Persistent Insulin Resistance in Podocytes Caused by Epigenetic Changes of SHP-1 in Diabetes

Diabetic kidney disease is one of the most devastating microvascular complications of diabetes that can lead to end-stage renal disease (1). Hyperglycemia and insulin resistance are believed to be the major risk factors for the development of diabetic nephropathy (DN). The Diabetes Control and Complications Trial (DCCT) demonstrated that intensive blood glucose control reduces the risk of diabetic microvascular complications (nephropathy, retinopathy, and neuropathy) compared with conventional insulin treatment (2). Moreover, the follow-up Epidemiology of Diabetes Interventions and Complications (EDIC) study showed that the cumulative incidence of nephropathy continues to differ between intensive and standard treatment groups for many years despite comparable HbA_1c (3,4). To explain this phenomenon, a new concept named hyperglycemic memory has been suggested to explain the persistence of hyperglycemia-induced metabolic derangements after normalization of blood glucose (5).

Podocytes are highly specialized epithelial cells essential for the integrity of the glomerular filtration barrier (6). Podocyte cell death is an early event of DN (7), and decreased glomerular podocyte density is the strongest predictor of DN progression (8). Insulin action itself is important in podocyte biology. Renal disease similar to DN can be observed in patients with a genetic mutation of insulin receptor (IR) signaling (9,10), and several studies have supported a link between insulin resistance and renal pathology in type 1 diabetes. In a clinical study, 14 patients with type 1 diabetes and microalbuminuria had significantly lower glucose disposal during euglycemic-hyperinsulinemic clamp studies than matched control subjects, even after accounting for blood pressure and BMI (11). Moreover, Welsh et al. (12) uncovered the importance of insulin action in podocytes, showing that mice depleted of the IR specifically in podocytes developed albuminuria and podocyte loss, which are characteristic of DN pathology. However, the mechanisms of podocyte and kidney insulin unresponsiveness induced by the hyperglycemic memory effect have not been explored.

The regulation of protein tyrosine phosphatase, such as SHP-1, is an important mechanism for developmental control and homeostasis of an array of cellular processes, including cell growth, differentiation, and oncogenic transformation. SHP-1 is a cytosolic tyrosine phosphatase expressed primarily in hematopoietic and epithelial cells and has been shown to dephosphorylate a wide spectrum of phosphoproteins involved in cell signaling of the receptors of the tyrosine kinase family. SHP-1 knockout mice are markedly glucose tolerant and insulin sensitive compared with wild-type controls. Dubois et al. (13) showed that these observations are a result of the enhanced insulin signaling through the IR substrate 1/2 (IRS1/2)/PI3K/Akt axis in the liver. Our group reported that in the kidney, elevated SHP-1 expression levels in mouse and human podocytes exposed to high-glucose (HG) concentrations result in reduced insulin and nephrin actions and podocyte dysfunction (14,15).

Current efforts have focused on the implication of epigenetic modifications as a potential mechanism underlying glycemic memory effects (16). Patients with albuminuria and microalbuminuria have shown higher levels of global DNA methylation than control subjects (17). More specifically in DN, Natarajan’s group (18) showed that the activation mark H3K9/14ac, generally associated with transcriptionally active genome regions, is increased, whereas H3K9me2, H3K9me3, and H4K27me3, which are associated with repressed regions, are decreased in diabetic rodents. With consideration of the hyperglycemic memory effect as the major obstacle for optimal prevention and treatment of DN, we hypothesized that epigenetic changes cause persistent SHP-1 expression after glycemic control, resulting in sustained insulin signaling inhibition and DN progression.

