mTORC1 activation in podocytes causes mesangial expansion.

To address whether activation of mTORC1 in podocytes contributes to podocyte injury and the development of glomerulopathy, we generated mice (PcKOTsc1) with a conditionally ablated Tsc1 gene in podocytes using a Cre expression cassette under the control of the Neph2 promoter (23). Because TSC1 is an upstream negative regulator of mTORC1, loss of TSC1 should result in activation of the mTORC1 pathway (24). Conventional Tsc1-KO mice die in embryonic stages due to cardiac and liver dysfunction (16), but PcKOTsc1 mice were born at normal Mendelian ratios and grew normally (Supplemental Figure 2A). We monitored mTORC1 activity over time by assaying the level of phospho-S6. Specific enhancement of phospho-S6 in podocytes was observed in PcKOTsc1 mice around 2 weeks after birth, suggesting that Cre recombination of the floxed Tsc1 gene was completed soon after birth (Supplemental Figure 2B). At 4 weeks of age, phosphorylation of S6 in the majority of PcKOTsc1 podocytes was dramatically enhanced, and rapamycin treatment attenuated this as expected, indicative of a high mTORC1 activity in podocytes of PcKOTsc1 mice (Figure ​(Figure2A).2A). Western blot analysis using glomerular proteins also showed that S6 was highly phosphorylated in a rapamycin-sensitive manner, whereas the levels of phosphorylation of Akt (Ser473), the site regulated by mTORC2 activity, was stable in PcKOTsc1 glomeruli (Figure ​(Figure2B).2B). PcKOTsc1 podocytes stained with WT1, a podocyte marker, were larger than those of wild-type mice (Figure ​(Figure2A2A and Supplemental Figure 2C). We found that large amounts of eosin- and PAS-positive materials were present in the mesangial area (Figure ​(Figure2A2A and Supplemental Figure 2, D and E) of PcKOTsc1 mice. Furthermore, total glomerular cell number was decreased in PcKOTsc1 mice (Supplemental Figure 2F), indicating that the glomerular phenotype seen in PcKOTsc1 mice was not associated with cell proliferation. Again, these phenotypic changes were not observed in rapamycin-treated PcKOTsc1 glomeruli (Figure ​(Figure2A2A and Supplemental Figure 2, C–F), suggesting that high mTORC1 activity in PcKOTsc1 podocytes contributed to mesangial expansion in this model. Consistent with these observations, both mRNA and protein levels of type IV collagen and fibronectin were significantly increased in PcKOTsc1 glomeruli (Figure ​(Figure2,2, C and D). Moreover, we also observed that mRNA and protein levels of TGF-β1 were significantly enhanced (Supplemental Figure 2, G and H), whereas the protein expression but not amount of mRNA for angiotensin type 2 receptor (AT2R), which may have renoprotective functions, was reduced in PcKOTsc1 mouse glomeruli (Supplemental Figure 2, G and I) (25–27).

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Figure 2

Characterization of PcKOTsc1 mice.

(A) Enhanced S6 phosphorylation, podocyte hypertrophy, and mesangial expansion in PcKOTsc1 glomeruli. Renal tissues from 4-week-old mice of the indicated genotypes were stained with phospho-S6, WT1, H&E, and PAS as indicated. Representative glomeruli from wild-type, PcKOTsc1 (KO), and rapamycin-treated PcKOTsc1 (KO+rapa) are shown. Rapamycin treatment (1 mg/kg, i.p., 3 times/wk) was performed from 2 to 4 weeks of age. (B) S6 but not Akt phosphorylation was enhanced in PcKOTsc1 glomeruli. Levels of phospho-S6, S6, phospho-Akt (Ser473), and Akt in glomeruli from mice of the indicated genotypes were determined by Western blot. The levels of phosphorylation of each protein were quantified. Data are expressed as fold increase in the ratio of phospho-protein to total protein. *P < 0.001 versus other groups; mean ± SEM, n = 3. (C) Enhanced matrix protein mRNAs in PcKOTsc1 glomeruli. The expression levels of fibronectin (FN) and type IV collagen (COL4) mRNAs in the indicated mouse glomeruli were determined by quantitative RT-PCR. Data were normalized to Hprt1. *P < 0.001 versus other groups; mean ± SEM, n = 7. (D) Enhanced deposition of the matrix proteins in PcKOTsc1 glomeruli. Representative glomerular images of staining for fibronectin and type IV collagen in the indicated animals are shown. The ratio of positive staining area to glomerular area in the indicated animals was determined (10 glomeruli/mouse, 3 mice/group). *P < 0.001 versus other groups, mean ± SEM. Original magnification, ×400 (A and D).

mTORC1 activation in podocytes causes podocyte injury and proteinuria.

