Identification of a Klotho-derived peptide that blocks TGF-β signaling
We first designed and synthesized 18 overlapping peptides that encompassed the KL1 domain of human Klotho protein. Using the TGF-β1-treated normal rat kidney interstitial fibroblast (NRK-49F) cells as an in vitro system, we screened these peptides for their ability to inhibit myofibroblastic activation of NRK-49F cells (Supplementary Fig. 1). After several rounds of screening, we identified a peptide with 30 amino acids encompassing from Phe57 to Lys86 of Klotho protein (Fig. 1a), designated as the Klotho-derived peptide 1 (KP1), which abolished TGF-β1 action in kidney cells. Bioinformatics analyses revealed a high degree of homology in the amino acid sequence of KP1 among different species ranging from human to fruit fly (Fig. 1a), suggesting high evolutionary conservation of KP1.
a Amino acid sequence alignment of KP1 in various species. A light green background indicates the regions with identical amino acids in different species. Differences in amino acids among various species are highlighted by red color. b KP1 inhibited the induction of α-SMA mRNA by TGF-β1 in NRK-49F cells. NRK-49F cells were incubated with TGF-β1 (2 ng/ml) in the absence or presence of KP1 (10 µg/ml) or sKlotho (10 µg/ml). The expression of α-SMA mRNA was assessed by qRT-PCR. P values (from left to right): <0.001, 0.001, <0.001. n = 4 biologically independent cells. c–e KP1 inhibited the protein expression of fibronectin and α-SMA. NRK-49F cells were preincubated with KP1 (10 µg/ml; 3 µM) for 1 h and then treated with TGF-β1 (2 ng/ml) for 24 h. Western blot (c) and quantitative data (d, e) are presented. P values (from left to right): 0.001 and 0.004 (fibronectin); 0.017 and 0.026 (α-SMA). n = 3 biologically independent cells. f KP1 inhibited TGF-β1-mediated fibronectin expression and deposition by NRK-49F cells. Representative micrographs with immunofluorescence staining of fibronectin are shown. Scale bar, 50 µm. Arrows indicated positive staining. g–i KP1 inhibited the expression of fibronectin and α-SMA in primary tubular cells. Western blot analyses (g) and quantitative data of fibronectin (h) and α-SMA (i) are shown. P values (from left to right): 0.001 and 0.001 (fibronectin); 0.001 and 0.003 (α-SMA). n = 3 biologically independent cells. j–l The protein level of fibronectin and collagen I expressed by primary cardiomyocytes. Western blot analyses (j) and quantitative data of fibronectin (k) and collagen I (l) are shown. P values (from left to right):0.001 and 0.001 (fibronectin); <0.001 and <0.001 (collagen I). n = 3 biologically independent cells. Ctrl controls. Data are presented as mean values ± SEM. Statistical significance was determined by one-way ANOVA followed by Fisher’s Least-significant Difference (LSD) post hoc test. Source data are provided as a Source Data file.
We then incubated NRK-49F cells with TGF-beta1 in the absence or presence of KP1. As shown in Fig. 1b, KP1 repressed the TGF-β1-induced mRNA expression of α-SMA, suggesting its ability to abolish the myofibroblastic activation of renal fibroblasts. The inhibitory effect of KP1 on α-SMA was comparable to soluble Klotho (sKlotho) (Fig. 1b). Similarly, KP1 inhibited α-SMA protein expression in NRK-49F cells after incubation with TGF-β1 (Fig. 1c, e). KP1 also inhibited fibronectin expression induced by TGF-β1 (Fig. 1c, d, f), as revealed by Western blotting and immunofluorescence staining.
