Pharmacological activation of autophagy by rapamycin attenuates UUO-induced tubulointerstitial fibrosis (TIF)

Rapamycin, the best characterized pharmacological inducer of autophagy, acts by inhibiting the mammalian target of rapamycin (serine/threonine kinase) complex 1 (MTORC1).¹³ To determine whether autophagy is associated with the evolution of kidney fibrosis, C57BL/6 mice were treated with daily rapamycin (1 mg/kg) or vehicle, starting 1 d prior to UUO operation. Compared with vehicle-treated control mice, rapamycin administration significantly increased the level of LC3-II and concomitant degradation of SQSTM1/p62, a receptor and selective substrate of autophagy,¹⁴ in the obstructed kidneys at d 7 after UUO (Fig. 2A). Notably, activation of autophagy by rapamycin markedly attenuated UUO-induced expression of COL1 (Fig. 2A and B). Consistent with the immunoblotting data, co-immunofluorescence with SQSTM1/p62 (red) and lotus tetragonolobus lectin (LTL, green), a marker of proximal tubules, revealed that abundant SQSTM1/p62-positive aggregates were located predominantly in the tubular cells of vehicle-treated mice following UUO, whereas the accumulation of SQSTM1/p62 was alleviated in mice treated with rapamycin (Fig. 2C). Accordingly, histological examination showed the degree of tubular injury (Fig. 2D) and the severity of interstitial fibrosis (Fig. 2E) were markedly reduced by rapamycin administration. In sham-operated kidneys, there was no statistical difference in morphological characterization between vehicle- and rapamycin-treated mice. Taken together, pharmacological induction of autophagy using rapamycin significantly ameliorated UUO-induced TIF, indicating renoprotective roles of autophagy in the pathogenesis of renal fibrosis.

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Figure 2.

Enhancement of autophagy by rapamycin (Rapa) ameliorates renal interstitial fibrosis in C57BL/6 mice. (A) Immunoblot analyses of LC3, SQSTM1/p62, and COL1 in the obstructed kidneys in different groups as indicated. (B) Quantitative determination of the relative abundance of the indicated proteins among different groups. Data are means ± SEM (n = 6); *, P < 0.001 vs. sham; #, P < 0.05 vs. UUO mice treated with vehicle. (C) Coimmunostaining of LTL (green) and SQSTM1/p62 (red) in kidney sections. Scale bar: 20 μm. (D) Semiquantitative analysis of tubular damage in the obstructed kidneys. Data are means ±SEM (n = 6); *, P < 0.05 vs. sham; #, P < 0.05 vs. UUO mice treated with vehicle. (E) Representative micrographs from indicated groups with either periodic acid-Schiff (PAS; upper panels) or Masson's trichrome staining (lower panels). Scale bar: 20 μm.

Proximal tubular epithelial cell-specific Atg5 deletion aggravates TIF

A series of autophagy-related genes are involved in the process of autophagy, and the ATG5 protein is required for the initiation of autophagosome formation. To further investigate the effects of autophagy on TIF, we generated mice with selective Atg5 deletion (atg5^−/−) in proximal tubular epithelial cells (TECs) by crossing Atg5^flox/flox mice with transgenic mice expressing Cre recombinase under the control of the Kap (kidney androgen regulated protein) promoter (Kap-cre), as previously described.¹⁵ We first performed immunoblot analysis to confirm a significantly decreased ATG12–ATG5 conjugation expression with reduced LC3-II accumulation and SQSTM1/p62 degradation in the kidney cortex of atg5^−/− mice, as compared with their wild-type (Atg5^+/+) littermates (Fig. 3A). Kidney fibrosis was induced by UUO in 8-wk-old male mice. As shown in Figure 3B and C, kidney tissue of wild-type mice showed a significant increase in protein level of LC3-II at d 7 after UUO, whereas this response was markedly abrogated in the atg5^−/− mice. Consistently, a substantial reduction of SQSTM1/p62 degradation was observed in fibrotic kidneys of atg5^−/− animals. The impaired autophagy in proximal tubular epithelial cells was concurrently accompanied by increased COL1 expression. Likewise, the amount and size of cytoplasmic SQSTM1/p62-positive aggregates were significantly greater in proximal tubular cells of atg5^−/− mice than those of Atg5^+/+ mice, as evaluated by immunofluorescence staining of renal tissue after UUO (Fig. 3D). Histological examination further confirmed dramatically elevated interstitial fibrosis (Fig. 3E) and collagen deposition (Fig. 3F), with more severe tubular injury (Fig. 3G) in the obstructed kidneys of atg5^−/− mice in contrast to Atg5^+/+ mice. These pathological changes were observed throughout the whole cortex, although the degree of TIF severity was not distributed evenly. Collectively, these findings indicate that aberrant activation of autophagy in proximal tubular cells is critical for promoting kidney fibrosis.

