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Podocyte injury has a pivotal role in the pathogenesis of diabetic nephropathy (DN). MicroRNA-27a (miR-27a), peroxisome proliferator-activated receptor γ (PPARγ) and β-catenin pathways have been involved in the pathogenesis of DN. Herein, we asked whether miR-27a mediates podocyte injury through PPARγ/β-catenin signaling in DN. The functional relevance of miR-27a, PPARγ and β-catenin were investigated in cultured podocytes and glomeruli of diabetic rats and patients using in vitro and in vivo approaches. Podocyte injury was assessed by migration, invasion and apoptosis assay. Biological parameters were analyzed using enzyme-linked immunosorbent assay. We found that high glucose stimulated miR-27a expression, which, by negatively targeting PPARγ, activated β-catenin signaling as evidenced by upregulation of β-catenin target genes, snail1 and α-smooth muscle actin (α-SMA) and downregulation of podocyte-specific markers podocin and synaptopodin. These changes caused podocyte injury as demonstrated by increased podocyte mesenchymal transition, disrupted podocyte architectural integrity and increased podocyte apoptosis. Furthermore, we provide evidence that miR-27a contributed to unfavorable renal function and increased podocyte injury in diabetic rats. Notably, miR-27a exhibited clinical and biological relevance as it was linked to elevated serum creatinine, proteinuria and reduced creatinine clearance rate. In addition, miR-27a upregulation and activation of PPARγ/β-catenin signaling were verified in renal biopsy samples from DN patients. We propose a novel role of the miR-27a/PPARγ/β-catenin axis in fostering the progression toward more deteriorated podocyte injury in DN. Targeting miR-27a could be a potential therapeutic approach for DN.
Diabetes mellitus, one of the fastest growing metabolic disorders in the world, contributes to about 3.2 million diabetes-related deaths annually.1, 2 Diabetic nephropathy (DN) is the most common cause of end-stage renal disease and cardiovascular events.3, 4 In the past decades, although the first-line therapy such as angiotensin-converting enzyme inhibitors/angiotensin II receptor blockers can slow but cannot stop the progression of DN, which urgently calls for innovative therapeutic strategies.5, 6
Adult podocytes are unique terminally differentiated glomerular epithelial cells critical in maintaining the integrity of the glomerular filtration barrier.7 Reduced podocyte number leads to proteinuria and glomerulosclerosis in diabetic8 and non-diabetic glomerular diseases.9 Loss of podocyte-specific molecules, such as podocin, synaptopodin and nephrin, causes podocyte injury and epithelial to mesenchymal transition.10, 11
Accumulating evidence has demonstrated that β-catenin activation is closely related to the emergence and development of various disease pathologies.12, 13 The pathway activates downstream target genes, such as snail1, c-myc and α-smooth muscle actin (α-SMA), thereby regulating many biological processes through a complex of β-catenin and the T-cell factor/lymphoid-enhancer factor 1 (TCF/LEF1) family.14 Wnt stabilizes cytosolic β-catenin, which then binds to TCF/LEF1 in the nucleus and recruits transcription factors Brg1 and CREB-binding protein to initiate Wnt-targeted gene expression.15, 16 β-Catenin activation triggers the activities of downstream transcription factors and activates the mesenchymal transition of podocytes.17, 18
Peroxisome proliferator-activated receptor gamma (PPARγ) is an important regulator of cell proliferation and differentiation.19 Our previous study has revealed that high glucose stimulated PPARγ phosphorylation in cultured renal tubular epithelial cells.20 Other studies have shown that activation of PPARγ can promote the degradation of β-catenin in mesenchymal stem cells and direct interaction with β-catenin.21 Recently, a PPARγ region and β-catenin binding domain were identified to have high homology with TCF/LEF, which facilitates its interaction with β-catenin.22 Undoubtedly, the PPARγ/β-catenin signaling is drawing increasing attention and has a critical role in the pathogenesis of diseases.
MicroRNAs (miRNAs) regulate gene expression after recognition of specific binding to the 3′-untranslated region (UTR) of target mRNAs, causing mRNA deadenylation and degradation or translational repression.23 Alterations of gene expression resulting from aberrant expression of miRNAs are associated with numerous diseases.24, 25
Along with others, we have shown that microRNA-27a (miR-27a), by targeting PPARγ, exacerbates renal tubulointerstitial fibrosis and mesangial cell injury in DN.20, 26 In this study, our aim was to explore whether miR-27a promotes podocyte injury through PPARγ-mediated β-catenin activation in DN.