Primary antibodies for immunoblotting and immunochemistry were obtained from commercial sources, including actin (horseradish peroxidase; I-19), SHP-1 (C-19), SHP-1 (D11; immunofluorescence), IR-β (C-19), phosphotyrosine (PY99), phosphatase and tensin homolog (PTEN; C-20-R), transforming growth factor-β (TGF-β; SC-146), protein kinase C-δ (PKC-δ; C-20), cadherin (C-19), WT-1 (C-19), and rabbit and mouse peroxidase-conjugated secondary antibody from Santa Cruz Biotechnology; protein kinase B (Akt), extracellular signal-regulated kinase (ERK), phospho-Akt (D9E), and phospho-ERK (D13.14.4E) from Cell Signaling; purified monoclonal antibody against PTP1B (610139) and SHP-2 (610621) from BD Biosciences; monoclonal antibody against IR-β (AB69508) from Abcam; nephrin GP-N2 from PROGEN Biotechnik; podocin (P-0372) from Sigma; collagen type IV from Novus Biological; Alexa Fluor 546–conjugated anti-rabbit and Alexa Fluor 488–conjugated anti-guinea pig from Jackson ImmunoResearch; and Alexa Fluor 594–conjugated anti-mouse from Invitrogen. All other reagents used, including RPMI medium, EDTA, leupeptin, phenylmethylsulfonyl fluoride, aprotinin, insulin, d-glucose, d-mannitol, fluorescein isothiocyanate-inulin, sodium orthovanadate, and actinomycin D, were purchased from Sigma-Aldrich; FBS and penicillin-streptomycin were obtained from Invitrogen.

C57BL/6J (Ins2^+/+) and diabetic heterozygous male Ins2^+/C96Y (Akita) mice were purchased from The Jackson Laboratory and bred in our animal facility. Throughout the study period, animals were provided with free access to water and standard rodent chow (Harlan Teklad). All experiments were conducted in accordance with the Canadian Council of Animal Care and Institutional Guidelines and were approved by the Animal Care and Use Committees of the University of Sherbrooke according to National Institutes of Health guidelines. After 4 months of uncontrolled diabetes, animals received two insulin implants (Linbit; LinShin Canada) subcutaneously per month to normalize blood glucose levels, and these implants were replaced every month over a 2-month period (Ins2^+/C96Y + ins). Blood glucose levels and animal weight were recorded every week.

Blood glucose was measured by a glucometer (Contour; Bayer). Twenty-four–hour urine collections were obtained from mice 1 day before death by housing them in individual mouse metabolic cages (Nalgene Nunc International) with free access to water and rodent mash. Urinary creatinine concentration was measured using alkaline picrate colorimetry based on Jaffe reaction (The Creatinine Companion; Exocell), and albumin levels were measured by using an indirect competitive ELISA according to the manufacturer’s instructions (Albuwell M; Exocell). The glomerular filtration rate (GFR) was evaluated by using fluorescein isothiocyanate-inulin clearance as described previously (19).

Right mouse kidneys were harvested for pathology examination, and sections were fixed in 4% paraformaldehyde (Sigma-Aldrich) and then transferred to 70% ethanol for immunohistochemistry. The tissue was embedded in paraffin, and 4-μm sections were stained with periodic acid Schiff stain and hematoxylin-eosin (Sigma-Aldrich). Left mouse kidneys were used for transmission and scanning electron microscopy to evaluate podocyte structure and foot process effacement as previously described (14).

Immunohistochemistry of kidney sections was performed with the ABC Kit from Vector Laboratories according to the manufacturer’s protocol. Coloration was obtained by incubating sections in 3,3′-diaminobenzidine solution (Vector Laboratories). Counterstaining of the nucleus was done using Gill’s Hematoxylin (Vector Laboratories).

Mesangial matrix expansion and glomerular hypertrophy were evaluated quantitatively as previously described (14). Apoptotic nuclei of kidney sections were detected using the TACS 2 Tdt-Fluor In Situ Apoptosis Detection Kit (Trevigen) according to the manufacturer’s instructions as previously described (14). Apoptotic podocytes were counted when both podocyte markers and Tdt-fluorescein–positive cells colocalized on all glomeruli (20–30) on the transverse section of the left kidney.

Kidneys from Ins2^+/+, Ins2^+/C96Y, and Ins2^+/C96Y mice treated with insulin implants were frozen in optimal cutting temperature compound (BDH Chemicals) embedding resin in cryomolds on a block of dry ice and sectioned at 8 μm (Leica cryostat). Immunofluorescence of the sections using nephrin, podocin (1:200), phospho-Akt, and SHP-1 antibodies were performed as previously described (14). Sections were examined with a Plan Apochromat 60× oil immersion objective (numerical aperture 1.42) mounted on an inverted spectral scanning confocal microscope (FV1000; Olympus, Tokyo, Japan) as previously described (15). Images of one experiment were taken at the same time under identical settings and handled similarly in Adobe Photoshop across all images.

