8. Detecting mRNA level of a-SMA, collagen type I and collagen type IV NRK-52E cells were divided into 6 groups as described above. Total RNA was extracted from each group of cells using a SV total RNA isolation kit. Reverse transcription PCR was used to examine the mRNA transcription of aSMA, collagen type I and collagen type IV. All the primers used were designed with Primer 5 software, including aSMA, sense- 5 0 -CAA CTG GTA TTG TGC TGG ACT C-3 0 , antisense 5 0 -ACA TCT GCT GGA AGG TAG ACA G-3 0 (a PCR product of 624 bp); collagen type I, sense 5 0 -ACT GTC CCA ACC CCC AAA A-3 0 , antisense 5 0 -GAC AGC ACC ATC GTT ACC AC-3 0 (739 bp); collagen type IV, sense 5 0 -CTG CTC TGT CTG CCA GTG TTT-3 0 , antisense 5 0 -AGC CAT TGT AGC CGT CCA TA-3 0 (278 bp); and glyceraldehyde phosphate dehydrogenase (GAPDH), sense 5 0 -ATC ACC ATC TTC CAG GAG CGA GA-3 0 , antisense 5 0 -GCT TCA CCA CCT TCT TGA TGT CA -3 0 (573 bp). The reverse transcription reactions were carried out at 50 C for 30 min, and then the samples were heated to 94 C for 2 min followed by a course of 35 cycles at 94 C for 30 s, 57 C for 30 s, and 72 C for 30 s. After the last cycle, a final extension step was taken at 72 C for 7 min. The results were normalized on GAPDH gene expression. The RT-PCR products were electrophoresed on 1.5% agarose gels, stained with Goldview dye, and quantified by densitometric analysis (Image Pro-Plus), and all the results were described as a ratio of relative optical density (ROD) value of the band of the interested gene to GAPDH.

Fig. 4. Solid phase binding assay of CTGF and P2 to NRK-52E cells. 5 · 103 lmol/L CTGF, P2, P2c or BSA was used at 50 ll/well to pre-coat 96-well plate at 4 C for 16 h. The wells were blocked with PBS containing 1% BSA, 100 ll of NRK-52E cell suspension (2.5 · 105 cells/well) were added into the wells. After 2 h incubation, the culture medium was discarded. 200 ll of CyQUANT GR dye/cell lysis buffer was added into each well and the plates were incubated in the dark for 5 min. Adherent cells were counted by measuring the fluorescence intensity using a Bio-Tek Fluorescence Micro-plate Reader FLx800 with an excitation of 485 nm and an emission of 530 nm. All experiments were repeated 3 times. P values were significant against blank control. *P < 0.05 vs. CTGF group, and nP < 0.05 vs. P2 group. No significant difference existed between CTGF and P2 group.

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Fig. 5. Competitive binding between CTGF and P2. 104 lmol/L rhodamine-labeled CTGF and various dosage of P2 or P2c at 0, 105, 104, 103 or 102 lmol/L were added to the medium simultaneously. After 1 h incubation, the combined red fluorescence of rhodamine-labeled CTGF was analyzed by flow cytometer. With the increasing concentration of P2, as was shown in (B) to (F), the fluorescence intensity decreased. When 104 or 102 lmol/L P2c was added, the fluorescence intensity was not changed (G and H). (A) was set as the isotype control.

Fig. 6. Effects of CTGF and P2 on cytoskeleton remodeling. To explore the effect of CTGF and P2 on cell cytoskeleton, NRK-52E cells were incubated in DMEM with 104 lmol/L CTGF or P2 for 48 h, BODIPY FL was used to stain the F-actin. These figures were the representative examples of the F-actin remodeling induced by CTGF (A) or P2 (B) and (C) is the negative control. Figures showed that treated with CTGF, F-actin was significantly upregulated, and large microfilaments developed in CTGF-treated cells, whereas no such changes were observed in P2-treated cells, as compared with the control cells. Furthermore, CTGF-treated cells presented enlarged size (LSCM · 600).