To generalize this finding, we examined the effect of KP1 on other types of cells after TGF-β stimulation. To this end, we utilized four additional types of primary cells or cell lines from mouse, rat, and human. As shown in Fig. 1g–i and Supplementary Fig. 2a–c, KP1 also inhibited fibronectin and α-SMA expression induced by TGF-β1 in mouse primary kidney tubular epithelial cells and human proximal tubular cells (HKC-8). Similarly, KP1 abolished TGF-β1-triggered fibronectin and collagen I expression in rat primary cardiomyocytes (Fig. 1j–l) and rat primary cardiac fibroblasts (Supplementary Fig. 2d–f). Together, these results show that KP1 is capable of blocking matrix gene expression by intercepting TGF-β signaling in various types of cells.
KP1 binds to TβR2 and disrupts TGF-β signaling
We next investigated the mechanism by which KP1 inhibits TGF-β1 signaling. As the binding of TGF-β with its type 2 receptor (TβR2) is the initial step in TGF-β signal transduction, we first examined the potential interaction between KP1 and TβR2. To this end, KP1 was labeled with fluorescein isothiocyanate (FITC) and then incubated with NRK-49F cell lysate. As shown in Fig. 2a, FITC-labeled KP1 was detected in the immunocomplexes precipitated by anti-TβR2 antibody, suggesting its interaction with TβR2. In the reciprocal experiment, TβR2 was also readily identified in the immunocomplexes precipitated by anti-FITC antibodies (Fig. 2b). To study the binding affinity of KP1 to TβR2, we employed the surface plasmon resonance (SPR) analyses with Biacore T200, an optical and label-free technique used to measure molecular interactions in real time29,30,31. As shown in Fig. 2c, the binding affinity between KP1 and TβR2 was strong and the equilibrium dissociation constant (KD) calculated was 1.41 µM, whereas KP12, a negative control peptide that did not inhibit TGF-β action (Supplementary Fig. 1), showed less affinity with TβR2. The KD between KP12 and TβR2 was 14.6 μM (Fig. 2d, Supplementary Table 1). We also assessed the binding between KP1 and TβR2 by directly incubating FITC-KP1 on a TβR2-coated microplate. As shown in Fig. 2e, significant binding of FITC-KP1 to TβR2 was evident, as a substantial amount of fluorescence was detected after washing. Under the same conditions, much less FITC-KP12 was detected when it was incubated on the TβR2-coated microplate (Fig. 2e). As a native control, neither FITC-KP1 nor FITC-KP12 bound to bovine serum albumin (BSA) coated on the microplate (Fig. 2e), suggesting the specificity of KP1 and TβR2 interaction.
a, b Co-immunoprecipitation (Co-IP) demonstrated that KP1 bound to TβR2. NRK-49F cell lysates (500 µg) and FITC-KP1 (10 µg) were immunoprecipitated (IP) with the anti-TβR2 antibody at 4 °C overnight, followed by immunoblotted (IB) for FITC and TβR2, respectively (a). In the reciprocal experiment, mixtures of cell lysates and FITC-KP1 were immunoprecipitated with anti-FITC, followed by immunoblotted for TβR2 and FITC (b). c The affinity of TβR2 to KP1 was shown. SPR analyses showed concentration-dependent (0.78–25 μM) binding of KP1 to TβR2 immobilized on a sensor chip. A representative sensorgram of two independent experiments is presented, in which the curves (color) with an overlay of the fitting (black) are shown. The fitted constants are ka = 407.7 M−1 S−1; kd = 5.8 × 10−4 S−1; KD = 1.4 × 10−6 M. d The affinity of TβR2 to KP12 was shown. SPR analyses showed concentration-dependent (1.56–100 μM) binding of KP12 to TβR2 immobilized on a sensor chip. A representative sensorgram of two independent experiments is presented, in which the curves (color) with an overlay of the fitting (black) are shown. The fitted constants are ka = 159.5 M−1 S−1; kd = 2.3 × 10−3 S−1; KD = 14.6 × 10−6 M. e The specific binding of FITC-KP1 to TβR2 coated on a microplate. The fluorescence intensity of FITC-KP1 (arbitrary units) was assessed after incubating on a microplate coated with TβR2 protein. P values (from left to right): <0.001, <0.001, and <0.001 (one-way ANOVA with Fisher’s LSD post hoc test). (n = 4 biologically independent cells). Data are presented as mean values ± SEM. f KP1 inhibited the interaction between TβR2 and TGF-β1 dose-dependently. NRK-49F cells were pre-incubated with different amounts of KP1 for 1 h, and then treated with TGF-β1 (2 ng/ml) for 5 min. Cells were collected and immunoprecipitated with anti-TβR2 or anti-IgG. g Negative peptide KP12 did not block the binding of TβR2 and TGF-β1. h KP1 inhibited the binding of TβR1 and TβR2. i KP1 inhibited the interaction of TβR2 and sKlotho. Cells were transfected with sKlotho plasmid for 24 h and incubated with TGF-β1 in the absence or presence of KP1. Cells were collected and immunoprecipitated with anti-TβR2. Source data are provided as a Source Data file.