Next we monitored proteinuria in PcKOTsc1 mice. Consistent with the time course of mTORC1 activation in podocytes (Supplemental Figure 2B), PcKOTsc1 mice showed selective albuminuria starting at 2 weeks of age. The majority of PcKOTsc1 mice showed massive and nonselective proteinuria around 3–4 weeks age (Supplemental Figure 3A). In contrast, neither heterozygous nor wild-type mice had proteinuria even after 8 weeks. Importantly, rapamycin treatment completely eliminated the incidence of proteinuria in PcKOTsc1 mice (Figure ​(Figure3,3, A and B).

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Figure 3

PcKOTsc1 mice show proteinuria and podocyte loss.

(A) Rapamycin treatment prevents proteinuria in PcKOTsc1 mice. Scheme of administration of rapamycin is shown above. Urine (1 μl ) from mice of the indicated genotypes was subjected to SDS-PAGE with Coomassie blue staining (n = 6 each group). veh, vehicle. Asterisk indicates a leak of sample from lane 6. (B) Urinary albumin/creatinine ratio is shown. Urinary albumin and creatinine concentrations in 4-week-old mice were determined by ELISA. *P < 0.001 versus other groups, mean ± SEM, n = 6. Note logarithmic scale for the y axis. Rapamycin treatment was performed from 2 weeks of age as indicated in Figure ​Figure2A.2A. (C) TEM analyses reveal abnormal foot process and GBM in PcKOTsc1 and wild-type mice. Representative TEM images of the indicated animals (3 mice/group) from 4 and 8 weeks of age are shown. Rapamycin treatment was as in Figure ​Figure2A.2A. Original magnification, ×13,500 (top row), ×7,900 (bottom row). Scale bars: 1 μm. (D) Scanning EM analyses show abnormal podocyte in PcKOTsc1 mice, which was prevented by rapamycin treatment. Rapamycin treatment was as in Figure ​Figure2A.2A. Original magnification, ×2,000. Scale bars: 5 μm (top row); 1.25 μm (bottom row). (E) Podocyte loss in PcKOTsc1 mice. The ratio (the number of WT1-positive cells/glomerular tuft area [μm²]) was determined in 15~30 glomeruli from the indicated animals. *P < 0.001 versus wild-type and/or KO with rapamycin treatment, mean ± SEM, n = 3–5 mice.

Transmission EM (TEM) analyses revealed an effacement of podocytes, as well as mild thickening and irregular shape of the GBM in 4 week-old PcKOTsc1 mice (Figure ​(Figure3C).3C). At 8 weeks, no podocyte foot processes could be identified (Figure ​(Figure3C),3C), and local loss of the fenestrations was observed (data not shown) in PcKOTsc1 mice. As expected, rapamycin treatment prevented all these pathological changes in PcKOTsc1 mice, indicating that mTORC1 activation in podocytes causes cell-autonomous phenotypic changes and dysfunction that lead to malfunction of filtration barrier. Strikingly, scanning EM revealed that mTORC1 activation in podocytes disrupted the normal architecture of foot processes with enlarged podocyte body (Figure ​(Figure3D3D and Supplemental Figure 2C).

In both type 1 and type 2 DN, the number and density of podocytes are reduced, and the decreased number of podocytes per glomerulus is the strongest predictor of progression of DN (2, 3, 28, 29). To investigate podocyte number in PcKOTsc1 mice, we determined average numbers of podocytes per glomerular tuft area by counting WT1-positive podocytes in PcKOTsc1 mice at various ages (Figure ​(Figure3E).3E). Indeed, the number of podocytes in 3- and 4-week-old PcKOTsc1 mice but not those in 1- to 2-week-old PcKOTsc1 mice was significantly decreased compared with those in wild-type mice. Furthermore, rapamycin treatment for 2 weeks prevented podocyte loss, indicating that mTORC1 activation causes loss of podocytes. Interestingly, these results also suggest that the podocyte loss may not be the sole reason for the initial proteinuria in PcKOTsc1 mice, because 2-week-old PcKOTsc1 mice exhibited proteinuria before apparent podocyte loss.

Given the fact that mTORC1 activation in podocytes induces autonomous podocyte injury and proteinuria, our data support the idea that diabetes-induced mTORC1 activation in podocytes may have a direct role in podocyte injury and proteinuria, thereby contributing to the progression of DN.

mTORC1-dependent podocyte injury and dysfunction are reversible.

