Role of endothelial-to-mesenchymal transition induced by TGF-β1 in transplant kidney interstitial fibrosis

Abstract

Chronic allograft dysfunction (CAD) induced by kidney interstitial fibrosis is the main cause of allograft failure in kidney transplantation. Endothelial-to-mesenchymal transition (EndMT) may play an important role in kidney fibrosis. We, therefore, undertook this study to characterize the functions and potential mechanism of EndMT in transplant kidney interstitial fibrosis. Proteins and mRNAs associated with EndMT were examined in human umbilical vein endothelial cells (HUVECs) treated with transforming growth factor-beta1 (TGF-β1) at different doses or at different intervals with western blotting, qRT-PCR and ELISA assays. Cell motility and migration were evaluated with motility and migration assays. The mechanism of EndMT induced by TGF-β1 was determined by western blotting analysis of factors involved in various canonical and non-canonical pathways. In addition, human kidney tissues from control and CAD group were also examined for these proteins by HE, Masson's trichrome, immunohistochemical, indirect immunofluorescence double staining and western blotting assays. TGF-β1 significantly promoted the development of EndMT in a time-dependent and dose-dependent manner and promoted the motility and migration ability of HUVECs. The TGF-β/Smad and Akt/mTOR/p70S6K signalling pathways were found to be associated with the pathogenesis of EndMT induced by TGF-β1, which was also proven in vivo by the analysis of specimens from the control and CAD groups. EndMT may promote transplant kidney interstitial fibrosis by targetting the TGF-β/Smad and Akt/mTOR/p70S6K signalling pathways, and hence, result in the development of CAD in kidney transplant recipients.

Keywords: Akt/mTOR/p70S6K; Smad; chronic allograft dysfunction; endothelial-to-mesenchymal transition; kidney interstitial fibrosis; kidney transplantation; transforming growth factor-beta1.

Figure 1

Indirect immunofluorescence staining of cells extracted from human umbilical cord segments. (A) The monolayer cells are observed to be arranged like ‘paving stones’ under the light microscope (×100). (B) CD31 (represented by green fluorescent signals) was mainly distributed in the cytoplasm of HUVECs (×200). (C) The nuclei of HUVECs were stained with DAPI (blue) (×200). (D) Positive staining signals for CD31 and DAPI (merged) in cells extracted from human umbilical cord segments prove that the cells were endothelial cells (×200).

Figure 2

TGF‐β1 promotes α‐SMA and collagen I expression and suppresses VE‐cadherin and CD31 expression in the HUVECs. (A and B) Equal amounts of protein from whole cell lysates were analysed by western blotting with antibodies against α‐SMA, collagen I, VE‐cadherin, CD31 and GAPDH after incubation of HUVECs with 5 ng/ml TGF‐β1 for the indicated time points (A) or stimulation of HUVECs with various concentrations of TGF‐β1 for 48 hrs (B). As can be seen, TGF‐β1 promoted the expression of α‐SMA and collagen I and suppressed the expression of VE‐cadherin and CD31 in a time‐ and dose‐dependent manner. (C–J) HUVECs were stimulated with TGF‐β1 (5 ng/ml) for the indicated time points (C, E, G and I) or stimulated with various concentrations of TGF‐β1 for 48 hrs (D, F, H and J). Total RNA was isolated and reverse transcribed, and the resultant RNA was subjected to quantitative real‐time PCR to detect the gene expression of α‐SMA, collagen I, VE‐cadherin and CD31. The results of quantitative real‐time PCR were normalized to β2‐macroglobulin expression and expressed as the fold‐change relative to unstimulated control cells. Relative abundance of mRNAs is presented as the mean ± S.D. value of three independent experiments. The PCR results were in agreement with the western blot results. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 versus the control group, as determined by one‐way anova (C‐J).

Figure 3

TGF‐β1 promotes extracellular matrix secretion in HUVECs. (A–D) HUVECs were incubated with various concentrations of TGF‐β1 for 48 hrs (A and C) or stimulated with 5 ng/ml TGF‐β1 for the indicated time points (B and D). The supernatant of the cultured HUVECs was collected for ELISA to determine the total concentration of collagen I (A and B) and fibronectin (C and D). The relative abundance of proteins was presented as the mean ± S.D. values of three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 versus the control group by one‐way anova (A–D).