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Figure 3.

Proximal tubular epithelial cell-specific Atg5 deletion exacerbates UUO-induced renal fibrosis. (A) Immunoblot analyses of ATG12–ATG5 conjugation, LC3 and SQSTM1/p62 in the kidneys of Atg5^+/+ and atg5^−/− mice. (B) Immunoblot analyses of proteins in the kidneys from sham-operated and at d 7 after UUO in Atg5^+/+ and atg5^−/− mice. (C) Relative expression levels of the indicated proteins normalized to ACTB in Atg5^+/+ and atg5^−/− mice. Data are means ± SEM (n = 6); *, P < 0.001 vs. sham; #, P < 0.05 vs. Atg5^+/+ mice with UUO. (D) Coimmunostaining of LTL (green) and SQSTM1/p62 (red) on kidney sections. Scale bar: 20 μm. (E) Representative micrographs of kidney histology with Masson's trichrome staining from Atg5^+/+ and atg5^−/− mice. Scale bar: 80 (upper panels), 40 (middle panels), and 20 (lower panels) μm. (F) Representative images of Sirius red-stained sections of sham-operated and 7 d-obstructed Atg5^+/+ and atg5^−/− mice. Scale bar: 20 μm. (G) Semiquantitative analysis of tubular damage in the obstructed kidneys. Data are means ± SEM (n = 6); *, P < 0.05 vs. sham; #, P < 0.05 vs. Atg5^+/+ mice with UUO.

Atg5 deficiency abolishes the protective effect of rapamycin against renal fibrosis

Next, we sought to determine whether UUO-induced renal fibrosis in Atg5-deficient mice was rescued by rapamycin. As shown in Figure 4A and B, in the sham-operated kidneys, treatment of Atg5^+/+ mice with rapamycin caused a significant increase in the expression levels of ATG12–ATG5 conjugation compared with those in vehicle-treated Atg5^+/+ mice. Rapamycin-treated atg5^−/− mice demonstrated a very slight increase of endogenous ATG12–ATG5 conjugation and a decrease in SQSTM1/p62 as compared to those in vehicle-treated atg5^−/− mice, but the difference was not statistically significant. Administration of rapamycin resulted in dramatic differences in protein levels of LC3-II, SQSTM1/p62, and ATG12–ATG5 conjugation between Atg5^+/+ and atg5^−/− mice. As expected, in obstructed kidneys at d 7, administration of rapamycin significantly upregulated autophagy and reduced COL1 expression when compared with those in vehicle-treated Atg5^+/+ mice. Whereas, the protective effects of rapamycin were not significant in atg5^−/− mice (Fig. 4C and D). Similarly, obstructed kidneys from Atg5^+/+ mice demonstrated less TIF in rapamycin-treated versus vehicle-treated mice, as indicated by a positive blue color in Masson's trichrome-stained sections. In contrast, these changes were not apparent in the obstructed kidneys from atg5^−/− mice treated with or without rapamycin (Fig. 4E and F). Our results suggest that the protective effect of rapamycin against renal fibrosis appear to be at least in part mediated through enhanced autophagy via ATG5.

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Figure 4.