Clinical experience suggests that tight control of blood glucose levels and/or the proper calorie restriction ameliorate proteinuria in early DN (30). Therefore, we wanted to determine whether mTORC1-dependent podocyte injury is a reversible phenotypic change. To this end, we started rapamycin treatment from 4 weeks age in PcKOTsc1 mice whose podocytes were already injured (Figure ​(Figure3).3). Surprisingly, after only 2 weeks of treatment, the massive proteinuria observed at 4 weeks was largely diminished and finally disappeared at 10 weeks age (6 weeks treatment) in PcKOTsc1 mice (Figure ​(Figure4,4, A and B). Histological and TEM analyses of glomeruli revealed that the eosin-positive compartments were cleared in the mesangial region (Figure ​(Figure4C)4C) and that foot process formation was significantly restored, although some irregular shape of the GBM still remained (Figure ​(Figure4D).4D). Scanning EM studies revealed that podocyte foot processes were regenerated around the podocyte body after 2 weeks of rapamycin treatment (Figure ​(Figure4E).4E). Although we observed different stages of glomeruli in which foot processes were regenerating in the same kidney, it appeared that a single major process sprouted from the podocyte cell body at the beginning, and foot processes developed from this structure (Figure ​(Figure4E).4E). These results indicate that mTORC1-dependent podocyte injury and the associated pathological phenotypes are reversible upon inactivation of mTORC1 by rapamycin and lowering mTORC1 activity may induce podocyte replenishment. Importantly, however, rapamycin treatment failed to rescue the phenotypes of 8-week-old PcKOTsc1 mice (data not shown), suggesting that once severe podocyte damage had occurred (Figure ​(Figure3C),3C), such as a severe loss of podocytes with sclerotic glomeruli, these changes might not be rescued with rapamycin treatment. Taken together, the results indicate that reduction of mTORC1 activity may rescue both the decrease in podocyte numbers and the dysfunction of filtration barrier when applied prior to a certain threshold. This is consistent with the observations that patients with DN having microalbuminuria can be effectively treated, but overt proteinuria is often refractory to intensive therapies (30, 31).

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Figure 4

Rapamycin reverses established phenotypes in PcKOTsc1 mice.

(A and B) Rapamycin treatment ameliorated the established proteinuria in PcKOTsc1 mice. Proteinuria-positive 4-week-old PcKOTsc1 mice (n = 9) were treated with rapamycin for 2 and 6 weeks. The levels of urinary protein from 9 mice were tested at the indicated time points and visualized by Coomassie staining (n = 9) (A), and the ratio of albumin to creatinine concentration is shown (B). *P < 0.001 versus other groups, mean ± SEM, n = 9; note logarithmic scale for y axis. The ratios for 4- and 10-week-old wild-type mice are also shown. (C) H&E staining of renal tissue from 4-week-old untreated and 10-week-old treated PcKOTsc1 mice (rapamycin treatment for 6 weeks). Eosin-positive area in glomerulus was measured in 30 glomeruli from 4-week-old wild-type and PcKOTsc1 mice as well as 10-week-old treated PcKOTsc1 mice. *P < 0.001 versus other groups, no statistical significance between WT 4 weeks and KO with rapamycin 10 weeks, mean ± SEM, n = 3~5. (D) TEM analyses of 10-week-old PcKOTsc1 mice (rapamycin treatment for 6 weeks). Foot process width at the level of slit diaphragm was measured (20~50 foot processes/glomerulus, 3 glomeruli/mouse, 3 mice/group). *P < 0.001 versus other groups, mean ± SEM. (E) Scanning EM analyses of 6-week-old PcKOTsc1 mice (rapamycin treatment for 2 weeks). Representative glomerulus from 2 different rapamycin-treated PcKOTsc1 mice is shown. Original magnification, ×400 (C), ×7,900 (D), ×2,000 (E). Scale bars: 10 μm (top row); 2.5 μm (bottom row).

High mTORC1 activation in podocytes of adult mice also leads to glomerular sclerosis.