The insulin signaling pathway was evaluated by injecting insulin 5 mU/g i.v. in Ins2^+/+, Ins2^+/C96Y, and Ins2^+/C96Y mice treated with insulin implants. The right kidney was removed before insulin injection and served as an internal control (nonstimulated). Fifteen minutes after the insulin injection, the left kidney was removed for protein extraction.

The mouse podocyte cell line was used and cultured as previously described (14,20). After differentiation of podocytes, the medium was changed to RPMI 0.1% FBS containing normal glucose (NG) (5.6 mmol/L + 19.4 mmol/L mannitol to adjust osmotic pressure) or HG (25 mmol/L) up to 120 h. To mimic the reversibility phenomenon, podocytes were exposed to HG for 96 h and then NG for the last 24 h (HG + NG). Adenoviral vectors containing green fluorescent protein (Ad-GFP) and dominant-negative SHP-1 (Ad-dnSHP-1) were used to infect podocytes as reported previously (14).

Immunoprecipitation assays were performed as described previously (14). For immunoblot analysis, 10–50 μg of lysate proteins were separated by SDS-PAGE and then transferred to a polyvinylidene fluoride membrane, which was blocked with 5% skim milk. Proteins were identified by enhanced chemiluminescence (Pierce Thermo Fisher Scientific).

Real-time PCR was performed to evaluate mRNA expression of SHP-1, SHP-2, PTP1B, PKC-δ, IRS-1, IRS-2, IR-β, collagen type IV, TGF-β, and fibronectin as previously described (14). PCR primers are listed in Supplementary Table 1. GAPDH mRNA expression was used for normalization.

Caspase-3 and -7 enzymatic activities were determined by quantification of cleaved substrate using luminescent assay Caspase-Glo 3/7 (Promega) as previously described (14).

We assessed phosphatase activity of SHP-1 by using a tyrosine phosphatase assay system (V2471; Promega) according to the manufacturer’s instructions. SHP-1 was immunoprecipitated from cell lysates with a polyclonal antibody (C-19) prebound to Protein A Sepharose beads. The phosphatase activity was determined using the Infinite 200 PRO NanoQuant (Tecan Group).

Renal cortexes of three kidneys (one per mouse) were combined and minced, and all tissue was passed through a 200-μm, 150-μm, and 75-μm sieve. The glomeruli that remained at the top of 75-μm sieve were collected with PBS and centrifuge for 10 min at 500g. The glomeruli samples (three kidneys per sample, four samples per group) were then used for chromatin immunoprecipitation (ChIP) assay.