To investigate the effect of high glucose stimulation on the expression of miR-27a and downstream genes, podocytes were cultured in different conditions and biological behaviors were observed. First, the level of miR-27a was increased in high glucose (30mM) cultured podocytes as determined by quantitative real-time RT-PCR (qRT-PCR) analysis (Figure 1a). As shown in Figure 1b, high glucose suppressed the expression of PPARγ and podocyte-specific marker podocin, but activated β-catenin and its target genes, snail1 and α-SMA. Mannitol (30mM), however, had no effect on the expression of these molecules. Next, we investigated the effect of time and glucose concentration on miR-27a expression. We found that miR-27a was increased in a time-dependent (Figure 1c) and dose-dependent (Figure 1d) manner as detected by qRT-PCR. Then, we examined whether miR-27a modulated PPARγ-mediated downstream gene expression levels in high glucose. As illustrated by qRT-PCR (Figures 1e and f) and western blot analyses (Figures 1g and h), the decreased expression level of PPARγ was time and dose dependent. In addition, β-catenin and its target genes, snail1 and α-SMA, were markedly increased and podocyte-specific marker podocin was downregulated. We finally asked whether miR-27a-induced downstream changes depended on PPARγ. Of note, the luciferase activity of wild-type 3′-UTR of PPARγ was significantly increased when treated with miR-27a inhibitor (miR-27ai) but decreased with miR-27a mimics (miR-27am) as compared with the mutant (Figure 1i), indicating that PPARγ is a direct downstream target of miR-27a. These results indicate a correlation between miR-27a and PPARγ-mediated downstream gene expression levels in high glucose cultured podocytes.
Having discovered the upregulation of miR-27a in high glucose condition, we next explored the underlying mechanisms of miR-27a-induced podocyte injury. As shown in Figure 2a, miR-27ai decreased miR-27a expression in podocytes by qRT-PCR analysis. Furthermore, high glucose-stimulated PPARγ phosphorylation was attenuated by miR-27ai (Figure 2b). In addition, miR-27ai inactivated the expression of β-catenin and its target genes, snail1 and α-SMA, but increased podocin level in high glucose instead of the normal glucose condition as shown by western blot (Figure 2b) and qRT-PCR analyses (Figure 2c). These data suggest underlying mechanisms may be involved in the post-translational modification of PPARγ.
This observation led us to ask whether PPARγ physically interacted with β-catenin in mediating the downstream events. To test this hypothesis, we carried out co-immunoprecipitation analysis and examined the interaction between phosphorylated PPARγ and active β-catenin. As shown in Figure 2d, physical interaction between phosphorylated PPARγ and active β-catenin were detected in podocytes and was strengthened by high glucose stimulation. Strikingly, miR-27ai rendered podocytes less vulnerable to the deleterious effects of high glucose concentrations, leading to decreased abilities of migration (Figure 2e), invasion (Figure 2f) and decreased apoptosis (Figure 2g). In contrast, miR-27a enrichment with miR-27am had the opposite effects (Supplementary Figures 1a–f). These data collectively demonstrate that miR-27a, via activation of PPARγ/β-catenin signaling, induces high glucose cultured podocytes to undergo mesenchymal transition and apoptosis, hence incurring podocyte injury.
To decipher whether PPARγ mediates Wnt/β-catenin signaling and podocyte injury in high glucose, we further examined the function of PPARγ using loss- and gain-of-function studies with PPARγ siRNA or its agonist rosiglitazone.27 Notably, we found that in high glucose condition, PPARγ siRNA decreased PPARγ phosphorylation, activated β-catenin and its target genes, snail1 and α-SMA, and downregulated podocin (red dashed box, Figure 3a). These changes, however, were not detected in normal glucose (green dashed box, Figure 3a). As shown by qRT-PCR analysis (Figure 3b), in high glucose, PPARγ depletion decreased the mRNA level of PPARγ and podocin, and increased β-catenin and its target genes, snail1 and α-SMA. These results suggest that a high glucose microenvironment is essential for PPARγ phosphorylation. Furthermore, PPARγ knockdown promoted the migration and invasion ability of high glucose cultured podocytes (Figures 3c and d) and increased apoptosis (Figure 3e). Conversely, PPARγ overexpression with rosiglitazone exerted the opposite effects (Supplementary Figures 2a–e). Taken together, these results indicate the pivotal role of PPARγ in mediating podocyte injury via activating Wnt/beta-catenin signaling in high glucose.