Atg5 deficiency abolishes the protective effect of rapamycin against renal fibrosis in mice. (A) Immunoblot analyses of proteins in the sham-operated kidneys of Atg5^+/+ and atg5^−/− mice treated with or without rapamycin. (B) Relative expression levels of the indicated proteins normalized to ACTB in Atg5^+/+ and atg5^−/− mice treated with or without rapamycin. Data are means ± SEM (n = 6); *, P < 0.001 vs. vehicle-treated Atg5^+/+ mice; #, P < 0.05 vs. rapamycin-treated Atg5^+/+ mice. (C) Immunoblot analyses of proteins in the kidneys at day 7 after UUO in Atg5^+/+ and atg5^−/− mice treated with or without rapamycin. (D) Relative expression levels of the indicated proteins normalized to ACTB in Atg5^+/+ and atg5^−/- mice treated with or without rapamycin. Data are means ± SEM (n = 6); *, P < 0.001 vs. vehicle-treated Atg5^+/+ mice; #, P < 0.05 vs. rapamycin-treated Atg5^+/+ mice. (E) Representative micrographs of kidney histology with Masson's trichrome staining from Atg5^+/+ and atg5^−/− mice. Scale bar: 80 (upper panels), 40 (middle panels), and 20 (lower panels) μm.

Autophagy deficiency associates with enhanced tubular cell G₂/M arrest in the fibrotic kidney

Because autophagy is involved in cell cycle progression in several cancer types,¹⁶ the relationship among autophagy, the cell cycle and ECM deposition was examined in a UUO-induced fibrosis model. To this end, we first analyzed cells in proliferation and arrested at the G₂/M phase of the cell cycle by costaining MKI67 with phosphorylation of histone H3 at Ser10 (p-H3) to evaluate the cell cycle distribution. Compared with sham-operated C57BL/6 mice, the percentage of TECs in the G₂/M phase (p-H3 positive) among all proliferative TECs (MKI67 positive) elevated to 68.7 ± 10.1 on d 7 and was even increased to 78.7 ± 4.0 on d 14 after UUO (Fig. 5A and B). The UUO-induced time-dependent G₂/M phase arrest in TECs was virtually identical to those observed for ECM accumulation, indicating that G₂/M arrest correlates with kidney fibrogenesis.

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Figure 5.

Tubular cell G₂/M arrest is increased in UUO-induced fibrotic kidney. (A) Dual immunofluorescence of MKI67 (red) and p-H3 (green) in sham-operated and UUO kidneys on d 7 and 14 in C57BL/6 mice. DAPI (blue) was used for nuclear staining. Scale bar: 20 μm. (B) The percentage of MKI67⁺ p-H3⁺ cells among total MKI67⁺ tubular cells in different groups. Data represented as mean ± SEM (n = 6); *, P < 0.05 vs. sham. #, P < 0.05 vs. UUO 7 d. (C) Representative immunofluorescence staining of p-H3 (red), COL1 (green), and DAPI (blue) on kidney sections of Atg5-deficient mice from different groups. Scale bar: 20 μm. (D) Quantification of tubular cells with p-H3 positive. Data represented as mean ± SEM (n = 6); *, P < 0.05 vs. sham; #, P < 0.01 vs. obstructed kidney of Atg5^+/+ mice.

Next, we attempted to verify our findings utilizing renal tubular epithelial cell-specific atg5 knockout mice following UUO. As shown in Figure 4C, very few TECs labeled positively with p-H3 antibody in both sham-operated atg5^−/− and Atg5^+/+ mice. At 7 d following UUO, increased p-H3-positive cells with more COL1 accumulation were evident in Atg5^+/+ animals, whereas the responses were substantially aggravated in the atg5^−/− mice (Fig. 5C and D). Therefore, ATG5 has a negative influence on cell cycle progression, and inhibition of autophagy may facilitate kidney fibrosis through a G₂/M arrest-dependent mechanism in TECs.

Autophagy blocks AGT II-induced COL1 expression in vitro

To ascertain the role of autophagy in ECM accumulation, the renal epithelial cell line HK-2 was treated with AGT II, an important mediator of tubulointerstitial fibrosis. As shown in Figure 6A and B, a time-course experiment in which HK-2 cells were treated with AGT II (10⁻⁶ mol/L) for various durations (0–48 h) resulted in increased levels of LC3-II starting from 6 h, reaching the highest levels at 24 h and reducing to baseline at 48 h. Whereas, the expression of COL1 was significantly upregulated after 48 h of treatment. These data are in agreement with in vivo findings that renal epithelial cells make attempts to trigger autophagy, followed by COL1 expression after 24 h. Additionally, pre-incubation with the lysosomal protease inhibitors E64d and pepstatin A (E/P), which block the lysosome-dependent autophagic degradation, appeared to increase LC3-II level in cells untreated with AGT II, and further to a greater content in cells exposed to AGT II at 24 and 48 h (Fig. 6C and D). Consistently, pretreatment with E/P increased the accumulation of LC3 puncta by immunofluorescence staining in AGT II-treated cells (Fig. 6E). Our data suggest AGT II stimulation causes a significant autophagy flux.