At the end of each treatment, ChIP assays were performed as described by Kuo and Allis (21), with minor modifications as previously described (22). At the end of treatment, podocytes or glomeruli were cross-linked by adding formaldehyde 1.1% for 15 min at room temperature and glycine 1 mol/L for the last 5 min. Cells and glomeruli were then washed two times in PBS, and the pellet was resuspended in 200 μL of ChIP lysis buffer (1% SDS, 10 mmol/L EDTA, 50 mmol/L Tris-HCl [pH 8.0], and protease inhibitors) and sonicated with a Branson Sonifier 450 (Danbury, CT) at power setting 2 with 10-s pulses at duty cycle 90 (to obtain fragments <500 base pairs). The chromatin solution was diluted 10-fold in ChIP dilution buffer (0.01% SDS, 1.1% Triton X-100, 1.2 mmol/L EDTA, 16.7 mmol/L Tris [pH 8.1], 16.7 mmol/L NaCl, and protease inhibitors). One-tenth of the lysate was used for purification of total DNA. Each sample was precleared by incubating with 80 μL salmon sperm DNA/protein A-agarose 50% gel slurry (Upstate Biotechnology, Lake Placid, NY) for 30 min at 4°C. An aliquot of 5 μg of H3K4me1, H3K4me3, or H3K4ac (Abcam) were added and immunoprecipitated at 4°C overnight. The immunoprecipitate was collected using salmon sperm DNA/protein A-agarose and washed once with the following buffers in sequence: low-salt wash buffer (0.1% SDS, 1% Triton X-100, 2 mmol/L EDTA, 20 mmol/L Tris-HCl [pH 8.1], 150 mmol/L NaCl), high-salt wash buffer (0.1% SDS, 1% Triton X-100, 2 mmol/L EDTA, 20 mmol/L Tris-HCl [pH 8.1], 500 mmol/L NaCl); LiCl wash buffer (0.25 mol/L LiCl, 1% IGEPAL CA-630, 1% sodium deoxycholate, 1 mmol/L EDTA, 10 mmol/L Tris-HCl [pH 8.1]); and TE buffer (10 mmol/L Tris-HCl [pH 8.0], 1 mmol/L EDTA). Beads were incubated with 1% SDS/0.1 mol/L NaHCO₃ solution for 30 min at 65°C to isolate the DNA. DNA-histone cross-links were reversed by incubation at 65°C overnight followed by proteinase K treatment. DNA was recovered by purification with the QIAquick PCR Purification Kit (QIAGEN). Immunoprecipitated DNA was measured by quantitative PCR. Total DNA was used as reference (input). Primers for the open reading frame of the gene were used as shown in Supplementary Fig. 3 as a control to demonstrate specificity of amplification of DNA associated with immunoprecipitated chromatin.

Data are presented as mean ± SD for each group. Statistical analysis was performed by unpaired t test or by one-way ANOVA followed by Tukey test correction for multiple comparisons. Data in each group were checked for normal distribution using D’Agostino and Pearson normality test based on α = 0.05. All results were considered statistically significant at P < 0.05.

We and others have previously shown that the genetically modified type 1 diabetic model (Ins2^+/C96Y) exhibited renal dysfunction (elevated urine albumin-to-creatinine ratio and GFR) and pathology (glomerular hypertrophy and mesangial cell expansion) compared with control littermates (14,23). As expected, diabetic Ins2^+/C96Y mice exhibited an elevated urine albumin-to-creatinine ratio by 3.3-fold (P < 0.0001) and GFR (0.206 vs. 0.395 mL/min, P < 0.0001) compared with nondiabetic Ins2^+/+ mice (Fig. 1A and B). To evaluate the hyperglycemic memory phenomenon, diabetic mice were treated with insulin implants for 2 months. Mice in the interventional glycemic group gained weight and normalized glucose levels during these last 2 months (Supplementary Table 2). Of note, despite euglycemia for the last 2 months, albuminuria and GFR remained significantly higher in the diabetic mice treated with insulin implants compared with nondiabetic counterparts, suggesting that renal dysfunction persisted despite glucose normalization for 2 months after uncontrolled hyperglycemia for 4 months. Diabetic Ins2^+/C96Y mice demonstrated glomerular hypertrophy (P < 0.0001) (Fig. 1C and D) and expansion of the mesangium (P < 0.0001) (Fig. 1E and F) as well as increased collagen type IV– and TGF-β–positive staining compared with nondiabetic mice (Fig. 1G and H). Similarly, blinded assessment of glomerular hypertrophy (P < 0.0001), mesangium expansion (P < 0.0001), and collagen type IV and TGF-β expression remained elevated despite systemic glucose normalization with insulin implants for the last 2 months in diabetic mice (Fig. 1C–H).

Podocyte effacement and cell death are early markers of glomerular dysfunction in DN. Both transmission (Fig. 2A) and scanning (Fig. 2B) electron microscopy showed podocyte effacement (arrows and boxes) in diabetic Ins2^+/C96Y mice as well as in diabetic mice treated with insulin implants. Immunofluorescence analyses were performed to evaluate podocyte death. Increased positive apoptotic signal (TUNEL) in podocytes (WT-1 staining) was observed in diabetic Ins2^+/C96Y mice compared with nondiabetic Ins2^+/+ mice. Podocyte loss was also detected in diabetic mice treated with insulin implants for the last 2 months after 4 months of uncontrolled hyperglycemia. Quantitative PCR analysis demonstrated that mRNA expression of collagen type IV, TGF-β, and fibronectin, recognized markers of glomerular injury, were significantly increased in diabetic Ins2^+/C96Y mice compared with nondiabetic counterparts, and the normalized systemic glucose levels for the last 2 months with insulin implants did not reduce their expression (Fig. 3A–C).