Figure 4

TGF‐β1 promotes the motility and migration of HUVECs. (A and B) HUVECs were wounded with a pipette and treated with 5 ng/ml TGF‐β1 for the indicated time points (A). The migrated cells were quantified by manual counting, and the motility index was determined using the formula ‘motility index = the number of cells that migrated in the control group/the number of cells that migrated in the TGF‐β1 group’ (B). (C and D) A total of 5 × 10⁴ HUVECs were seeded in the top chamber and treated with 5 ng/ml TGF‐β1 for the indicated time points (C). Cells that migrated through the membrane were stained and quantified. The migration index was determined using the formula ‘migration index = the number of cells that had migrated at 48 hrs/the number of cells that had migrated at 0 hr’ (D). The motility and migration indexes were expressed as the mean ± S.D. value of five independent experiments. **P < 0.01, ***P < 0.001 versus the control group, by one‐way anova (B) or Student t test (D).

Figure 5

TGF‐β1 upregulates α‐SMA expression in HUVECs through the TGF‐β/Smad and Akt/mTOR/p70S6K signalling pathways. (A) HUVECs were treated with TGF‐β1 (5 ng/ml) for the indicated time points. Equal amounts of protein were collected from whole cell lysates and analysed by western blotting with antibodies against phosphorylated Smad 2, Smad 3, Akt, mTOR, p70 S6K, p38 MAPK, Erk1/2 and c‐Jun. (B, D, E, G, I and K) HUVECs were pre‐treated for 1 hr with MK2206 (10 μmol/l) (B and D), SB431542 (10 μmol/l) (E), SB203580 (10 μmol/l) (G), UO126 (10 μmol/l) (J) or SP600125 (5 μmol/l) (K), and specific chemical inhibitors of Akt, Smad, p38MAPK, Erk1/2 and JNK. Subsequently, cells were treated with TGF‐β1 (5 ng/ml) for the indicated time points. Cells were collected 1 hr after TGF‐β1 stimulation. Equal amounts of protein from whole cell lysates were analysed by western blotting with antibodies against phosphorylated and total Akt (B and D), phosphorylated and total mTOR (D), phosphorylated and total p70S6K (D), phosphorylated and total Smad 2 (E), phosphorylated and total Smad 3 (E), phosphorylated and total p38 MAPK (G), phosphorylated and total Erk1/2 (I), and phosphorylated and total c‐Jun (K). (B, E, G, I and K) HUVECs were collected 48 hrs after TGF‐β1 stimulation. Equal amounts of protein from whole cell lysates were analysed by western blotting with antibodies against α‐SMA and GAPDH. The ratio of α‐SMA to GAPDH density was expressed as the fold‐change relative to unstimulated control cells (C, F, H, J and L). The results of densitometric determination of the relative abundance of α‐SMA are presented as the mean ± S.D. value of three independent experiments. ^a P > 0.05 versus the control cells; ^b P < 0.01 versus the control cells; ^c P < 0.01 and ^d P > 0.05, cells treated with TGF‐β1 versus cells treated with TGF‐β1 and specific inhibitors, by Student t test (C, F, H, J and L).

Figure 6

Increase in TGF‐β1, α‐SMA, collagen I and collagen III expression; decrease in CD31 expression; and kidney interstitial fibrosis in kidney tissues from renal transplant recipients with CAD. (A and B) Representative tissue sections from the control and CAD group were stained with HE (A) and Masson‐Trichrome (B) stain. (C) Quantitative analysis of the fibrosis intensity of kidney sections stained with Masson‐Trichrome was performed. The results demonstrated a significant degree of fibrosis in kidney tissues from renal transplant recipients with CAD. (D–H) Distribution and expression of TGF‐β1 (D), α‐SMA (E), collagen I (F), collagen III (G) and CD31 (H) in the CAD and control group were assessed by immunohistological staining assays. Fibrosis percentage and relative abundance of proteins were presented as the mean ± S.D. value of three independent experiments. Representative images of kidney tissues from the control group (n = 25) and CAD group (n = 25) are shown. ****P < 0.0001 versus the control group, as determined by the Student t test (C, D–H).

Figure 7

TGF‐β1 promotes the development of EndMT via the TGF‐β/Smad and Akt/mTOR/p70S6K signalling pathways in kidney tissues from renal transplant recipients with CAD. (A–D) Kidney tissues from the control group (A) and renal transplant recipients with CAD (B–D) were examined. The distribution and expression of α‐SMA (represented by green fluorescent signals) and vWF (represented by red fluorescent signals) were analysed by the indirect immunofluorescence double‐staining assay. (E, F) Equal amounts of proteins from human kidney tissues were analysed by western blotting with antibodies against TGF‐β1 (E), α‐SMA (E), collagen I (E), VE‐cadherin (E), CD31 (E), phosphorylated Smad 2 (F), Smad 3 (F), Akt (F), mTOR (F), p70 S6K (F) and GAPDH (F). (G) A model is proposed to illustrate the fibrotic mechanism involved in EndMT induced by TGF‐β1 in the pathogenesis of kidney interstitial fibrosis in kidney transplant recipients with CAD.