PcKOTsc1 mice develop glomerulosclerosis and die due to renal failure by 14 weeks after birth (Figure ​(Figure5A).5A). The causes of early death by 6 weeks may include acute hypovolemic shock due to loss of plasma proteins. We observed massive ascites in these PcKOTsc1 mice (Supplemental Figure 3B). The levels of serum creatinine concentration but not glutamic pyruvic transaminase (sGPT) activity were significantly elevated above the normal reference range in 8-week-old PcKOTsc1 mice (Supplemental Figure 4, A and B) (32). To examine the possibility of strain or genetic background effects on survival and renal phenotype, we also transferred these alleles to the C57BL/6 strain. Although survival was slightly improved for PcKOTsc1 mice in C57BL/6 (Figure ​(Figure5A,5A, orange line), they developed proteinuria and glomerulosclerosis at a similar rate (Supplemental Figure 5, A–C), and all died by 15 weeks of age. Since the glomeruli in young mice are still developing, it is possible that mis-coordination of growth or development between the podocytes and the other glomerular compartments could be a reason for the phenotypic changes in podocytes in the PcKOTsc1 mice. Moreover, DN normally develops in adults with diabetes. To test whether podocyte mTORC1 activation in adult mice also causes podocyte injury, we treated PcKOTsc1 mice with rapamycin until 15 weeks of age; withdrew the treatment; and monitored proteinuria, histological changes of glomeruli, and survival. Interestingly, these mice rapidly developed proteinuria within 1 week of withdrawal of rapamycin (Figure ​(Figure5B).5B). Seven weeks after withdrawal of rapamycin treatment, the eosin- and PAS-positive components were accumulated in the mesangial area, and the formation of a tuft adhesion to Bowman’s capsule was marked, indicative of glomerulosclerosis in PcKOTsc1 mice (Figure ​(Figure5C).5C). Furthermore, the trend of the survival curve of these mice was essentially similar to that of young PcKOTsc1 mice not treated with rapamycin (Figure ​(Figure5A).5A). These data indicate that mTORC1 activation in podocytes also causes injury and glomerulopathy in developed glomeruli, suggesting a role of mTORC1 activation in the acquired glomerulopathy.

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Figure 5

mTORC1 activation in podocytes causes proteinuria and mesangial expansion in adult mice.

(A) Rapamycin prevents death in PcKOTsc1 mice. Mice were treated with rapamycin as indicated. Survival curve of the indicated animals is shown. Wild-type, n = 50; heterozygous (Het), n = 50; KO, n = 50; KO with rapamycin treatment (off-on), n = 15; KO with rapamycin treatment (off-on-off), n = 10; KO C57BL/6 (B6) background, n = 30. (B) mTORC1 activation in podocytes causes proteinuria and mesangial expansion in adult mice. Withdrawal of rapamycin treatment in adult PcKOTsc1 mice led to rapid onset of proteinuria and glomerulopathy. Scheme of administration of rapamycin is shown at top. Urinary albumin concentrations of the indicated animals were determined (WT 4 wk; KO 15 wk [rapamycin treatment from 2 to 15 weeks]; KO 16 wk, 19 wk, and 21 wk [off-on-off; 1 wk, 4 wk, and 6 wk after withdrawal of rampamycin treatment]). *P < 0.001 versus WT and KO 15 wk, mean ± SD, n = 6. (C) Withdrawal of rapamycin treatment in adult PcKOTsc1 mice led to mesangial expansion. Scheme of administration of rapamycin is shown at top. H&E and PAS staining of the representative glomeruli are shown. PAS-positive area in glomerulus was measured in 30 glomeruli from the indicated animals. *P < 0.001 versus other groups, n = 4. Original magnification, ×400.

mTORC1 regulates the localization of slit diaphragm proteins.

Dysfunction of slit diaphragm or cytoskeletal proteins in podocytes leads to foot process effacement with proteinuria and is followed by podocyte loss as seen in genetic glomerulopathies (33, 34). In DN, there are mixed reports regarding slit diaphragm expression of proteins such as nephrin in early diabetic glomeruli, though it seems consistent that nephrin expression is reduced in the later stages of DN, possibly due to the decreased number of podocytes (35–38). Since mTORC1 activation in podocytes causes cell-autonomous morphological changes, we hypothesized that mTORC1 activation may inhibit gene expression of the slit diaphragm protein in podocytes. However, mRNA expression of representative slit diaphragm proteins such as nephrin in 3-week-old PcKOTsc1 mice, which have already developed proteinuria, was enhanced compared with that in control mice (Supplemental Figure 2G). However, immunogold nephrin staining characterized by TEM revealed that nephrin expression at interpodocyte filtration slit areas was largely diminished in PcKOTsc1 mice (Figure ​(Figure6A).6A). We found that total nephrin expression in glomeruli isolated from 3-week-old PcKOTsc1 mice was maintained at levels similar to those in wild-type mice (Figure ​(Figure6B),6B), raising the possibility that nephrin is mislocalized in the PcKOTsc1 podocytes. Consistent with this hypothesis, confocal microscopic analyses revealed that nephrin was predominantly expressed in cytoplasm, with little found on the plasma membrane in the podocytes of PcKOTsc1 mice (Figure ​(Figure6C).6C). Furthermore, nephrin failed to colocalize well with synaptopodin, which is abundant in the vicinity of podocyte membranes in PcKOTsc1 mice. Importantly, nephrin was seen on the plasma membrane and colocalized with synaptopodin in PcKOTsc1 mice treated with rapamycin (from 2 to 3 weeks of age). Together these data suggest that abnormal mTORC1 activation results in mislocalization of nephrin, thereby potentially disrupting the integrity of the slit diaphragm and contributing to the effacement of foot processes and the genesis of proteinuria. We found that podocin was also expressed in cytoplasm in PcKOTsc1 mice (data not shown); however, Neph1 was properly membrane localized, suggesting that mTORC1 activation leads to mislocalization of the specific slit diaphragm proteins (Supplemental Figure 6A).