2.9. Detecting protein expression of a-SMA, collagen type I and collagen type IV NRK-52E cells were divided into 6 groups pretreated with different concentration of CTGF and P2 as described above. Total cell protein was extracted using M-PER Mammalian Protein Extraction Reagent. The protein

concentration was determined by the Bradford method (Bio-Rad). Ten microliters of total protein in 10 · loading buffer of each groups was loaded and separated on 10% sodium dodecyl sulfate–polyacrylamide gel electropheresis (SDS–PAGE). The protein was transferred to PVDF membrane at 400 mA for 1 h in transfer buffer (25 mM Tris, 0.2 M glycine, and 20% methanol). Membrane was blocked

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diluted collagen type IV rabbit polyclonal antibody, and 1:1 000 diluted b-actin mouse monoclonal antibody were used to detect the expression of a-SMA, collagen type I and collagen type IV, respectively. b-Actin was used as the loading control. The proper species and diluted HRPlabeled second antibodies were added. Western blot results were detected by the SuperSignal West Pico Chemiluminescent Substrate with a 30 s exposure time, the films were developed in developer and the results were scan by an Epson scanner. 3. Results 3.1. Expression of integin av and b3 subunits in NRK-52E cells Immunocytochemistry staining showed that both the integrin av and b3 subunits were positively expressed on the cells surface (Fig. 2). Strong fluorescence was also detected by flow cytometry on the surface of cells incubated with anti-integrin av or b3 subunits antibodies. (Fig. 3). 3.2. Binding of CTGF and P2 to NRK-52E cells Solid phase adhesion assay was used to characterize the interaction between P2 and NRK-52E cells. The fluorescence intensity detected by micro-plate reader represented the remained cells. After 2 h incubation and four-time washes with PBS, a large amount of cells adhered to the surface precoated with CTGF or P2, while only a few cells attached to the uncoated surface or P2c and BSA coated surface. (P < 0.05) (Fig. 4). In the competitive binding assay, the same concentration of rhodamine-labeled CTGF and variant amount of P2 or P2c were added to the medium simultaneously. The fluorescence intensity of rhodamine represented the quantity of bound CTGF. In P2-free group, high fluorescence intensity presented, and the increasing concentration of P2 was coupled with the decreased fluorescence intensity. However, P2c had no effect on the CTGF binding (Fig. 5). 3.3. Cytoskeleton remodeling effect of CTGF and P2 Fig. 7. Immunocytochemistry staining of the expression of E-cadherin in cell membrane. Cells were treated with 104 lmol/L CTGF or P2 for 48 h. Rabbit anti-rat E-cadherin antibody (with a dilution of 1:200) was added, HRP-labeled secondary antibody and DAB were used to visualize the positive expression. (A) The blank control, which showed that E-cadherin was positively expressed in cell membrane. (B and C) Cells treated with P2 or CTGF, respectively. In CTGF group, E-cadherin was negatively or very weakly expressed, and some cells presented elongated shape and enlarged size and lost contact with their neighbor. However, in P2 group, positive expression of E-cadherin was presented and no phenotype changes happened. (DAB · 400).

in Tris-buffered saline–Tween 0.1% (TBST) with 7% skimmed milk powder for 1 h at room temperature. The 1:100 diluted a-SMA mouse monoclonal antibody, 1:400 diluted collagen type I goat polyclonal antibody, 1:200

The effect of CTGF and its homologous peptide P2 on cytoskeleton remodeling in NRK-52E cells were evaluated. It was observed that after being cultured in DMEM with CTGF for 48 h, F-actin, the important component of cytoskeleton, was significantly upregulated, and large microfilaments were developed, whereas no such changes were observed in P2 -treated cells as compared with the control cells (Fig. 6). 3.4. Regulation of a-SMA and E-cadherin expression induced by CTGF and P2 E-cadherin is considered as a membrane protein marker on the surface of epithelial cells. The disappearance of E-