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Figure 6.

Overexpression of ATG5 augments AGT II-mediated autophagy in vitro. (A) Serum-deprived HK-2 cells were treated with 10⁻⁶ mol/L of AGT II for the indicated time period. Cell lysates were probed with antibodies against LC3 and COL1. ACTB was used as a loading control. (B) Densitometry of LC3-II and COL1 proteins in immunoblots (relative to ACTB). Data are means ± SEM (n = 3); *, P <0.001 vs. control group. #, P <0.001 vs. AGT II-treated cells for 12 or 24 h. (C) Cells were treated with AGT II at 10⁻⁶ mol/L for 24 and 48 h in the presence or absence of E64d and pepstatin A (E/P). Induction of autophagy was assessed by analyzing the expression of LC3. ACTB was used as a loading control. (D) Densitometry of LC3-II in immunoblots (relative to ACTB). Data are means ± SEM (n = 3); *, P < 0.001 vs. control group; #, P < 0.05 vs. AGT II-treated cells at the same time point in the absence of E/P. (E) Cells were treated with AGT II (10⁻⁶ mol/L) for 24 and 48 h in the absence or presence of E/P. Formation of LC3 puncta was analyzed by immunofluorescence under confocal microscopy (×400). (F) Cells were transfected with either pcDNA3.1-ATG5 or pcDNA3.1-HA vector followed by incubation with 10⁻⁶ mol/L of AGT II for 24 h. LC3 and ATG12–ATG5 conjugation protein levels were examined by Western blotting. ACTB was used as a loading control. (G) LC3 and ATG5 contents were quantitatively analyzed using a densitometer. Values are mean ± SEM (n = 3); *, P < 0.001 vs. control group in the same condition; #, P < 0.05 vs. AGT II-treated cells in the corresponding condition without E/P; §, P < 0.05 vs. cells transfected with empty vector in the corresponding condition.

Given that atg5^−/− mice showed an increase in COL1 accumulation and interstitial fibrosis (Fig. 3) compared with wild-type mice, we sought to determine the role of ATG5 in AGT II-induced COL1 production in vitro. To this end, we transfected HK-2 cells with either ATG5 expression or control vectors. As shown in Figure 6F and G, enhanced expression of ATG5 alone induced a small increase in basal LC3-II levels, and this response became pronounced in the presence of E/P. Therefore, elevated LC3-II in ATG5-overexpressing cells does not appear to be a result of inhibition of autophagic flux. ATG5 overexpression markedly increased LC3-II content upon AGT II stimulation for 24 h both in the presence and absence of protease inhibitors, compared with the control vector. In contrast, the protein level of COL1 was dramatically reduced after 48 h of AGT II exposure in ATG5-overexpressing cells (Fig. 7A and B).

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Figure 7.

ATG5 inhibits AGT II-induced COL1 expression in vitro. (A) HK-2 cells were transfected with either pcDNA3.1-ATG5 or pcDNA3.1-HA vector before stimulation with 10⁻⁶ mol/L of AGT II for 24 or 48 h. Cell lysates were analyzed by immunoblotting with the indicated antibodies. (B) Densitometry of LC3-II and COL1 proteins in immunoblots. Data are means ± SEM (n = 3); *, P < 0.05 vs. negative and empty control; #, P < 0.05 vs. control and AGT II-treated cells for 24 h with or without ATG5 overexpression; §, P < 0.05 vs. control and AGT II-treated cells for 48 h without ATG5 overexpression. (C) Expression of LC3, ATG12–ATG5 conjugation, SQSTM1/p62, and LRP2/MEGALIN in primary TECs from Atg5^+/+ and atg5^−/− mice were assessed by Western blot analysis. (D) Primary TECs from Atg5^+/+ and atg5^−/− mice were stimulated with 10⁻⁶ mol/L of AGT II for 24 or 48 h. Cell lysates were probed with antibodies against the indicated proteins.