Having shown a reciprocal interplay between miR-27a and PPARγ, we next investigated whether the effect of miR-27a on downstream gene expression levels and podocyte functions depended on PPARγ. To this end, high glucose cultured podocytes were co-transfected with PPARγ siRNA and miR-27ai. As shown by immunofluorescence microscopy (Figure 3f), PPARγ siRNA-induced β-catenin activation, which was attenuated by co-transfection with miR-27ai. In this context, we hypothesized that PPARγ indispensably mediated β-catenin-induced downstream gene expression levels. This hypothesis was verified as exemplified by downregulation of β-catenin target genes, snail1 and α-SMA, and podocin upregulation by qRT-PCR (Figure 3g) and western blot analyses (Figure 3h). In addition, the increased ability of migration (Figure 3i) and invasion (Figure 3j) induced by PPARγ abolishment were mitigated by co-transfection with miR-27ai. Collectively, these results suggest that PPARγ is required in miR-27a-induced β-catenin activation and podocyte injury in high glucose.
We finally investigated the in vivo relevance of the interplay between miR-27a and PPARγ/β-catenin signaling. MiR-27ai or miR-27am were administered to diabetic rats and the functional effects examined. We found that miR-27ai significantly improved renal function as evidenced by decreased level of serum creatinine (Scr), serum blood urea nitrogen (BUN), urine albumin excretion rate (UAER), urine albumin to creatinine ratio (UACR) and increased creatinine clearance rate (Ccr) (Table 1). Conversely, miR-27am exerted opposite effects (Table 2). To confirm the localization of miR-27a in glomeruli, we next examined miR-27a expression using in situ hybridization (ISH). Compared with normal control rats, miR-27a was upregulated in podocytes of diabetic rat kidney tissues, as well as in renal tubular epithelial cells, which was significantly abolished by miR-27ai and enriched by miR-27am (Figures 4a and b). Furthermore, miR-27ai reduced PPARγ phosphorylation and β-catenin activation in podocytes. Concomitantly, the expression of β-catenin target genes snail1 and α-SMA were downregulated and podocyte-specific marker synaptopodin was markedly upregulated. MiR-27am, however, had the opposite effects (Figures 4c and d). Further, elevated miR-27a level was found in microdissected glomeruli (Figure 4e) and plasma samples (Figure 4f) of diabetic rats, which was decreased by treatment with miR-27ai and increased by miR-27am. These data further illustrate the functional interplay between miR-27a and PPARγ/β-catenin signaling in diabetic rats in vivo.
Given that miR-27a induces abolishment of slit diaphragm-associated protein synaptopodin, we next examined whether miR-27a affected podocyte number and architectural integrity. We found that in diabetic glomeruli, podocyte architectural integrity was disrupted as evidenced by a decrease of the podocyte marker synaptopodin, loss of WT1-positive podocytes and effaced podocyte foot processes. These changes were mitigated by miR-27ai and exaggerated by miR-27am (Figures 5a–f). Next, we asked whether miR-27a-dictated podocyte injury depended upon the interplay between PPARγ and β-catenin activation. To this end, we carried out immunofluorescence microscopy and co-immunoprecipitation analyses to further explore their correlations. The results showed that phosphorylated PPARγ and active β-catenin colocalized and interacted with each other (Figure 5g), which was strengthened both in diabetic conditions and by miR-27am treatment (Figure 5h). These results together suggest that miR-27a induces podocyte injury via the PPARγ/β-catenin signaling.
We verified the expression patterns of miR-27a and PPARγ/β-catenin-related markers in human renal biopsy samples from DN patients. Similar to changes detected in animal studies, compared with healthy transplant donor kidney biopsies, ISH analysis shows that miR-27a level was upregulated in podocytes of renal biopsies from DN patients (Figures 6a and b). Concomitantly, activation of phosphorylated PPARγ and β-catenin were observed in podocytes of renal tissues from DN patients. Consistently, β-catenin activation also led to increased expression of snail1 and α-SMA but decreased synaptopodin (Figures 6c and d). In addition, podocyte injury was demonstrated by loss of the slit diaphragm-associated protein synaptopodin and decreased number of WT1-positive podocytes as detected by immunofluorescence microscopy (Figures 6e–g). Colocalization of phosphorylated PPARγ and active β-catenin was also identified in podocytes of DN samples (Figures 6h–j), further supporting the notion that PPARγ-mediated β-catenin activation promotes podocyte injury in DN. A hypothetical model illustrated that miR-27a, via PPARγ-mediated β-catenin activation, promotes podocyte injury in DN (Figure 6k).