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Figure 6

Hyperactivation of mTORC1 leads to mislocalization of slit diaphragm proteins.

(A) Nephrin expression at interpodocyte filtration slit area is diminished in PcKOTsc1 mice. Representative electron micrographs of immunogold nephrin staining in 3-week-old mice are shown. Number of immunogold particles for nephrin at the slit diaphragm was measured in 290 slit diaphragms from 2 wild-type and 170 slit diaphragms from 2 PcKOTsc1 mice. *P < 0.001 versus wild-type, mean ± SEM. (B) Nephrin and actin expression in the indicated glomeruli were determined by Western blotting. Levels of nephrin expression in the indicated genotypes were quantified (nephrin/actin). Data are expressed as fold increase. P = 0.67, mean ± SEM, n = 3. (C) Nephrin is mislocalized in PcKOTsc1 podocytes. Confocal microscopic analyses of nephrin and synaptopodin expression in the indicated animals (3 weeks old) are shown. Rapamycin treatment was performed for 1 week. Areas indicated by squares in the Merge column are shown at higher magnification in the right two columns. Original magnification, ×13,500 (A), ×400 (C).

To investigate whether this regulation between mTORC1 and nephrin could be observed in vitro, we examined the localization of nephrin in COS7 cells, which normally do not express podocyte-specific proteins, by ectopically expressing nephrin and Rheb, a potent mTORC1 activator. Transfected nephrin was localized in both plasma membrane and peri-nuclear regions (Figure ​(Figure7A).7A). As seen in the PcKOTsc1 podocytes, activation of mTORC1 by Rheb resulted in loss of plasma membrane localization of nephrin. Importantly, under these conditions, rapamycin increased nephrin plasma membrane localization. These results suggest that mTORC1 may have a specific role in the regulation of nephrin localization.

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Figure 7

Mislocalization of nephrin by abnormal mTORC1 activation and diabetes.

(A) Membrane localization of nephrin is disrupted by Rheb. Exogenous nephrin expression pattern was analyzed by confocal microscopy. Human nephrin (Myc-Nephrin) was expressed with or without Rheb (HA-Rheb) in COS7 cells. Nephrin and Rheb expression was determined by rabbit polyclonal nephrin and mouse monoclonal HA antibodies, respectively. The arrowheads indicate membrane expression of nephrin. Rapamycin (20 nM) treatment was for 12 hours. (B) Rapamycin treatment restores membrane localization of nephrin in db/db mice. The pattern of nephrin expression of the indicated animals (25 weeks of age) was monitored by confocal microscopy. Rapamycin treatment was performed from 8 to 25 weeks. The arrowheads indicate the accumulation of nephrin in cytoplasm. (C) Membrane localization of nephrin was reduced in human type 2 DN. Nephrin staining showed a granular pattern expression in cytoplasm with reduced membrane expression (NOR, normal subject; DN p1, DN patient 1). Original magnification, ×600 (A), ×400 (B, and C).

To investigate whether nephrin mislocalization is a common pathologic phenotype associated with mTORC1 activation, we examined the pattern of nephrin expression in diabetic animals using confocal microscopy. In nondiabetic (db/+) mice, a majority of nephrin was linearly expressed on the membrane of podocyte foot process. In contrast, nephrin was diffusely expressed and accumulated in some organelles in the cytoplasm in db/db mice podocytes (Figure ​(Figure7B).7B). Rapamycin treatment diminished cytoplasmic nephrin, and almost all of the nephrin was relocated along with plasma membrane in foot processes. Consistent with these observations in the mouse model, human nephrin expression in normal subjects exhibited a liner pattern of expression, whereas it showed a more granular pattern with reduction of its membrane expression in human type 2 DN glomeruli (Figure ​(Figure7C).7C). Collectively, the above data indicate that proper control of mTORC1 activity in podocytes is important for maintaining intact localization of certain slit diaphragm proteins.

mTORC1 activation converts podocytes to a fibroblastic phenotype and causes podocyte detachment.