To evaluate whether the inhibition of insulin actions persisted in the diabetic intervention group, insulin 5 mU/g i.v. was injected in all groups of mice. The right kidney was removed before the insulin injection and served as the internal control (nonstimulated). Fifteen minutes after the injection, insulin-stimulated Akt and ERK phosphorylation was significantly blunted in the renal cortex (Fig. 4A) and podocytes (using podocin as a podocyte marker) (Fig. 4B) of both nontreated and insulin implant–treated diabetic mice compared with Ins2^+/+ mice. The inhibition of the insulin signaling pathway in diabetes was not due to any change in the expression of IR-β, IRS1, or IRS2 (Fig. 4C–E).

Elevated SHP-1 expression has been previously reported in retinal and renal cortex using two different mouse models of type 1 diabetes (14,24,25). As expected, SHP-1 mRNA and protein expression were elevated in the renal cortex of Ins2^+/C96Y mice. Despite systemic euglycemia with insulin implants for the last 2 months, SHP-1 expression remained elevated in diabetic mice (Fig. 5A and B). We have shown that SHP-2 and PTP1B, other protein phosphatases that have been implicated in the insulin signaling pathway, were unchanged in renal cortex of diabetic mice and remained unaffected by treatment with insulin implants (Fig. 5A and B). Furthermore, with use of immunofluorescence analysis of nephrin, a podocyte-specific marker, we confirmed that the increase of SHP-1 expression continued to be elevated in podocytes of diabetic Ins2^+/C96Y mice treated with insulin implants (Fig. 5C).

Caspase-3/7 enzymatic activity, a marker of cell death, was measured in podocytes cultured in 5.6 mmol/L (NG) or 25 mmol/L (HG) glucose concentrations for up to 120 h. To replicate the hyperglycemic memory phenomenon, podocytes were exposed to HG for 96 h before returning glucose levels to 5.6 mmol/L (HG + NG). Exposing podocytes to HG levels for 120 h increased caspase-3/7 enzymatic activity by 32% (P = 0.0312) (Fig. 6A). Podocytes exposed to 96 h of HG and NG levels for the last 24 h showed similar caspase-3/7 enzymatic activity compared with 120 h of HG exposure, suggesting that the apoptotic signal continued to be active despite returning glucose concentration to a normal range. A longer period in NG after the HG treatment produced similar results. Of note, adding 10 nmol/L insulin for the last 24 h did not prevent HG effects on podocyte caspase-3/7 activity even when the 24-h insulin treatment was done in NG after the HG exposure (HG + NG) (Fig. 6A). These data suggest that HG-induced insulin unresponsiveness is not easily reversed.

To follow with our findings that podocyte cell death induced by HG concentrations was associated with increased SHP-1 expression (14), we evaluated whether the persistent caspase-3/7 enzymatic activity in podocytes exposed to NG after being cultured in HG concentrations displayed elevated SHP-1 expression. The data demonstrate that mRNA and protein expression as well as phosphatase activity of SHP-1 were significantly increased in cultured podocytes exposed to HG concentrations (Fig. 6B–D). No change was observed in the expression of SHP-2 and PTP1B. Similar to in vivo data, the mRNA and protein expression of SHP-1 as well as its phosphatase activity remained elevated despite returning podocytes to NG levels for 24 h (Fig. 6B–D) and 48 h (Supplementary Fig. 1A and B) after HG exposure. Importantly, the half-life of SHP-1 mRNA by treating podocytes with actinomycin D 5 mg/mL was similar in NG and HG conditions, suggesting that persistent expression of SHP-1 is not due to abnormal degradation or decay of SHP-1 in HG conditions (data not shown).