Podocytes, the terminally differentiated visceral epithelial cells, uniquely reside along the glomerular basement membrane.28, 29 Podocyte foot processes form one of the most critical components of glomerular filtration barrier and keep the integrity of the glomerulus.30 Pathologically, multiple injurious stimuli have been found to contribute to podocyte dedifferentiation, featured by loss of podocyte-specific properties and gain of mesenchymal features, thus leading to podocyte dysfunction and defects of glomerular filtration barrier.31 Diabetes is a condition in which podocytes are universally exposed to high glucose and its resulting metabolites, such as advanced glycation end products (AGEs) and oxidative stress.23, 32, 33
Several miRNAs, such as microRNA-130b,34 microRNA-182,35 microRNA-146a36 and microRNA-200a,37 have been found dysregulated in diabetes. They function as a ubiquitous class of noncoding RNA modulators of mammalian physiological responses acting via posttranscriptional regulation of gene expression.38 In this study, we have identified that high glucose induces the expression of miR-27a in cultured podocytes. In addition, miR-27a upregulation has also been detected in animal models of diabetes and DN patients. This finding highlights the significance of miR-27a in podocyte pathophysiology in DN.
The most novel finding in this study is that in DN miR-27a induces podocyte injuries and worsens renal function via PPARγ-mediated β-catenin activation. This conclusion is authenticated by several lines of evidence. First, miR-27a directly targets the 3′-UTR of PPARγ mRNA and provokes the PPARγ-mediated downstream events (Figure 1). Second, PPARγ phosphorylation causes β-catenin activation, which triggers a series of β-catenin-dependent reprogramming in podocytes, such as enhanced epithelial–mysenchymal transition, loss of podocyte-specific markers and increased apoptosis (Figures 2 and and3).3). Third, miR-27a disrupts podocyte architectural integrity and reduces podocyte number in diabetic rats. Finally, miR-27a deteriorates renal function as revealed by elevated albuminuria and dropped Ccr in diabetic rats. The finding that miR-27a targets PPARγ and controls the expression of β-catenin target genes provides novel and mechanistic insights into how miRNAs contribute to the development and progression of DN. This notion inspires us to develop new combinational therapeutic strategies to simultaneously regulate miR-27a downstream signaling, such as PPARγ and β-catenin.
Another intriguing finding in this study is that miR-27a-induced β-catenin activation depends on PPARγ phosphorylation in DN. We also found that miR-27ai increased PPARγ transcriptional activity but PPARγ protein level remained unchanged in high glucose. This observation led us to reason it is likely that inhibiting PPARγ phosphorylation is an alternative mechanism for anti-diabetic effects of PPARγ ligands. Indeed, this speculation has been supported by emerging lines of evidence showing that many PPARγ-based novel drugs, such as SR166439 and UHC1,40 have a separate biochemical activity, blocking the obesity-linked phosphorylation of PPARγ by Cdk5.20 These novel synthetic compounds have a unique mode of binding to PPARγ while lacking classical transcriptional agonism, and hence with fewer side effects such as fluid retention, bone fracture, weight gain or congestive heart failure.41 We believe that miR-27ai has promising efficacies in inhibiting PPARγ phosphorylation similar to the above mentioned new synthetic compounds. More broadly, the potential of ‘partial agonists' to modulate protein phosphorylation may be feasible, possibly allowing for identification of novel miR-27a targeted drugs. Exactly how miR-27a affects PPARγ phosphorylation is unknown at this stage and warrants additional investigation.
The therapeutic efficacy of miR-27a blockage by its inhibitors results in reversal of the mesenchymal transition and architectural defects of the podocyte. It also combats proteinuria and renal injury in diabetic rats (Figure 4,Tables 1 and and2).2). This finding is in line with the critical role of β-catenin inactivation in a series of studies.42, 43 It is also consistent with our previous study illustrating the inhibitory effect of miR-27a/PPARγ axis in renal tubulointerstitial fibrosis in DN.44 In glomerular mesangial cells, miR-27a has been reported to induce progression of DN by targeting PPARγ.26 The similarities and differences between our present study and the above studies are as follows: (1) by targeting PPARγ, miR-27a could initiate each individual event provoking different downstream signalings in different cell types in diabetic kidney, which may have amplifying effects in exacerbating the disease; (2) miR-27a could contribute to the formation of the most typical pathological hallmark of diabetic glomerular lesions, for example, extracellular matrix accumulation and podocyte foot process effacement; and (3) miR-27a is a robust biomarker in DN, which may be used for evaluating the disease severity. It should also be noted that PPARγ and β-catenin are widely expressed in the kidney, so the effects of systemic administration of miR-27ai or miR-27am may affect multiple cell types. Therefore, it raises the topic of combinational therapy in DN in that by synergistically targeting miR-27a/PPARγ/β-catenin and miR-27a/PPARγ/TGF-β signalings, both podocyte injuries and tubulointerstitial fibrosis could be reversed or even prevented. It is also possible that by targeting miR-27a alone, β-catenin and TGF-β45, 46, 47, 48 downstream events may be inhibited. Future studies will be needed to explore the efficacies of miR-27ai in clinical settings.