Further, we characterized the effect of ATG5 specifically on COL1 expression by using primary renal proximal TECs isolated from 3-wk-old Atg5^+/+ or atg5^−/− mice. As shown in Fig. 7C, the expression of ATG12–ATG5 conjugation and LC3-II were almost completely suppressed, and SQSTM1/p62 accumulated in primary atg5^−/− TECs. As expected, the protein levels of LC3-II and COL1 in primary TECs from Atg5^+/+ mice were similar in pattern to those in HK-2 cells after AGT II treatment for 24 and 48 h. Remarkably, AGT II-stimulated COL1 production was exacerbated in primary atg5^−/− TECs (Fig. 7D). Taken together, these data suggest that Atg5-deficient TECs are more sensitive to induction of COL1 expression in response to AGT II.

ATG5 prevents ECM deposition through blocking G₂/M arrest

As our observations indicated that AGT II induces autophagy and COL1 production at 2 distinct time points, we sought to determine whether these appearances of 2 biological events were due to the heterogeneity of cellular populations. As shown in Fig. 8A and B, stimulation of HK-2 cells with AGT II for 24 h did not alter cell cycle progression. However, AGT II treatment at 48 h caused a striking increase in the percentage of cells in G₂/M phase by 50% as compared to the cells untreated or treated for 24 h. Moreover, we synchronized HK-2 cells at G₁/S or G₂/M phase by incubation with hydroxyurea or nocodazole, respectively.¹⁷ Immunoblot analysis showed that the expression of LC3-II in G₁/S-phase-synchronized cells was higher than that in G₂/M-phase-synchronized cells after exposure to AGT II for 24 and 48 h. Interestingly, synchronization of cells at the G₁/S phase almost completely blocked AGT II-induced COL1 production. In contrast, a dramatic increase of COL1 expression was observed in cells synchronized at the G₂/M phase after 48 h of treatment with AGT II (Fig. 8C and D). These data suggest that the cell cycle state determines cell fate propensity, and that AGT II induces autophagy preferentially in the G₁/S phase.

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Figure 8.

AGT II-induced autophagy is dependent on cell cycle status in HK-2 cells. Cells were treated with10⁻⁶ mol/L of AGT II for the indicated time period. (A) Cells were exposed to AGT II, followed by PI staining and flow cytometry. (B) Cell cycle distribution among different groups. Data are means ± SEM (n = 3); *, P < 0.05 vs. control and AGT II-treated cells for 24 h. Symbol indicates the fraction of G₂/M phase cells. (C) Cells synchronized at G₁/S or G₂/M phase were treated with AGT II for the indicated time period. Cell lysates were probed with antibodies against LC3 and COL1. (D) Densitometry of LC3-II and COL1 proteins in immunoblots. Data are means ± SEM (n = 3); *, P < 0.05 vs. respective control group; #, P < 0.05 vs. control and cells synchronized at G₁/S phase treated with AGT II for 24 h; §, P < 0.05 vs. cells synchronized at G₂/M phase treated with AGT II for 0–24 h, or cells synchronized at G₁/S phase treated with or without AGT II.

Next, we investigated the functional role of ATG5 in cell cycle progression. Compared with empty vector, ATG5 overexpression significantly reduced the proportion of cells in G₂/M arrest (Fig. 9A), and was accompanied by a decrease in the levels of CDKN1A/p21, phosphorylated (p)-CDK1 and CCNB1 (cyclin B1) after treatment with AGT II for 48 h (Fig. 9B and C). In the absence of AGT II, neither empty vector nor ATG5 plasmid alone had effects on the distribution of the cell cycle and expression of cell cycle regulatory proteins in HK-2 cells (Fig. 9A–C). Further, treatment of primary atg5^−/− TECs with AGT II for 48 h greatly led to an accumulation of cells in the G₂/M phase with a concomitant reduction of cells in the G₀/G₁ phase in contrast to primary Atg5^+/+ TECs. Quantitative analysis revealed a 20% increase of the cell population in the G₂/M phase in atg5^−/− TECs compared to that in wild-type TECs (Fig. 9D). The changes in expression of regulatory proteins were consistent with the cell cycle distribution (Fig. 9E and F). Taken together, our results indicate that ATG5 may attenuate COL1 production primarily by blocking G₂/M cell cycle arrest.