We previously reported that the increased expression SHP-1 was associated with the incapacity of insulin to induce Akt phosphorylation (14). The current results indicate that the persistent increased expression and activity of SHP-1 in podocytes reduced the capacity of insulin to phosphorylate its receptor (IR-β) and subsequently Akt (Fig. 7A). To demonstrate the continuous inhibition of IR activity, podocytes were infected with Ad-dnSHP-1. In podocytes overexpressing Ad-GFP, insulin-induced phosphorylation of IR-β was completely blunted during HG exposure. In contrast, the overexpression of Ad-dnSHP-1 rescued insulin action of IR-β phosphorylation (Fig. 7B).

The data demonstrate both in vivo and in vitro that SHP-1 was persistently expressed in type 1 diabetic mouse glomeruli and podocytes exposed to HG despite glucose level normalization. Our group has previously demonstrated that PKC-δ activation by HG levels is responsible for the increase expression of SHP-1 in podocytes (25). However, although PKC-δ activity was elevated, there was no significant change of PKC-δ mRNA and protein expression in podocytes exposed to HG (Supplementary Fig. 2). Thus, epigenetic regulation, such as histone modifications, could be an explanation for the persistent SHP-1 expression. Therefore, we performed ChIP assays of the SHP-1 promoter region with antibodies against histone 3 (H3) acetylation, monomethylation (me1), and trimethylation (me3). The data show that the promoter region of the SHP-1 gene was enriched with H3K9/14/27ac and H3K4me1 (Fig. 8A and B), whereas no change was observed in H3K4me3 (Fig. 8C). Moreover, we evaluated whether H3K9/14 and H3K4me1 enrichment in cultured podocytes occurred in vivo. Isolation of glomeruli from each group of mice demonstrated that H3K9/14ac and H3K4me1 marks were enhanced in diabetic glomeruli and remained elevated in glomeruli of diabetic mice treated with insulin implants (Fig. 8D and E). These data strongly support the notion that persistent expression of SHP-1 causing permanent insulin inhibition and dysfunction in podocytes was triggered by epigenetic modifications at the SHP-1 promoter region.

The metabolic memory effect (or legacy effect) has received much interest since 2003, but the biological mechanisms of this glycemic memory phenomenon remain elusive. The metabolic memory effect is generally defined as a period of good metabolic control that results in a long-term beneficial effect on vascular end points. On the other hand, the metabolic changes caused by the persistence of hyperglycemic stress despite glucose normalization has been defined as the hyperglycemic memory phenomenon (26,27). This observation was initially reported in dogs, rats, and mice with alloxan- and streptozotocin-induced diabetes, where a poor glycemic period followed by a period of good control led to similar rates of retinopathy and expression of fibronectin mRNA in the kidney cortex (28–30). These experiments implied that some irreversible biological changes could be implicated in the progression of microvascular complications after a prolonged period of hyperglycemia. Thus, a better understanding of the pathways leading to hyperglycemic memory may contribute to designing mechanism-driven strategies to blunt vascular damage in patients with diabetes to improve vascular outcomes. The current study provides strong evidence that SHP-1 is persistently activated and expressed in diabetic podocytes, causing sustained inhibition of insulin signaling and podocyte dysfunction and cell death. Furthermore, these data demonstrate that the persistent activation of SHP-1 can be explained by epigenetic changes at the SHP-1 promoter region, thus providing a potential explanation for the hyperglycemic memory effect in podocytes and progression of DN despite glucose level normalization.