Our study is limited in that we did not tailor the treatment strategy according to the disease stage in the animal studies. Future studies will be needed to explore how to select the appropriate time point for using single/combinational therapies, for example, miR-27ai at the early stage and inhibitors of β-catenin and TGF-β in later stages. In addition, whether cross-talks between glomerular and tubular cells or cross-talks between podocytes and mesangial cells initiate miR-27a-mediated downstream signalings remains unknown at these stage. These areas warrant further additional exploration.
The results in this study, for the first time, demonstrate that miR-27a/PPARγ axis, as an upstream regulatory signaling, dictates the expression of genes associated with podocyte biology via β-catenin pathway. Thus, miR-27a/PPARγ and β-catenin are intimately connected to constitute a pathologic axis having a crucial role in the pathogenesis of DN.
In summary, we have shown that miR-27a, via activating PPARγ/β-catenin signaling, exacerbates podocyte injury as evidenced by increased capacity of migration, invasion and apoptosis. In addition, we have identified that miR-27ai is effective in reversing proteinuria and renal dysfunction in DN. This study provides a mechanistic interplay between miR-27a and PPARγ/β-catenin activation in the pathogenesis of DN.
A conditionally immortalized murine podocyte cell line was maintained as described previously.49 Briefly, cells were cultured at 33°C in RPMI medium 1640 (Sigma-Aldrich, , St. Louis., MO, USA) supplemented with 10% fetal bovine serum (FBS; Life Technologies, Carlsbad, CA, USA) and 10units/ml mouse recombinant interferon-γ (IFN-γ, R&D Systems, Minneapolis, MN, USA). To induce differentiation, podocytes were grown under non-permissive conditions at 37°C in the absence of IFN-γ for 14 days before experiments.
Podocytes were then maintained in normal glucose (5mM) for 1 week, grown to 75–85% confluence and made quiescent by incubation overnight in a serum-free medium. Podocytes were next exposed to mannitol (30mM) or high glucose (30mM) for time periods as individual experiments required.
The predicted 3′-UTRs sequence of PPARγ interacting with miR-27a and mutated sequences within the predicted target sites were synthesized and inserted into the pRL-TK control vector (Promega, Madison, WI, USA). Podocytes transfected with 120ng miR-27ai or negative controls (NCs), followed by co-transfection with 30ng of the wild-type or mutant 3′-UTR of PPARγ using 0.45μl of Fugene (Promega). Luciferase assay was carried out on extracts from the cells 48h post transfection and measured using Dual-Luciferase Assay System (Promega). pRL-TK expressing Renilla luciferase was co-transfected as an internal control. Data were normalized by the ratio of Firefly and Renilla luciferase activities.
MiR-27ai, miR-27am or the appropriate NCs of miRNA inhibitor (miR-iNC) and miRNA mimics (miR-NC), respectively, were purchased from GenePharma (Shanghai, China) and transfected at a final concentration of 50–100nM in the cells using HiPerFect Transfection Reagent (Qiagen, Hilden, Germany) according to the manufacturer's recommendations. Expression of murine PPARγ was knocked down with small interfering RNA (siRNA) duplexes using Oligofectamine (Invitrogen, Carlsbad, CA, USA). The target sequence for PPARγ mRNA was: 5′-AAUAUGACCUGAAGCUCCAAGAAUAAG-3′. Non-targeting siRNA pool (D-001206-13-05; Dharmacon, Fisher Scientific, Pittsburgh, PA, USA) was used as a NC. Podocytes were transfected with 1μg of siRNA in reduced serum medium (OPTI-MEM-I; Invitrogen, Carlsbad, CA, USA) according to the manufacturer's protocol and harvested 72h post transfection. The RNA and protein were extracted and analyzed, respectively.
PPARγ agonist rosiglitazone (Rosi.; 50nM; R2408; Sigma-Aldrich) was used to treat podocytes. Cells in each group were treated for 72h and then harvested for further analyses.