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Fig. 8. FCM analysis of the expression of E-cadherin on the surface of cells treated with different concentration of CTGF and P2. Cells were divided into 6 groups, group 1 (A) was set as a blank control, neither CTGF nor P2 was added; group 2 (B), 104 lmol/L CTGF was added; group 3 (C), 104 lmol/L CTGF and 105 lmol/L P2 were added; group 4 (D), 104 lmol/L CTGF and 104 lmol/L P2 were added; group 5 (E), 104 lmol/L CTGF and 102 lmol/L P2 were added and group 6 (F), only 104 lmol/L P2 was added. Cells were cultured for 48 h, and then 106 cells were resuspended in PBS with 1 lg/ml anti-E-cadherin antibody for 1 h. Then cells were washed and 1 lg/ml FITC-labeled secondary antibodies were added and incubated for 1 h. The fluorescence of FITC was detected by flow cytometry.

cadherin is a milestone for EMT. Using immunocytochemistry staining, positive expression of E-cadherin was detected in the normal cultured cells (blank control group) and the P2 treated cells. However, in CTGF treated group, only very weak expression of E-cadherin was presented (Fig. 7). FCM analysis was carried out to investigate the E-cadherin expression in NRK-52E cells membrane. As was shown in Fig. 8, CTGF sharply suppress the expression of E-cadherin. However, when P2 was added in the medium, the suppression effect of CTGF on E-cadherin was inhibited in a dose-dependent manner. When 104 lmol/ L CTGF and 102 lmol/L P2 were added simultaneously, CTGF effect almost been eliminated. a-SMA, a specific molecular marker of activated ECM producing cells, was also measured by reverse transcription PCR and Western blot. a-SMA expression was markedly elevated at both mRNA levels (Fig. 9) and protein levels (Fig. 10) in CTGF-treated cells, but remained unchanged in P2-treated cells. Further more, the upregulating effect of CTGF on a-SMA was inhibited by P2 in a dose-dependent manner (Figs. 9 and 10).

3.5. Effect of CTGF and P2 on the expression of collagen type I and collagen type IV The mRNA of collagen type I and collagen type IV was markedly upregulated by 104 lmol/L CTGF, but not by P2. Moreover, the upregulating effect of CTGF on mRNA

and protein levels of collagen type I and IV was significantly inhibited by P2 in a dose-dependent manner. (Figs. 9 and 10). 4. Discussion To determine whether the synthesized peptide P2 bind to NRK-52E cells and block up the CTGF signal transduction, we tested the binding ability of P2 to NRK-52E cells and its effect on the epithelial–mesenchymal transition (EMT) induction. Our studies show that both CTGF and P2 can bind to NRK-52E cells. Integrin avb3 might be the membrane receptor of CTGF. P2 competitively inhibit CTGF binding to the cells, however, P2 has no effect on the F-actin remodeling, EMT and extracellular matrix induction. These results indicate that P2 might be used as a CTGF inhibitor to block up the CTGF signal pathway in EMT induction and renal interstitial fibrosis. As a para- and endocrine cytokine, CTGF receptor in cell membrane is still uncertain [26,27,31]. CTGF comprises four modules and its C-terminus is a cysteine rich region which contains the binding domain for its receptors [32]. Several integrins are predicted to be potential receptors of CTGF [26,27,33–35], such as integrin avb3, which plays a crucial role in the activation of hepatic stellate cells (HSCs) during liver fibrosis [27]. In our studies, CTGF receptor on NRK-52E cells was not clarified, however, we confirmed that both the integrin av and b3 subunits positively express on the cell surface. Similar to HSCs, integrin avb3 is the receptor and binding domain for CTGF on