It has been reported that podocytes display fibroblast-like phenotypic changes and this may cause podocyte loss/detachment in DN (10, 39–43). These phenotypes include the expression of desmin and the redistribution of zona occludens–1 (ZO-1). However, the molecular mechanism by which diabetes causes this phenotypic change is not well understood. Confocal microscopic analyses showed that ZO-1 expression is redistributed in the cytoplasm (loss of liner expression pattern) and desmin is expressed in the cytoplasm of the WT1-positive cells, indicating that podocytes display a fibroblastic phenotypic change in PcKOTsc1 mice (Figure ​(Figure8A).8A). Moreover, rapamycin treatment blocked these phenotypic changes. These results suggest that high mTORC1 signaling may be a potential mechanism for fibroblastic phenotypic changes in the podocytes seen in diabetes. In 4-week-old PcKOTsc1 mice, whose podocyte number was significantly decreased (Figure ​(Figure3E),3E), the levels of desmin expression in the remaining podocytes were further enhanced (Supplemental Figure 8, A–D). This observation suggests that the fibroblast-like phenotypic change in podocytes with desmin expression might be a trigger for podocyte loss. Consistent with this hypothesis, we observed cells expressing podocyte-specific mRNAs such as nephrin and podocin in the urine from PcKOTsc1 mice but not those from wild-type mice, suggestive of a detachment of podocytes from the GBM in PcKOTsc1 mice (Figure ​(Figure8B).8B). However, we did not observe a significant increase in apoptotic cells in PcKOTsc1 mouse glomeruli (data not shown). These observations suggest that mTORC1 activation may induce podocyte loss mainly due to detachment along with epithelial-mesenchymal transition–like (EMT-like) phenotypic changes.

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Figure 8

Hyperactivation of mTORC1 induces ER stress and EMT-like phenotype in podocytes.

(A) EMT-like phenotypic changes were induced in PcTSC1KO podocytes. Double staining with desmin and WT1 antibodies was performed in renal tissues from the indicated animals and analyzed by confocal microscopy (top row). Arrowheads indicate desmin expression in the podocytes. Rapamycin treatment was performed for a week. ZO-1 expression was analyzed by confocal microscopy (bottom row). Arrowheads indicate the liner expression pattern of ZO-1. (B) Detection of nephrin and podocin mRNA in PcTSC1KO urinary sediments. Twenty-four-hour urine samples were collected from the indicated animals. RT-PCR was performed using purified mRNA from the sediments. Glomerular mRNA was used as a positive control (p). (C) Accumulation of GRP78 in PcKOTsc1 podocytes (3 weeks of age). Double staining using anti-GRP78 and anti-synaptopodin antibodies in the indicated glomeruli is shown. Rapamycin treatment was performed for 1 week. (D) PBA treatment protects podocyte loss in PcKOTsc1 mice. Staining with GRP78 plus synaptopodin, nephrin, H&E, and WT1 is shown in the indicated animals (8 weeks of age). PBA treatment (400 mg/kg/d, i.p.) was performed for 6 weeks. (E) Podocyte number in the indicated glomerular sections from 8-week-old animals was measured. The ratio (number of WT1-positive cells/glomerular tuft area [μm²]) was determined in 15~30 glomeruli from the indicated animals as in Figure ​Figure3E.3E. *P < 0.001 versus other groups, mean ± SEM, n = 4~8. (F) Urinary albumin concentrations were measured in the indicated animals. *P < 0.001 versus other groups; no statistical significance was observed between KO and KO with PBA treatment; mean ± SEM, n = 4~8. Original magnification, ×400 (A, C, and D).

ER stress is an essential factor for podocyte loss in PcKOTsc1 mice.