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Figure 9.

Overexpression of ATG5 attenuates AGT II-induced G₂/M cell cycle arrest in HK-2 cells and primary TECs. (A) HK-2 cells were transiently transfected with either pcDNA3.1-ATG5 or pcDNA3.1-HA vector followed by treatment with 10⁻⁶ mol/L of AGT II for 48 h, and then cell cycle distribution was analyzed by flow cytometry. Data are means ± SEM (n = 3); *, P < 0.01 vs. AGT II-untreated cells; #, P < 0.05 vs. AGT II-treated cells without ATG5 overexpression. Symbols indicate the fraction of G₂/M phase cells. (B) HK-2 cells were treated as described above. Expression of CDKN1A/p21, p-CDK1, and CCNB1 (cyclin B1) was examined by immunoblot. (C) Protein levels were quantified with scanning densitometry and standardized to ACTB. Data are expressed as mean ± SEM (n = 3); *, P < 0.01 vs. control or cells treated with AGT II for 24 h; #, P <0.05 vs. cells treated with AGT II for 48 h without ATG5 overexpression. (D) Primary renal proximal TECs from Atg5^+/+ and atg5^−/− mice were stimulated with 10⁻⁶ mol/L of AGT II for 48 h. Cell cycle distribution was assessed by flow cytometry. Data are expressed as mean ± SEM (n = 3); *, P < 0.01 vs. respective control; #, P < 0.05 vs. AGT II-treated cells from Atg5^+/+ mice. Symbols indicate the fraction of G₂/M phase cells. (E) Primary renal proximal TECs were treated as described in Figure 5D. Protein expression was detected by immunoblotting. (F) The expression of CDKN1A/p21, p-CDK1, and CCNB1 was quantitatively analyzed using a densitometer. Values are mean ± SEM (n = 3); *, P < 0.05 vs. respective control or cells treated with AGT II for 24 h; #, P < 0.05 vs. cells from Atg5^+/+ mice treated with AGT II for 48 h.

ATG5-mediated inhibition of G₂/M arrest is autophagy-dependent

Aside from its function as a critical regulator of autophagy induction, ATG5 has been associated with proliferation and apoptosis under stress conditions.^18-20 These previous findings encouraged us to investigate whether autophagy induction is required for ATG5-mediated cell cycle progression. To this end, HK-2 cells were transfected with a plasmid expressing a mutant form of ATG5 (ATG5^K130R), which is unable to conjugate with ATG12 and loses autophagic activity.²¹ Indeed, we found that overexpression of ATG5^K130R did not enhance formation of the ATG12–ATG5 conjugate with or without AGT II exposure, and had no influence on AGT II-induced autophagy, as assessed by LC3-II immunoblotting (Fig. 10A and B). Interestingly, when compared with empty vector, overexpression of ATG5 reversed the percentage of G₂/M-arrested cells following AGT II treatment for 48 h, whereas ATG5^K130R expression failed to show such inhibitory effects. On the contrary, inhibition of autophagy by pharmacological inhibitor, 3-methyladenine (3-MA), or specific knockdown of ATG5 with its siRNA, significantly increased G₂/M arrest under the same conditions (Fig. 10C and D). In the absence of AGT II, empty vector, ATG5, ATG5-K130R, ATG5 siRNA, or 3-MA did not affect cell cycle progression. Collectively, these data indicate that blockage of G₂/M arrest by ATG5 is dependent on its function in autophagy.

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Figure 10.