To investigate the functional consequence of KP1 and TβR2 interaction, we examined its influence on the binding of TGF-β and TβR2. To this end, NRK-49F cells were pretreated with KP1 and then incubated with TGF-β1. As shown in Fig. 2f, TGF-β1 was able to bind to TβR2 and formed a complex with it, as revealed by co-immunoprecipitation. However, such TGF-β1/ TβR2 interaction was abolished by incubation with KP1 in a dose-dependent manner (Fig. 2f), suggesting that KP1 blocks TGF-β signaling by disrupting the ligand–receptor engagement. Under the same conditions, KP12 was unable to disrupt the TGF-β1/TβR2 interaction (Fig. 2g). Consistently, KP12 failed to inhibit TGF-β1-induced fibronectin and α-SMA expression in NRK-49F cells (Supplementary Fig. 3a–c). TβR2 forms heterodimer with TβR1 after TGF-β1 binding to TβR212. Thus, we tested whether KP1 inhibited the binding and recruitment of TβR1. As shown in Fig. 2h, KP1 interrupted the binding of TβR1 and TβR2. As Klotho is also able to bind to TβR2, we further examined whether KP1 can compete with it for binding. As shown in Fig. 2i, sKlotho was able to interact with TβR2, which was abolished in the presence of KP1. These results suggest that KP1 acts as an antagonist of TGF-β signaling by binding to TβR2 and disrupting TGF-β/TβR2 engagement.
KP1 inhibits multiple downstream signaling of TGF-β in vitro
We further investigated the effect of KP1 on the downstream signaling of TGF-β. We first assessed the phosphorylation and activation of Smad-2 and -3 (Smad2/3), the canonical pathway mediating TGF-β actions. As shown in Fig. 3a, Smad2/3 was phosphorylated and activated after incubation with TGF-β1 in NRK-49F cells, which was inhibited by KP1. Quantitative determination of p-Smad2/3 protein levels in different groups was presented in Fig. 3b, c. Immunofluorescent staining revealed that p-Smad3 was accumulated in the nuclei of NRK-49F cells after TGF-β1 stimulation (Fig. 3d). However, incubation with KP1 largely abolished the TGF-β1-triggered p-Smad3 nuclear accumulation (Fig. 3d).