It has been postulated that ER stress is enhanced in tissues such as pancreatic β cells as well as renal tubular and glomerular cells under diabetic conditions (44, 45). However, the role of diabetes-induced ER stress in podocytes remains unclear. We found that GRP78, which is induced during ER stress, was expressed at higher levels in the podocytes of PcKOTsc1 mice. The accumulation of GRP78 in PcKOTsc1 podocytes was significantly reduced by rapamycin treatment, suggestive of the elevated ER stress being induced by high mTORC1 activity (Figure ​(Figure8C).8C). Proper protein post-translational modifications such as glycosylation and oxidation in the ER are essential for nephrin to localize at the membrane (46); furthermore, it has been reported that ER stress triggers EMT in thyroid cells (47). To study the role of ER stress in podocyte phenotypic changes in PcKOTsc1 mice, we utilized 4-phenyl butyric acid (PBA), a chemical chaperone that has been used to reduce ER stress in the pancreatic β cells under diabetic conditions (48). We found that PBA treatment effectively reduced the accumulation of GRP78 (Figure ​(Figure8D)8D) in the podocytes and partially but significantly prevented the loss of podocytes in 8-week-old PcKOTsc1 mice (Figure ​(Figure8,8, D and E). Furthermore, loss of nephrin and synaptopodin expression was moderately restored in PBA-treated 8-week-old PcKOTsc1 mice (Figure ​(Figure8D).8D). These data indicate that high mTORC1-induced ER stress causes podocyte loss in PcKOTsc1 mice. In agreement with this hypothesis, reduction of ER stress with PBA treatment in PcKOTsc1 mice significantly attenuated desmin expression in podocytes (Supplemental Figure 8, A–D), suggesting that the mTORC1-induced change in podocytes to a fibroblastic phenotype was at least in part due to ER stress. It is worth noting that PBA treatment has little effect on S6 phosphorylation in PcKOTsc1 mouse podocytes (Supplemental Figure 8, B, C, and E), indicating that unlike rapamycin, PBA only attenuates the mTORC1-induced ER stress pathway. Intriguingly, despite the protective role of PBA in the fibroblastic phenotypic change and loss of podocytes in PcKOTsc1 mice, it had little effect on proteinuria in the course of the treatment in PcKOTsc1 mice (Figure ​(Figure8F).8F). Consistent with this observation, PBA treatment failed to restore nephrin localization to the plasma membrane (Supplemental Figure 8F). These data indicate that ER stress induced by mTORC1 activation may contribute to podocyte damage in this model, but does not explain nephrin mislocalization.

Human materials.

Human kidney biopsy material was collected after receipt of informed consent from the patients, with the protocol approved by the Scientific-Ethical Committee of the Hospital of Milan (Ospedale S. Carlo Borrome). Frozen kidney sections used for the study were those remaining after completion of diagnosis of type 2 diabetic nephropathy class 1, according to the classification previously reported (74). Control kidneys were from the unaffected pole of tumor nephrectomies.

Animals and animal care.

To generate mice with Tsc1 deletion specifically in podocytes (PcKOTsc1), we crossed Tsc1^fl/fl mice (129S4/SvJae) with Tg (Nphs2 [podocin]-Cre) (C57BL/6J) mice (63). [Tg (Nphs2-Cre), Tsc1^fl/+] × [Tsc1^fl/fl] mating was used for this study. Mice were maintained on the segregating (129S4/SvJae; C57BL/6J) genetic background. To generate C57BL/6J background PcKOTsc1 mice, Tg (Nphs2-Cre), Tsc1^fl/+ (129S4/SvJae; C57BL/6J) mice were crossed with C57BL/6J mice, followed by 6 sequential generations of backcrossing with C57BL/6J mice. Offspring from the backcrossed [Tg (Nphs2-Cre), Tsc1^fl/+] × [Tsc1^fl/fl] mating were used for the study. For podocyte-specific Raptor-KO diabetic mice, C57BL/6J background [Tg (Nphs2-Cre), Raptor^fl/+] mice were crossed with BKLS background db/m mice to generate male [Tg (Nphs2-Cre), Raptor^fl/+, db/+]F₁ mice (49). [Tg (Nphs2-Cre), Raptor^fl/+, db/+]F₁ male mice were crossed with db/m females, followed by additional sequential generations of backcrossing with db/m mice. F₃ [Tg (Nphs2-Cre), Raptor^fl/+, db/+] male mice were crossed with F₃ females [Raptor^fl/+, db/+] to generate [Tg (Nphs2-Cre), Raptor^fl/+, db/db] mice. To minimize genetic variations, the full and half siblings were grouped into control ([Raptor^fl/+, db/+] and [Raptor^fl/fl, db/+]), DM ([Raptor^fl/+, db/db] and [Raptor^fl/fl, db/db]), and DM Raptor^+/– ([Tg (Nphs2-Cre), Raptor^fl/+, db/db]) mice and used for the study. Animal experiments were conducted following a protocol approved by the University of Michigan Committee on the Use and Care of Animals (UCUCA).

Immunohistochemistry.