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Fig. 9. Gene transcription level of a-SMA, collagen type I and collagen type IV in NRK-52E cells. NRK-52E cells were treated with various concentrations of CTGF and P2, total RNA was purified and one step reverse transcription PCR was carried out. Cells were divided into 6 groups, group 1 was set as a blank control, neither CTGF nor P2 was added (lane 1); group 2, 104 lmol/L CTGF was added (lane 2); group 3, 104 lmol/L CTGF and 105 lmol/L P2 were added (lane 3); group 4, 104 lmol/L CTGF and 104 lmol/L P2 were added (lane 4); group 5, 104 lmol/L CTGF and 102 lmol/L P2 were added (lane 5) and group 6, only 104 lmol/L P2 was added (lane 6). The upper illustrate (A) Gave one of the three times repeated RT-PCR results of a-SMA, collagen type I, collagen type IV and the internal control GAPDH. The lower illustrate (B) Showed that with the increasing concentration of P2 in the medium, the transcriptional levels of a-SMA, collagen type I and collagen type IV were downregulated.

Fig. 10. NRK-52E cells were treated with various concentrations of CTGF and P2, Western blot was carried out to detect the protein expression of a-SMA, collagen type I and collagen type IV. Cells were divided into 6 groups, group 1 was set as a blank control, neither CTGF nor P2 was added; group 2, 104 lmol/L CTGF was added; group 3, 104 lmol/L CTGF and 105 lmol/L P2 were added; group 4, 104 lmol/ L CTGF and 104 lmol/L P2 were added; group 5, 104 lmol/L CTGF and 102 lmol/L P2 were added and group 6, only 104 lmol/L P2 was added. (A–C) Gave one of the three times Western blot results of a-SMA, collagen type I and collagen type IV, respectively, and (D), the expression of b-actin served as loading control.

the surface of NRK-52E cells, blockage of the interaction between CTGF and integrin avb3 may obstruct the CTGF signal pathway. Previous study has elucidated the unique binding domain in CTGF to integrin avb3 [27]. In this study, the 16-residual peptide homologous to the binding domain was synthesized and named as P2. Our results show that,

similar to the whole CTGF protein, P2 acted as an adhesion molecule to mediate solid phase adhesion of NRK52E cells. These results suggest that P2 is a potential ligand for integrin avb3. P2c, the antisense sequence of P2, could not mediate the cell binding. To investigate whether competitive binding exist between CTGF and P2, rhodamine-labeled CTGF and P2 were added in the medium simultaneously. As was seen in the flow cytometric analyses, the increasing concentration of P2 made the fluorescence of CTGF decrease. The results indicate that this much smaller molecule P2 is capable of binding to the cells preferably and can competitively inhibit CTGF binding. When the P2 concentration was raised to 100 times more than CTGF, CTGF binding would almost be eliminated. However, P2c had no effect on CTGF binding. EMT is a complex process in which renal TECs lose their polarized phenotype and acquire new features characteristic of myofibroblasts, the major effector cells responsible for the excess deposition of interstitial ECM under pathologic conditions [36]. E-cadherin is an adhesive junction protein expressed in differentiated and polarized epithelial cells [37], such as tubular epithelial cells. Under pathologic conditions, tubular epithelial cells lose E-cadherin. The disappearance of E-cadherin in epithelial cell indicates that the cell undergoes an EMT process [38]. In this study, we reveal that induced by CTGF, the expression of E-cadherin in NRK-52E cells is markedly suppressed, even though the phenotype change is not apparent. On the contrary, treated with P2, the expression of E-cadherin shows no different compared with the control cells. Further more, when P2 is added in the medium, the EMT induction effect of CTGF is sharply suppressed. These results suggest that P2 might have no effect on EMT induction. In addition, P2 might act as a CTGF inhibitor, when both CTGF and P2 are added in the medium, P2 will occupy the binding domain of CTGF on NRK-52E cells, and thus CTGF could not exert its function. Both clinical studies and animal experiments indicate a strong positive correlation between the loss of renal function and the emergence of myofibroblasts. a-SMA is a hallmark of the activated myofibroblasts, and its positive expression in tubular epithelial cells means that the cells lose their epithelial phenotype and obtain myofibroblasts phenotype [39–41]. Our studies have shown that when CTGF alone is added in the culture medium, markedly elevated a-SMA mRNA and protein are detected in NRK52E cells. As a specific molecular marker of activated myofibroblasts, the high expression level of a-SMA is associated with the NRK-52E cells that differentiate from epithelial cells to myofibroblasts. These findings suggest that CTGF can activate the signal pathway to invoke the cell transition. When P2 along is added, cells show no change in aSMA expression. Both collagens type I and IV are the fundamental extracellular matrix proteins [42,43]. Once secreted by ECM producing cells, collagens type I and IV accumulate in