Decreased podocyte number and podocyte foot process effacement have been shown to occur in patients with diabetes during the early phases of kidney damage (31), even before the onset of microalbuminuria and more-advanced DN (32). In addition, several clinical observations have suggested that disruption of normal insulin actions is a part of the etiology of DN. A previous study indicated through protein analysis that podocytes have the highest levels of both IR-β and IRS1 expression compared with other glomerular cells, such as renal endothelial and mesangial cells (33). Although the primary defect in type 1 diabetes is diminished insulin secretion, insulin resistance occurrence in type 1 diabetes is widely recognized (34,35), providing support for a renal insulin resistance in type 1 diabetes (11,36). Insulin resistance was shown to precede microalbuminuria in 16 patients with uncomplicated type 1 diabetes (37). Coward et al. (38) showed that podocytes are insulin-responsive cells. By using both a type 1 and a type 2 diabetic rat model, a study demonstrated that the insulin responses in renal endothelial cells were reduced in part by the hyperglycemia-induced polyubiquitination of IRS-1, whereas in the tubular compartment, the insulin signaling pathway was unaffected (33). However, the current study supports the notion that the insulin resistance phenomenon occurs in podocytes and is caused by hyperglycemia-induced persistent expression and activity of SHP-1, resulting in decreased IR-β phosphorylation. Moreover, loss of Akt2, specifically in podocytes, resulted in a rapid disease progression in a mouse model of glomerular diseases (39). Taken together, the capacity of insulin to stimulate Akt is important for podocyte function, and any disruption of its signaling will result in podocyte loss.

Several articles explored the possibility that epigenetic modifications induced by hyperglycemia explain the hyperglycemic memory phenomenon. Studies have demonstrated that HG exposure prompted long-lasting activation of epigenetic changes by promoting Set7 recruitment and H3K4me1 enrichment at the promoter region of the p65 subunit of the nuclear factor-κB in bovine aortic endothelial cells (40,41). In general, acetylation of lysine residues of histone (i.e., H3K9/14) at the gene promoters is associated with transcriptional activation. In contrast, lysine methylation can result in either repression or activation of gene transcription, depending on the amino acid residue modified and the degree of methylation. For example, mono- (me1), di- (me2), and trimethylation (me3) of H3K4 and H3K36 are associated with transcriptionally active genome regions, whereas methylation of H3K9 and H3K27 are associated with gene repression (42). Previous studies have shown that TGF-β–induced collagen IV, connective tissue growth factor, and plasminogen activator inhibitor 1 expression is associated with activation of lysine marks in mesangial cells (43). Studies that used podocytes exposed to HG concentrations showed increased p66^Shc expression in an activated protein C–dependent manner that is accompanied by hyperacetylation of histone at the p66^Shc promoter (44). The current data demonstrate that HG exposure induced an enrichment of H3K4me1 and H3K9/14ac at the SHP-1 promoter region and that these epigenetic modifications were sustained despite glucose level normalization. These results may explain why elevated activity and expression levels of SHP-1 persisted in podocytes and renal glomeruli of diabetic mice. Another study reported that altered activity of Sirt1 and HADC4 have been implicated in the pathogenesis of diabetes-induced renal injury and DN (45,46), suggesting that other epigenetic modifications may contribute to the pathogenesis of DN.

In conclusion, this study provides new insight into the persistent activation and expression of SHP-1 in podocytes, causing sustained insulin resistance and podocyte dysfunction that may account for the observed hyperglycemic memory phenomenon responsible for the continued progression of diabetic renal complications. Because metabolic memory remains a major obstacle in the effective prevention and treatment of diabetic complications, targeting epigenetic changes of SHP-1 could be beneficial to halt the progression of renal injury.

Acknowledgments. The authors thank Anne Vezina (Department of Cell Anatomy, Université de Sherbrooke) for assistance with the histology services.

Funding. F.L. and B.D. are the recipients of a scholarship award from Fonds de Recherche du Québec–Santé. This work was supported by grants from the Canadian Institutes of Health Research (201209PNI, 201505MOP to P.G.), JDRF International (27-2012-531 to P.G), and Diabète Estrie. This work was performed at the Research Center of CHU de Sherbrooke and funded by Fonds de Recherche du Québec–Santé. P.G. currently holds the Canadian Research Chair in Vascular Complications of Diabetes and is the recipient of a Scholarship Award from the Canadian Diabetes Association.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. F.L., B.D., and P.G. contributed to the experiments, data analysis, and writing of the manuscript. A.G. contributed to the animal care and data research. N.G. contributed support and assistance with the ChIP assays. A.M.C. contributed to the data analysis and writing of the manuscript. P.G. is the guarantor of this work and, as such, had full access to all the data and takes responsibility for the integrity of the data and the accuracy of the data analysis.