Cells (1.0 × 106 cells/ml) in serum-free medium were added to the top chamber of 24-well transwell plates (8mm pore size; Corning Star, Cambridge, MA, USA) and 600μl of complete medium with 10% FBS was in the bottom chamber. The assembled chamber was incubated at 37°C in a humidified, 5% CO2 cell culture incubator for 24h and fixed with 10% formalin and stained with crystal violet for visualization.
Podocytes (2.0 × 105 cells per well) were plated in six-well plates and grown to 80% confluence. The individual wells were wounded by scratching with a pipette tip and incubated with serum-free medium to various time points. Wells were photographed under phase-contrast microscopy ( × 10).
All animal experiments were specifically approved by the Animal Ethics Committee at Nanfang Hospital, Southern Medical University, Guangzhou, China. The study protocols conform to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication no. 85-23, revised 1996).
Male Sprague Dawley rats, 6–8 weeks of age, were kept in the Animal Center of Nanfang Hospital according to the policy of the Committee for Animal Usage. Rat models of diabetes were induced with a single intraperitoneal injection of either streptozotocin (65mg/kg; S0130; Sigma-Aldrich) in 0.1M citrate buffer (pH 4.5) or 0.1M citrate buffer according to the protocol described previously.34, 50, 51 Rats with a blood glucose level over 16.7mmol/l were considered diabetic and thus included in the study, followed by treatment with the intraperitoneal administration of miR-27ai or miR-27am (4ng/mm3) or the appropriate scrambled control RNAs every day till killing at week 12 of diabetes.
Six groups of rats were used: (1) normal control (NC, n=6), (2) diabetic control (DM, n=7), (3) diabetic rats treated with NC miRNA inhibitor (DM_miR-iNC, n=8), (4) diabetic rats treated with miR-27ai (DM_miR-27ai, n=7), (5) diabetic rats treated with NC miRNA mimic (DM_miR-NC, n=6) and (6) diabetic rats treated with miR-27a mimic (DM_miR-27am, n=6). No adverse or toxic effects were observed.
Blood glucose level was measured every week. The treatment continued until the rats were killed. At 12 weeks after the induction of diabetes, rats were housed in metabolic cages and urine was collected for the determination of UAER and UACR. Then, rats were anesthetized with pentobarbital sodium (P3761, 30mg/kg, Sigma-Aldrich) and the venous blood was drawn from the eye orbit of rats before killing. Left kidneys were fixed by retrograde aortic perfusion using phosphate-buffered saline (PBS, pH 7.4). Cortex of the left kidneys was obtained for electron microscopy analysis and the rest was fixed in 10% formalin in PBS for 24h, rinsed in buffer, dehydrated and embedded in paraffin for histological analysis. The non-perfused right kidney was cut into small pieces, snap-frozen and stored at −80°C for further analysis. Rats were killed by exsanguination without any previous intervention following guidelines recommended from the Animal Research: Reporting In Vivo Experiments (ARRIVE) (http://www.nc3rs.org.uk/page.asp?id=1357). All efforts were made to minimize suffering.
Blood glucose, BUN, serum and urine creatinine were analyzed using a Beckman Coulter AU480 Chemistry Analyzer (Beckman, Brea, CA, USA). Urine albumin was determined with an enzyme-linked immunosorbent assay kit specific for rat albumin (E111-125, Bethyl Laboratories, Montgomery, TX, USA). Ccr was calculated as urinary creatinine (umol/l) × urine volume (ml/min)/Scr (umol/l), and was expressed as ml/min/kg. All experiments were repeated in triplicate.
Several 1-mm cubes from cortex of left kidneys were cut, placed in 2.5% glutaraldehyde for at least 4h washed with cacodylate buffer, postfixed in 1% osmium tetroxide and block-stained in uranyl acetate before embedding in Poly/Bed812 resin (Polysciences, Inc., Warrington, PA, USA). Ultrathin sections were obtained from at least three randomly selected glomeruli from each animal, stained with uranyl acetate and lead citrate and examined using a Hitachi 7,700 transmission electron microscope (Tokyo, Japan). Representative micrographs of the podocytes from each rat were examined and imaged at a magnification of × 4000.