a KP1 inhibited the phosphorylation of Smad2 and Smad3 induced by TGF-β1. Numbers (1–3) indicate each individual wells of cells. Serum-starved NRK-49F cells were pre-treated with KP1 or vehicle for 1 h and then stimulated by TGF-β1 for 45 min. b, c Quantitative data of p-Smad3 (b) and p-Smad2 (c) in different groups are shown. P values (from left to right): <0.001 and 0.001 (p-Smad3); <0.001 and 0.001 (p-Smad2). n = 3 biologically independent cells. d Immunofluorescence staining of p-Smad3 showed that KP1 blocked TGF-β1-mediated Smad3 phosphorylation and nuclear translocation. DAPI denotes the nuclei. Arrows indicate p-Smad3 positive cells. Scale bar, 20 µm. e–h KP1 inhibited TGF-β1-induced phosphorylation of ERK1/2, JNK, and p38. Western blot analyses (e) and quantitative data of p-ERK1/2 (f), p-JNK (g), and p-p38 (h) are shown. P values (from left to right): <0.001 and 0.001 (p-ERK1/2); 0.001 and 0.002 (p-JNK); 0.005 and 0.010 (p-p38). n = 3 biologically independent cells. i–k KP1 inhibited TGF-β/Smad signaling in primary tubular cells. Western blot analyses (i) and quantitative data of p-Smad2 (j) and p-Smad3 (k) in different groups are shown. P values (from left to right): 0.032 and 0.034 (p-Smad2); 0.041 and 0.049 (p-Smad3). n = 3 biologically independent cells (l–n) KP1 was more effective in inhibiting TGF-β signaling compared to TβR2 inhibitor ITD-1. NRK-49F cells were pre-incubated with KP1 (10 µg/ml, 3 μM) or ITD-1 (3 μM) for 1 h, and then treated with TGF-β1 (2 ng/ml) for 45 min. Western blot analyses (l) and quantitative data of p-Smad3 (m) and p-p38 (n) are shown. P values (from left to right): <0.001, 0.008 and <0.001 (p-Smad3); <0.001, <0.001 and <0.001 (p-p38). n = 3 biologically independent cells. Ctrl controls. Data are presented as mean values ± SEM. P values were calculated by one-way ANOVA with Fisher’s LSD post hoc test (b, c, f, g, h, m, n) or Dunnett’s T3 test (j, k). Source data are provided as a Source Data file.
We also examined the effect of KP1 on MAPK activation induced by TGF-β. As shown in Fig. 3e–h, TGF-β1 induced the phosphorylation of ERK1/2, p38 MAPK, and JNK in NRK-49F cells, whereas KP1 hampered the phosphorylation and activation of these MAPKs (Fig. 3e–h). However, KP12 did not affect Smad2/3, ERK1/2, JNK, and p38 MAPK activation induced by TGF-β1 in NRK-49F cells (Supplementary Fig. 3d–j). KP1 also blocked TGF-β1-induced Smad2/3 phosphorylation in mouse primary tubular epithelial cells (Fig. 3i–k), rat primary cardiac fibroblasts (Supplementary Fig. 4a–c), and rat primary cardiomyocytes (Supplementary Fig. 4d–f). Furthermore, KP1 was more potent than ITD-1, a small molecule inhibitor of TGF-β signaling by inducing proteasomal degradation of the TβR232,33, in blocking Smad and MAPK activation (Fig. 3l–n). Collectively, these data demonstrate that KP1 is a potent inhibitor that blocks multiple downstream signaling of TGF-β in vitro.
KP1 preserves kidney function and reduces renal fibrosis after ischemia-reperfusion injury
In view of the effectiveness of KP1 in blocking TGF-β signaling, we sought to investigate its efficacy in ameliorating renal fibrosis in vivo. To this end, we first examined the tissue distribution of KP1 in vivo after intravenous injection. At 7 days after unilateral ischemia-reperfusion injury (UIRI), FITC-labeled KP1 was injected into UIRI mice through the tail vein. After 0.5 h, major organs of mice were collected for detecting the FITC fluorescence under in vivo imaging system. As shown in Fig. 4a, FITC-labeled KP1 was largely accumulated in the injured kidney, whereas other major organs including liver, lung, heart, and spleen, as well as the contralateral uninjured kidney, exhibited little accumulation of the FITC-labeled KP1, suggesting that KP1 is preferentially delivered to the diseased kidney.