For immunohistochemistry, we used both paraffin-embedded and frozen tissue sections in mouse models. For paraffin-embedded tissue, the renal tissues were fixed by transcardiac perfusion with PBS containing 4% paraformaldehyde. For frozen sections, transcardiac perfusion with PBS containing 20 mM NaF was performed to eliminate blood cells and to protect protein dephosphorylation during sample preparation. Staining was visualized by indirect/direct immunofluorescence or DAB. For human samples, acetone-fixed tissue sections were used for indirect immunofluorescence staining.

Immunoelectron microscopy.

For immunogold staining, renal tissues were fixed by transcardiac perfusion with PBS containing 4% para­formaldehyde. Tissues were dehydrated and embedded in LR white (Electron Microscopy Sciences). Ultrathin kidney cortical sections (70 nm) were mounted onto Formvar/carbon-coated nickel grids (Electron Microscopy Sciences). Aldehyde quenching with 0.05 mol/l glycine and antigen retrieval with citrate buffer (95°C for 10 minutes) were performed before blocking with normal donkey serum, bovine serum albumin, cold water fish gelatin, and Tween 20 (Electron Microscopy Sciences). The tissues were incubated with rabbit anti-nephrin antibody overnight at 4°C in blocking buffer plus 0.2% BSA-c (Aurion; Electron Microscopy Sciences). After washing, sections were incubated with a donkey anti-rabbit antibody conjugated to 10 nmol/l gold particles at 1:20. After rinsing, grids were fixed in 2.5% glutaraldehyde in 0.1 mol/l phosphate buffer and post-stained with uranyl acetate and lead citrate.

EM analysis.

The renal samples were fixed with Sorensen’s phosphate buffer containing 4% paraformaldehyde and 2.5% glutaraldehyde. The processed samples were analyzed by Philips CM100 TEM and AMRAY 1910 field emission scanning EM (FEG-SEM).

In vivo physiological studies.

Blood glucose levels were measured using an automated blood glucose reader (Accu-Chek, Roche). Urinary albumin and creatinine were determined using mouse-specific ELISA (Albuwell M kit) and Creatinine Companion kit (Exocell). Mouse serum creatinine (Arbor Assays) and GPT (Cayman Chemical) concentrations were also measured by ELISA. Glomerular tuft area, PAS-positive mesangial area, and other pixel densities obtained by immunohistochemical experiments were measured by ImageJ software. The number of total glomerular or WT1-positive glomerular cells (podocytes) was counted in glomerular cross-sections (4 μm).

Isolation of glomeruli and mRNA quantification by RT-PCR analysis.

Pure glomeruli were collected by magnetic bead–based isolation. The transcardiac perfusion was performed with PBS containing precleaned beads (Dynabeads, Invitrogen). The perfused renal cortex was briefly digested with collagenase, and the glomeruli choked with beads were isolated by magnets (IMag, BD). Total RNA was purified and used for reverse transcription by SuperScript First-Strand Synthesis System (Invitrogen). Quantitative PCR was carried out with specific primers (Supplemental Information). For quantitative analysis, the samples were normalized to Hprt1 (hypoxanthine phosphoribosyl transferase 1) gene expression using the ΔΔCT value method.

Western blot analysis.

The isolation of glomeruli was performed as above. Purified glomeruli from one mouse were lysed in lysis buffer (10 mM Tris-HCl [pH 7.5], 100 mM NaCl, 1% NP-40, 50 mM NaF, 20 mM β-glycerophosphate, 2 mM EDTA, 1 mM DTT, 1 mM PMSF, 10 μg/ml leupeptin, and 10 μg/ml aprotinin) as a lysate sample. Lysates were then boiled in SDS sample buffer (20 mM Tris [pH 6.8] 8% SDS, 0.05% bromophenol blue, 40% glycerol, 20% 2-mercaptoethanol) and subjected to SDS-PAGE and Western blotting according to standard techniques.

We thank A. Saltiel and F. Brosius (University of Michigan, Ann Arbor) for discussion and C. Tsui, R. Verma, and M. Uhm (University of Michigan, Ann Arbor) for technical assistance. We thank C. Edwards, S. Meshinchi, and D. Sorenson (Microscopy and Image Analysis Laboratory, University of Michigan, Ann Arbor) for EM analyses. This work was supported by a George M. O’Brien Renal Center Developmental Research (DR)/Pilot and Feasibility (P&F) grant (to K. Inoki), a Juvenile Diabetes Research Foundation Award (to K. Inoki), NIH grants DK083491 (to K. Inoki) and GM051586 (to K.-L. Guan), and the Excellence Initiative of the German Federal and State Governments (EXC 294 5o T.B. Huber).