Fresh snap-frozen rat kidney tissues were embedded in optimal cutting temperature (OCT) compound (Sakura Finetek Japan Co., Ltd, Tokyo, Japan). Ten-μm thick cryosections were cut and mounted on Membrane slides (Muster Membrane Slides 1.0 polyethylene naphthalate (PEN)-covered glass slide; 000757-11, Zeiss, Jena, Germany). On each unstained membrane slide, 50–100 individual glomeruli were laser microdissected and collected using the PALM MicroBeam LCM system (Zeiss) for RNA and protein extraction as described previously.50
Total RNA from podocytes and microdissected glomeruli were extracted using TRIzol reagent (MRC, Cincinnati, OH, USA). First-strand cDNA was synthesized using 2μg of total RNA treated with Moloney murine leukemia virus reverse transcriptase (Promega) according to the manufacturer's instructions. qRT-PCR analysis was performed in triplicate with Power PCR SYBR Green Master Mix (Applied Biosystems, Carlsbad, CA, USA) using the ABI PRISM 7500 FAST Real-TIME PCR System (Applied Biosystems) with results normalized to β-actin expression. The ΔΔCT method was used to calculate relative expression. Primer sequences used in qRT-PCR are shown in Supplementary Tables 1 and 2.
To assess the level of miRNA expression, total RNA extracted from the cells, human plasma samples, rat glomerular tissues or plasma samples was reversely transcribed into cDNA using miRScript PCR System and then analyzed by qRT-PCR with the miScript SYBR Green PCR Kit using the specific miR-27a miScript Primer Assays (Qiagen) according to the manufacturers' instructions. Expression levels were normalized to the average of U6-snuRNA. MiR-27a levels were calculated as fold change (2−ΔΔCT) with respect to normal controls. The mean value of miR-27a expression in glucose-free cultured cells was used as the calibrator. Target-specific reverse transcription and Taqman miRNA assays were performed using the Hairpin-it miRNA qPCR Quantitation Kit (GenePharma, Suzhou, China) according to the protocol. The reactions were performed using the ABI PRISM 7500 FAST Real-TIME PCR System (Applied Biosystems) with results normalized to U6-snuRNA expression. The 2−ΔΔCt method was used to calculate the relative expression. All experiments were performed in triplicate.
Lysates from podocytes and microdissected glomeruli from each experimental group were separated in parallel on two 10% denaturing sodium dodecyl sulfate-polyacrylamide gels, transferred onto nitrocellulose membranes, blocked with 5% nonfat milk in 0.1% tris-buffered saline with Tween-20 (TBST) and probed using primary antibodies against PPARγ (phospho S112) (1:100, Santa Cruz Biotechnology, Santa Cruz, CA, USA), PPARγ (1:100, ab45036, Abcam, Cambridge, UK), non-phospho (active) β-catenin (Ser45) (D2U8Y, 1:50, #19807, Cell Signaling Technology, Beverly, MA, USA), active β-catenin (05-665; EMD Millipore, Billerica, MA, USA), snail1 (1:100, ab53519, Abcam), α-SMA (1:100, ab5694, Abcam), NPHS2 (1:100, ab50339, Abcam), synaptopodin (1:50, 39067, GeneTex, San Antonio, TX, USA) and β-actin (1:200, ab6276, Abcam) at 4°C overnight. After extensive washing in TBST buffer, the secondary antibody (horseradish peroxidase-labeled IgG anti-rabbit/mouse antibody, Invitrogen, Cambridge, MA, USA) was used at 1:3000 dilution for 1h at room temperature. The supersignal-enhanced chemoluminescent substrate (Pierce Biotechnology, Inc., Rockford, IL, USA) was applied to the probed membrane and exposed for 10min before the protein bands were visualized on radiograph films (Super Rx, Fuji Photo Film, Tokyo). Quantification was performed by measurement of the intensity of the bands using ImageJ analysis software (National Institute of Health, Bethesda, MD, USA).
A total of 62 formalin-fixed paraffin-embedded renal biopsy samples were obtained from type 2 diabetic patients including 46 from the Department of Renal Pathology in King Medical Diagnostics Center in Guangzhou from 2012 to 2014 and 16 from the Division of Nephrology in the First Affiliated Hospital of Inner Mongolia Medical University in Hohhot from 2010 to 2015. The inclusion criteria were: (1) type 2 diabetic patients with no history of using renal toxic or herbal medicine; (2) the indications for performing the renal biopsy were proteinuria with or without microscopic hematuria and fast drop in renal function; (3) type 2 diabetic patients with no complications of other renal diseases. The Ethics Committee from King Medical Diagnostics Center and Inner Mongolia Medical University specifically approved the use of patient tissue samples in this study and written informed consent was obtained from each patient.