a Organ imaging showed that KP1 was preferentially accumulated in the injured kidney after UIRI. Relative levels of FITC-KP1 are shown in major organs. b Schematic diagram of the experimental design. The red line indicates the duration of surgery. The green arrowheads indicate the injections of KP1 (1 mg/kg body weight), whereas white arrowheads denote the injections of vehicle (0.01 M acetic acid). c, d KP1 preserved kidney function in UIRI mice. Serum creatinine (Scr) (c) and blood urea nitrogen (BUN) (d) levels are shown. P values (from left to right): 0.003, 0.010 (Scr); <0.001, P < 0.001 (BUN). n = 6 biologically independent animals. e Representative micrographs of Masson’s trichrome staining in different groups. Scale bar, 50 µm. Arrow indicates collagens deposition. f Quantitative determination of renal fibrotic lesions. P values (from left to right): <0.001, <0.001. n = 6 biologically independent animals. g Representative micrographs of immunohistochemical staining for fibronectin and α-SMA. Scale bar, 50 µm. h, i Quantitative data of fibronectin and α-SMA positive area in each high-power field. P values (from left to right): <0.001, <0.001 (fibronectin); <0.001, <0.001 (α-SMA). n = 6 biologically independent animals. j Representative Western blot showed renal protein levels of fibronectin, α-SMA, and collagen I in different groups. Numbers (1–3) indicate each individual animal in a given group. k–m Quantitative data of fibronectin (k), α-SMA (l), and collagen I (m) proteins in different groups. P values (from left to right): <0.001, 0.004 (fibronectin); 0.003, 0.045 (α-SMA); 0.001, 0.018 (collagen I). n = 6 biologically independent animals. *P < 0.05 versus sham; †P < 0.05 vs. UIRI alone. P values were determined by one-way ANOVA followed by Fisher’s LSD post hoc test (d, f, h, i) or Dunnett’s T3 test (c, k, l, m). Data are presented as mean values ± SEM. Source data are provided as a Source Data file.
We then employed a mouse model of UIRI and injected KP1 through daily intravenous injections, starting at 4 days after UIRI. As shown in Fig. 4b, at 10 days after UIRI, the uninjured kidney was removed through uninephrectomy (UNx) and experiments terminated at 11 days. To evaluate the therapeutic effect of KP1, we first examined renal function by measuring serum creatinine (SCr) and blood urea nitrogen (BUN). As shown in Fig. 4c, d, SCr and BUN were elevated in mice after UIRI, which were reduced after KP1 treatments. As the side effect of Klotho is hypophosphatemia and hypocalcemia34, blood phosphorus, and calcium level were examined. As shown in Supplementary Table 2, KP1 did not affect blood phosphorus and calcium level compared to the UIRI group. These results indicate that KP1 preserves kidney function after UIRI.
We next assessed the kidney fibrotic lesions in UIRI mice after treatment with KP1. As shown in Fig. 4e, f, Masson’s trichrome staining (MTS) revealed a substantial collagen deposition in the renal interstitial area at 11 days after UIRI, whereas KP1 alleviated the accumulation and deposition of interstitial collagens. Consistently, UIRI induced renal fibronectin accumulation and myofibroblast activation as assessed by immunohistochemical staining for fibronectin and α-SMA, both of which were mitigated by KP1 (Fig. 4g–i). Similar results were obtained when kidney lysates were analyzed for the expression of fibronectin, α-SMA, and collagen I proteins by Western blotting (Fig. 4j). Quantitative determination of fibronectin, α-SMA, and collagen I protein levels are presented in Fig. 4k–m, respectively. In short, these results indicate that KP1 ameliorates renal fibrotic lesions in a mouse model of IRI.
KP1 inhibits TGF-β signaling in UIRI mice in vivo
We then investigated the mechanism underlying the anti-fibrotic action of KP1. Based on the in vitro data (Fig. 2), we first examined whether KP1 affects the engagement of TGF-β and TβR2 in the diseased kidney in vivo. As shown in Fig. 5a, co-immunoprecipitation revealed a substantial binding between TGF-β1 and TβR2 at 11 days after UIRI. However, such ligand-receptor interaction virtually vanished after KP1 treatment (Fig. 5a), suggesting that, similar to in vitro situation, KP1 also disrupts the engagement of TGF-β with TβR2 in vivo.