Autophagy induction is required for ATG5-mediated cell cycle progression. (A) HK-2 cells were transiently transfected with plasmids encoding pcDNA3.1-ATG5-K130R or pcDNA3.1-HA followed by incubation with 10⁻⁶ mol/L of AGT II for 24 or 48 h. Expression of LC3, SQSTM1/p62, and ATG5 (ATG12-conjugated and monomeric) were examined by immunoblot. (B) Relative expression levels of the indicated proteins normalized to ACTB. Data are mean ± SEM (n = 3); *, P < 0.05 vs. negative control, empty vector, or ATG5-K130R control vector. (C) HK-2 cells were transiently transfected with plasmids encoding pcDNA3.1-ATG5, pcDNA3.1-ATG5-K130R or pcDNA3.1-HA, or scramble siRNA, ATG5 siRNA, or cells pretreated with 3-MA, and then cell cycle distribution was analyzed in the absence (upper panels) or presence (lower panels) of AGT II for 48 h. (D) HK-2 cells were treated as indicated and the histograms represent the distribution of cells cycle among different groups. Data are means ± SEM (n = 3); *, P < 0.05 vs. cells in the absence of AGT II; #, P < 0.05 vs. cells treated with AGT II without ATG5 overexpression. Symbols indicate the fraction of G₂/M phase cells.

Animals

Wild-type C57BL/6 mice were obtained from the Sun Yat-Sen University Animal Center (Guangzhou, China). GFP-Lc3 transgenic mice and Atg5^flox/flox mice were purchased from the RIKEN BioResource Center (Tsukuba, Ibaraki, Japan). The Kap-cre mice expressing Cre recombinase under the control of the Kap promoter were from the Jackson Laboratory (Bar Harbor, Maine, USA). All animal studies were approved by the Animal Care and Use Committee of the Sun Yat-Sen University.

Proximal tubule-specific atg5^−/− mice were generated by crossing an Atg5^flox/flox mouse with a Kap-cre mouse.¹⁵ All mice were maintained by backcross breeding to C57BL/6 mice. Genotyping was identified by polymerase chain reaction (PCR) analysis using transgene-specific primers, as described previously.^33-34 Male mice between 8 and 10 wk of age were used in the present study.

For the UUO model, mice underwent ligation of the left ureter with silk sutures, and sham-operated mice were used as controls.³⁵ To activate autophagy, rapamycin (Sigma, R0395) was dissolved in vehicle (0.01% [v/v] dimethyl sulfoxide (Sigma, D2650]) followed by daily intraperitoneal injection in C57BL/6, atg5^−/−, or Atg5^+/+ mice (1 mg/kg/d) 1 d before and until UUO or sham surgery. Control mice received the same dose of dimethyl sulfoxide. Mice (n = 6) were sacrificed at 3, 7, and 14 d after UUO. Both contralateral and obstructed kidneys were harvested for further studies.

Kidney histology analyses

The kidney tissues from C57BL/6, Atg5^+/+ and atg5^−/− mice were fixed with 4% paraformaldehyde (Sigma, P6148). Paraffin-embedded 3-μm kidney sections were stained with periodic acid-Schiff (Sigma, 395B-1KT), Masson's trichrome (Baso, BA4079A) or picrosirius red (Polysciences, 24901). The degree of tubular injury was assessed semiquantitatively by the point-counting method in a blinded fashion.³⁵ Samples from GFP-Lc3 transgenic mice were analyzed as described.³³

Briefly, mice were transcardially perfused with 4% paraformaldehyde in 0.1 M sodium phosphate buffer (pH 7.4). Kidney tissues were further fixed in the same buffer for an additional 4 h, immersed in 15% sucrose (Sigma, S0389) for 4 h, followed by 30% immersion in sucrose at 4°C overnight for cryoprotection. The sections (5–7 μm) were embedded and frozen with optimal cutting temperature (OCT) compound (Tissue-Tek, 4583) before being sectioned with a cryostat, air-dried, and stored at −20°C. Frozen tissue sections were mounted on a glass slide and viewed under a fluorescence microscope.

Immunofluorescence staining of the kidney was performed using an established procedure.³⁵ Briefly, the 4-μm paraffin-embedded tissue sections were deparaffinized, rehydrated and incubated with the indicated primary antibodies at 4°C overnight, followed by their corresponding secondary antibodies for 1 h at room temperature.

For immunofluorescent analysis, cells were seeded onto glass coverslips, exposed to different experimental conditions, and fixed in 4% phosphate-buffered saline (PBS; Boster, PYG0021)-buffered paraformaldehyde. Formation of autophagic vesicles was monitored by LC3 puncta formation in cells. DAPI (Sigma-Aldrich, D9542) was used to stain the nuclei. All images were collected by laser scanning confocal microscopy (Zeiss LSM 510 META; Carl Zeiss, Oberkochen, Germany).