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the basement or interstitial space. The disordered synthesis and secretion and unsatisfactory degradation lead to the excessive deposition of ECM components and destruction of tissue architecture [44]. Our data also show that P2 did not regulate the expression of collagen type I and IV in mRNA or protein levels and had no effect on the cytoskeleton remodeling. When CTGF and P2 were added in the medium simultaneously, P2 robustly attenuated CTGF-mediated EMT induction and fibrogenesis. Among the many strategies that have been applied to ameliorate the function of renal allograft [45–50], delay or reversion of EMT is considered to be most important one. Because of its important role in the interstitial fibrosis, CTGF is considered as a potential target for preventing renal allograft fibrosis [50–52]. P2, a small peptide homologous to CTGF, could act as a novel inhibitor of CTGF and provide a promising therapeutic strategy in CAN. In this in vitro study, rat TECs are used rather than human primary cells. Human TECs would be much better than rat cell line to study the P2 effect. However, as we will test its in vivo effect, the rat model may be better and P2 will be adequately modified for further in vivo studies. Acknowledgements This study was supported by a grant from the National Basic Research Program of China (No. 2003CB515504), by the Program for Changjiang Scholars and Innovative Research Team in University, Ministry of Education, and by the National Natural Scientific Foundation (No. 30571761) and the Foundation of China Medical Board of New York Inc. (CMB, Inc.). References [1] Okada H, Kalluri R. Cellular and molecular pathways that lead to progression and regression of renal fibrogenesis. Curr Mol Med 2005;5:467–74. [2] Hmiel SP, Beck AM, de la Morena MT, Sweet S. Progressive chronic kidney disease after pediatric lung transplantation. Am J Transpl 2005;5:1739–47. [3] Chatziantoniou C, Boffa JJ, Tharaux PL, Flamant M, Ronco P, Dussaule JC. Progression and regression in renal vascular and glomerular fibrosis. Int J Exp Pathol 2004;85:1–11. [4] Nangaku M. Mechanisms of tubulointerstitial injury in the kidney: final common pathways to end-stage renal failure. Intern Med 2004;43:9–17. [5] Nankivell BJ, Borrows RJ, Fung CL, O’Connell PJ, Allen RD, Chapman JR. The natural history of chronic allograft nephropathy. N Engl Med 2003;349:2326–33. [6] Djamali A, Reese S, Yracheta J, Oberley T, Hullett D, Becker B. Epithelial-to-mesenchymal transition and oxidative stress in chronic allograft nephropathy. Am J Transpl 2005;5:500–9. [7] Djamali A, Premasathian N, Pirsch JD. Outcomes in kidney transplantation. Semin Nephrol 2003;23:306–16. [8] Furness PN. Histopathology of chronic renal allograft dysfunction. Transplantation 2001;71(Suppl):SS31–36. [9] Schwarz A, Mengel M, Gwinner W, Radermacher J, Hiss M, Kreipe H, et al. Risk factors for chronic allograft nephropathy after renal transplantation: a protocol biopsy study. Kidney Int 2005;67:341–8.

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