a Co-immunoprecipitation showed that KP1 blocked the interaction between TGF-β1 and TβR2 in vivo. b, c KP1 inhibited Smad2/3 activation in vivo. Representative Western blot analyses (b) and quantitative data (c) of p-Smad3 and p-Smad2 are shown. P values: <0.001, <0.001 (p-Smad3); 0.002, 0.002 (p-Smad2). n = 6 biologically independent animals. d Representative micrographs of immunohistochemical staining of p-Smad3 and p-ERK1/2. Boxed areas are enlarged. Arrows indicate positive staining. Scale bar, 50 µm. e, f Quantitative data of p-Smad3 positive cells number (e) and p-ERK positive staining area (f) in each high-power field. P values: <0.001, <0.001 (p-Smad3); <0.001, <0.001 (p-ERK); n = 6 biologically independent animals. g KP1 blocked the activation of MAPK and TβR2 expression in vivo. h–k Quantitative data of p-ERK1/2 (h), p-JNK (i), p-p38 (j), and TβR2 (k) are shown. P values (from left to right): 0.006, 0.018 (p-ERK1/2); <0.001, 0.046 (p-JNK); < 0.001, 0.035 (p-p38); 0.002, 0.004 (TβR2). n = 6 biologically independent animals. Data are presented as mean values ± SEM. Statistical significance was determined by one-way ANOVA followed by Fisher’s LSD post hoc test (c, i, k) or Dunnett’s T3 test (e, f, h, j). Source data are provided as a Source Data file.
To further corroborate the inhibitory effects of KP1 on TGF-β signaling, we examined the downstream signaling of TGF-β. As shown in Fig. 5b, c, KP1 effectively inhibited Smad2/3 phosphorylation and activation in the diseased kidney, although it did not affect total Smad2/3 abundances after UIRI. Immunohistochemical staining exhibited that p-Smad3 was markedly induced and predominantly localized in the nuclei of tubular epithelial cells in the UIRI kidney, which was abolished by KP1 (Fig. 5d, e). Similarly, KP1 also repressed the phosphorylation and activation of ERK1/2, p38 MAPK, or JNK in the kidneys after UIRI (Fig. 5d, f, g–j). Of note, renal expression of TβR2 was also suppressed by KP1 in the injured kidney after UIRI (Fig. 5g, k).
KP1 ameliorates renal fibrosis and inflammation in obstructive nephropathy
To generalize the effect of KP1 on renal fibrosis, we sought to use another model by employing unilateral ureter obstruction (UUO), the most aggressive type of kidney fibrogenesis. As shown in Fig. 6a, mice were subjected to UUO for 7 days and KP1 was injected from day 1 to day 6. Organ distribution of FITC-labeled KP1 showed that KP1 was largely accumulated in the obstructed kidney, but not in other major organs and the contralateral unobstructed kidney (Fig. 6b). Tissue sections showed that FITC-KP1 was mainly localized in renal tubular epithelia of the obstructed kidney, but little was observed in other organs such as heart, liver, lung, and spleen (Fig. 6c). Similar results were obtained when KP1 was quantitatively assessed by liquid chromatography–tandem mass spectrometry (LC–MS/MS) analyses (Fig. 6d).
a Experimental design. The green arrowheads indicate the injections of KP1 (1 mg/kg body weight), whereas the white arrowheads indicate the injections of vehicle (0.01 M acetic acid). b The organ distribution of FITC-KP1 in vivo. Major organs as indicated were assessed for FITC-KP1 accumulation at 0.5 h after intravenous injection. c Representative micrographs of immunofluorescence of FITC-KP1 in organs. UUO mice were sacrificed 0.5 h after intravenous injection of FITC-KP1. Tissue cryosections were stained with an anti-FITC antibody. Kidney indicates UUO injured kidney. Scale bar, 50 µm. d The concentration of KP1 in each organ was assessed by LC–MS/MS at 0.5 h after intravenous injection. Two-sided t test P values (from left to right): 0.041, 0.014, 0.049, 0.049. n = 3 biologically independent animals. e Representative micrographs of Masson’s trichrome staining and immunohistochemical staining for vimentin showed that KP1 decreased interstitial matrix deposition and myofibroblasts activation. Arrows indicate positive staining. Scale bar, 50 µm. f, g Quantitative data of Masson’s trichrome staining and vimentin-positive cells number. P values: <0.001, <0.001 (Masson); <0.001, <0.001 (vimentin). n = 5 biologically independent animals. h–k Representative Western blot (h) showed renal expression of fibronectin, α-SMA, and collagen I in different groups. Quantitative data of renal fibronectin (i), α-SMA (j), and collagen I (k). P values (from left to right): 0.001, 0.001 (fibronectin); 0.033, 0.035 (α-SMA);0.026, 0.043 (collagen I). n = 5 biologically independent animals. Data are presented as mean values ± SEM. P values were determined by one-way ANOVA with Fisher’s LSD post hoc test in (f, g) or one-way ANOVA with Dunnett’s T3 test in (i–k). Source data are provided as a Source Data file.