In all specimens, the morphological diagnosis of DN was confirmed by two individual renal pathologists (JG and XB). Normal kidney tissues (n=13) were obtained from healthy transplant donor kidney biopsies and thus served as controls. Serum samples from 30 DN patients and 30 healthy volunteers were collected.
ISH for miR-27a (50nM) was performed as described previously.52 Frozen kidney samples from rats were fixed in 4% paraformaldehyde in 0.1M PBS (pH 7.4) containing 1/1000 diethylprocarbonate overnight at 4°C before staining. Samples were then placed in 1mol/l sucrose solution overnight at 4°C and embedded in OCT compound at −80°C. Ten-μm-thick cryosections were prepared and the ISH analysis was performed using digoxygenin (DIG)-labeled miRCURY Locked Nucleic Acid (LNA) microRNA Detection Probes (Exiqon, Vedbaek, Denmark) according to the manufacturer's protocol. Briefly, slides were pre-digested with proteinase-K (15μg/ml) at 37°C for 8min, pre-hybridized at 57°C for 15min, hybridized with double-carboxyfluorescein (FAM) labeled LNA probes (Exiqon A/S) diluted in hybridization buffer (PerfectHyTM Plus Hybridization Buffer; Sigma-Aldrich) at 56°C in a humidified chamber for 2h in a humidified chamber. After stringent washes with saline-sodium-citrate buffer (pH 7.0), the probes were detected with alkaline phosphatase-conjugated sheep anti-FAM Fab fragments, followed by incubation in 4-nitroblue tetrazolium and 5-bromo-4-chloro-3′-indolylphosphate (NBT/BCIP) solution (Roche, Basel, Switzerland) for 60min. The signal resulted in a dark blue staining. Photographs were taken under a BX-51 light microscope (Olympus, Tokyo, Japan) fitted with a DFC550 color video camera (Leica Microsystems, Wetzlar, Germany) using appropriate filters.
Podocytes, tissue samples from rats and patients were labeled with antibodies to pPPARγ (1:100), active beta-catenin (1:100), snail1 (1:100), α-SMA (1:100), synaptopodin (1:1000) and WT1 (1:100). For immunofluorescence staining, Alexa Fluor 594-conjugated goat anti-mouse IgG and Alexa Fluor 488-conjugated goat anti-rabbit IgG (1:1000, Invitrogen, Cambridge, MA, USA) were used for secondary antibodies, nuclei were counterstained with 4′,6-diamidino-2-phenylindole (DAPI, Sigma-Aldrich) and coverslipped with aqueous mounting medium (CTS011, BD Bioscience, Minneapolis, MN, USA). For immunohistochemistry, EnVision Detection Systems Peroxidase/diaminobenzidine (DAB), rabbit/mouse kit (K4065, Dako, Carpinteria, CA, USA) was used. Nuclei were counterstained with hematoxylin and coverslipped with Permount mounting medium (00-4960-56, eBioscience, San Diego, CA, USA).
Samples were evaluated semiquantitatively by systematically selecting without bias 20 fields for analysis. Immunofluorescence images were taken with a FV1000-IX81 confocal laser scanning microscope (Olympus) with appropriate filters. Staining intensity was measured using ImageJ analysis software (ImageJ 1.44, National Institute of Health). The number of cells positive for WT-1 was counted in all glomeruli at × 400 magnification in each section and the mean number was recorded as podocyte number in each sample. Immunohistochemical images were taken under a BX-51 light microscope (Olympus) fitted with a DFC550 color video camera (Leica Microsystems) using appropriate filters. PBS instead of primary antibodies served as a NC.
Data are presented as mean±S.D. Statistical significance between groups was evaluated using independent samples T-test or one-way ANOVA followed by post-hoc least-significant difference test. All statistical tests were performed using SPSS 12.0 (SPSS, Inc., Chicago, IL, USA). The significance level is set at 0.05 to indicate statistical significance.
This study was supported by National Nature and Science Young Investigator Grant (no. 81100496) from the National Natural Science Foundation of China, Special Fund from Chinese Society of Nephrology (no. 13030370422), Guangdong Natural Science Foundation (no. 2016A030313581) and Distinguished Young Scholar Fund from Nanfang Hospital (no. 2015J009) to XB. We thank Guangzhou King Medical Diagnostics Center for providing human renal biopsy samples. We gratefully acknowledge all lab members for their technical assistance.
Supplementary Information accompanies this paper on Cell Death and Disease website (http://www.nature.com/cddis)
Edited by G Calin
The authors declare no conflict of interest.