Western blotting analysis

Kidney cortex or cultured cells were homogenized in lysis buffer (Sigma, R0278) containing Complete Mini Protease Inhibitor Cocktail (Roche Applied Science, 05892791001). Protein concentration was measured with the Bradford assay. Samples were loaded and separated by 10–15% SDS polyacrylamide gels, electrotransferred onto a nitrocellulose membrane, blotted with the indicated antibodies, and then detected by enhanced chemiluminescence (Amersham Pharmacia Biotech, RPN2232).³⁵ Densitometric analysis was performed with an image analysis program (Fluorchem^TM 8900, Alpha Innotech).

Cell culture and treatments

A human proximal tubule epithelial cell line (HK-2) was purchased from American Type Culture Collection (ATCC®CRL-2190^TM) and cultured at 37°C, 5% CO₂ in DMEM/F-12 medium (GIBCO, 11330–057) supplemented with 10% fetal bovine serum (GIBCO, 10099) and antibiotics.

Mouse TECs were isolated and cultured as described previously.¹⁵ Briefly, kidneys were harvested from 3-wk-old male Atg5^+/+ or atg5^−/− mice under sterile conditions. The kidney cortex was diced into small pieces (<1 mm³), incubated in DMEM medium containing collagenase (Sigma, C1639) at 37°C for 20 min, and sieved through 2 nylon sieves (pore size 250 μm and 70 μm). The filtrated cells were resuspended cultured at 37°C, 5% CO₂ in DMEM/F-12 medium supplemented with 10% fetal bovine serum and antibiotics.

HK-2 and primary tubular epithelial cells between passages 3 to 5 were grown to approximately 80% confluence, followed by serum deprivation for 24 h prior to experimental manipulation. To induce autophagy, cells were treated with human AGT II (10⁻⁶ mol/L) for different time points. In some experiments, cells were pre-treated with 0.5 mmol/L hydroxyurea for 20 h, 0.25 ug/ml nocodazole for 20 h or 10 µg/ml E64d-pepstatin A for 60 min, and then stimulated with AGT II.

Plasmid constructs and transfection

The full-length ATG5 gene was amplified from genomic DNA of the HK-2 cell line by PCR and subcloned into the pcDNA3.1 vector (Invitrogen,V79020). The ATG5-K130R mutant replacing Lys130 with Arg was generated by PCR and cloned into the Xhol and Xbal sites of pcDNA3.1-HA.³⁶ The primers used for PCR were ATG5: sense 5′-GCGTCGACTCCTGGAAGAATGACAGAT-3′, antisense 5′-CGCCTCGAGTCCTTCAATCTGTTGGCT-3′ and ATG5-K130R mutant: sense 5′-CATGTATGAGAGAAGCTGATGCT-3′, antisense 3′-AGCATCAGCTTCTCTCATACATGA-5′. pcDNA3.1-HA mammalian expression vector was from Invitrogen Life Technologies (V79020). Scrambled and ATG5-specific siRNA products were from Shanghai GenePharma Co., Ltd (Shanghai, China). ATG5 siRNA knockdown was performed by transient transfection with validated ATG5 siRNA (sense, GCAACUCUGGAUGGGAUUGTT; antisense, CAAUCCCAUCCAGAGUUGCTT). Scrambled siRNA was used as control. Cells were transfected with ATG5, ATG5-K130R mutant, or ATG5 siRNA using Lipofectamine™ LTX and Plus reagent in Opti-mem (Invitrogen, 15338–100) according to the manufacturer's instructions. Overexpression efficacy was assessed by western blotting.

Cell cycle distribution analysis

Propidium iodide (PI; Sigma, P4170) staining was used to analyze DNA content and cell cycle profile. After treatment, cells were washed twice with ice-cold PBS, harvested, and fixed with 70% ethanol at 4°C overnight. Cells were resuspended in PBS, permeabilized with 0.125% Triton X-100 (Sigma-Aldrich, T9284) and stained with 40 μg/ml PtdIns for 30 min on ice. Cell cycle distributions were analyzed by flow cytometry (FACS Calibur, BD Biosciences).