Obstructive injury-induced substantial accumulation of collagens and increased fibroblast activation in the obstructed kidney, as demonstrated by MTS and immunohistochemical staining for vimentin (Fig. 6e–g). However, KP1 ameliorated these fibrotic lesions. Analyses of protein expressions of fibronectin, α-SMA, and collagen I by Western blotting gave rise to similar results (Fig. 6h). Quantitative determination of renal fibronectin, α-SMA, and collagen I abundances is presented in Fig. 6i–k.
We further examined the effect of KP1 on TGF-β signaling in the UUO model. As shown in Fig. 7a–c, KP-1 did not affect Smad2/3 abundances but markedly reduced the levels of p-Smad2 and p-Smad3 in the obstructed kidney, suggesting its inhibition of TGF-β canonical signaling. Immunostaining revealed that p-Smad3 was primarily localized in the nuclei of tubular epithelial cells after UUO, which was abolished by KP1 (Fig. 7d, e). Similarly, KP1 also hampered the phosphorylation and activation of ERK1/2, p38 MAPK, and JNK in the obstructed kidney (Fig. 7f–i), and reduced TβR2 expression (Fig. 7f, j). We also compared the efficacy of KP1 and ITD-1 for their ability to inhibit TGF-β signaling and renal fibrosis in vivo. As shown in Supplementary Fig. 5, KP1 was clearly more potent than ITD-1 in blocking TGF-β signaling and alleviating fibrotic lesions after UUO. KP1 was comparable with Klotho in inhibiting TGF-β signaling and renal fibrosis after UUO (Supplementary Fig. 6).
a–c KP1 inhibited activation of Smad signaling in the UUO kidney. Representative Western blot (a) and quantitative data of p-Smad3 (b) and p-Smad2 (c) in the obstructed kidney are shown. P values (from left to right): 0.003, 0.003 (p-Smad3); <0.001, 0.001 (p-Smad2). n = 5 biologically independent animals. d Immunohistochemical staining of p-Smad3 in the obstructed kidney in different groups as indicated. Arrows indicate the nuclear staining of p-Smad3. Scale bar, 50 µm. e Quantitative data of p-Smad3 positive cells number. P values (from left to right): <0.001, <0.001. n = 5 biologically independent animals. f KP1 inhibited MAPK activation and TβR2 expression in UUO. g–j Quantitative data of p-ERK1/2 (g), p-JNK (h), p-p38 (I), and TβR2 (j). P values (from left to right): 0.013, 0.022 (p-ERK); 0.020, 0.049 (p-JNK); 0.001, 0.001 (p-p38); <0.001, 0.004 (TβR2). n = 5 biologically independent animals. k, l Representative Western blot (k) and quantitative data of F4/80 (l). P values (from left to right): 0.001, 0.001. n = 5 biologically independent animals. m Immunohistochemical staining of CD3 and immunofluorescence of F4/80 are shown. Scale bar, 50 µm. n Quantitative data of CD3 positive T cells number. P values (from left to right): <0.001, 0.002. n = 5 biologically independent animals. Data are presented as mean values ± SEM. P values were calculated by one-way ANOVA with Fisher’s LSD post hoc test (c, e, j) or Dunnett’s T3 test (b, g–i, l, m). Source data are provided as a Source Data file.
We further examined the effect of KP1 on renal inflammation after UUO. As shown in Fig. 7k, l, UUO induced renal expression of F4/80, a marker of macrophages, and KP1 completely abolished its induction. Immunohistochemical staining also revealed that KP1 blocked renal infiltration of F4/80+ macrophages and CD3+ T cells in the obstructed kidneys (Fig. 7m, n).
KP1 restores endogenous Klotho expression in vivo
The kidney is the main source of endogenous Klotho in vivo. We wondered whether KP1 affects the expression of endogenous Klotho protein. To test this, we assessed the expression of Klotho protein in diseased kidneys in the absence or presence of KP1. As shown in Fig. 8a, b, Klotho was highly expressed and readily detectable by Western blotting in normal kidneys, and UIRI triggered the loss of renal Klotho expression. However, KP1 largely preserved renal expression of Klotho protein (Fig. 8a, b). Consistently, KP1 also preserved Klotho expression in the obstructed kidney after UUO (Fig. 8c, d). Immunohistochemical staining showed that Klotho was predominantly expressed in renal tubular epithelia in normal kidneys but completely lost in the obstructed kidney at 7 days after UUO, which was restored by KP1 (Fig. 8e, f). These data suggest that KP1 may elicit its protective activity at least partially through restoring endogenous Klotho expression.
a, b KP1 restored endogenous Klotho expression after UIRI. Representative Western blot (a) and quantitative data of Klotho (b) are shown. P value (from left to right): 0.007, 0.037 (one-way ANOVA with Fisher’s LSD test). n = 6 biologically independent animals. Data are presented as mean values ± SEM. c, d KP1 preserved endogenous Klotho protein after UUO. Representative Western blot (c) and quantitative data of Klotho (d) are shown. P value (from left to right): <0.001, <0.001 (one-way ANOVA with Fisher’s LSD test). n = 5 biologically independent animals). Data are presented as mean values ± SEM. e Immunohistochemical staining for Klotho protein in the obstructed kidney at 7 days after UUO. Arrows indicate positive staining. Scale bar, 50 µm. f Quantitative data of Klotho positive staining area in each high-power field. P value (from left to right): 0.001, 0.003. n = 5 biologically independent animals. Data are presented as mean values ± SEM. g, h Colocalization of Klotho and p-Smad3. Representative micrographs show Klotho localized in those tubules with nuclei negative for expression of p-Smad3. Nontumor kidney tissue from the patients who had renal cell carcinoma and underwent nephrectomy was used as normal controls (g). Sequential paraffin-embedded kidney sections from patients with membranous nephropathy (MN) were immunostained for Klotho and p-Smad3. Scale bar, 50 μm. Boxed areas are enlarged. h Scatter plots with linear regression show an inverse correlation between Klotho expression levels and p-Smad3 positive cells in human kidney sections. The Spearman correlation coefficient (r) and two side P value are shown. Source data are provided as a Source Data file.
To establish the clinical relevance of Klotho to TGF-β signaling in human CKD, we performed immunostaining for Klotho and p-Smad3 on serial sections of kidney specimens. As shown in Fig. 8g, abundant Klotho expression was evident in the tubular epithelium of normal human kidneys, whereas no or little p-Smad3+ cells were observed. However, Klotho expression was inhibited and Smad3 signaling was activated in human kidney biopsies from CKD patients with membranous nephropathy (MN). In tubules with high Klotho expression, a few p-Smad3+ cells were observed (Fig. 8g, asterisk), while in the adjacent tubules with low levels of Klotho, p-Smad3+ nuclei were abundant (Fig. 8g, arrowhead). There was clearly an inverse correlation between Klotho level and Smad3 activation in diseased kidneys (Fig. 8h), suggesting an intrinsic association of Klotho with TGF-β signaling in humans as well.
