Diabetes increases susceptibility of primary cultures of rat proximal tubular cells to chemically induced injury

Toxicology and Applied Pharmacology 241 (2009) 1–13

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Diabetes increases susceptibility of primary cultures of rat proximal tubular cells to chemically induced injury Qing Zhong, Stanley R. Terlecky, Lawrence H. Lash ⁎ Department of Pharmacology, Wayne State University School of Medicine, 540 East Canfield Avenue, Detroit, MI 48201, USA

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Article history: Received 27 May 2009 Revised 31 July 2009 Accepted 4 August 2009 Available online 12 August 2009 Keywords: Rat kidney Diabetes Proximal tubular cells Oxidative stress Mitochondrial function Glutathione

a b s t r a c t Diabetic nephropathy is characterized by increased oxidative stress and mitochondrial dysfunction. In the present study, we prepared primary cultures of proximal tubular (PT) cells from diabetic rats 30 days after an ip injection of streptozotocin and compared their susceptibility to oxidants (tert-butyl hydroperoxide, methyl vinyl ketone) and a mitochondrial toxicant (antimycin A) with that of PT cells isolated from agematched control rats, to test the hypothesis that PT cells from diabetic rats exhibit more cellular and mitochondrial injury than those from control rats when exposed to these toxicants. PT cells from diabetic rats exhibited higher basal levels of reactive oxygen species (ROS) and higher mitochondrial membrane potential, demonstrating that the PT cells maintain the diabetic phenotype in primary culture. Incubation with either the oxidants or mitochondrial toxicant resulted in greater necrotic and apoptotic cell death, greater evidence of morphological damage, greater increases in ROS, and greater decreases in mitochondrial membrane potential in PT cells from diabetic rats than in those from control rats. Pretreatment with either the antioxidant N-acetyl-L-cysteine or a catalase mimetic provided equivalent protection of PT cells from both diabetic and control rats. Despite the greater susceptibility to oxidative and mitochondrial injury, both cytoplasmic and mitochondrial glutathione concentrations were markedly higher in PT cells from diabetic rats, suggesting an upregulation of antioxidant processes in diabetic kidney. These results support the hypothesis that primary cultures of PT cells from diabetic rats are a valid model in which to study renal cellular function in the diabetic state. © 2009 Elsevier Inc. All rights reserved.

Introduction Nephropathy is a serious and frequent complication of both type 1 and type 2 diabetes and is a frequent cause of death in diabetic patients (Ibrahim and Hostetter, 1997). On the level of the whole organ, the diabetic kidney is characterized by increased perfusion, which likely leads to increased glomerular filtration and intraglomerular pressure (Remuzzi et al., 2006). Growth of renal tissue and enlargement of glomeruli occur, which lead to pathological changes in glomerular structure, initial microalbuminuria, progression to more extensive proteinuria, loss of tubular filtration, and ultimately renal failure. It is estimated that 35% to 40% of patients with diabetes will develop clinically manifest diabetic nephropathy (Geiss et al., 1993; Sedor, 2006; Susztak and Böttinger, 2006), indicating the importance of prevention and Abbreviations: AA, antimycin A; DCFH, 2,7-dichlorofluorescein; DMEM:F12, Dulbecco's Modified Eagle's Medium:Ham's F12; FACS, flow-activated cell sorting; GSH, glutathione; HE, dihydroethidium; JC-1, 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzoimidazolylcarbocyanine iodide; LDH, lactate dehydrogenase; MVK, methyl vinyl ketone; NAC, N-acetyl-L-cysteine; PT, proximal tubular; ROS, reactive oxygen species; STZ, streptozotocin; tBH, tert-butyl hydroperoxide. ⁎ Corresponding author. Fax: +1 313 577 6739. E-mail address: [email protected] (L.H. Lash). 0041-008X/$ – see front matter © 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.taap.2009.08.007

treatment of diabetic renal disease as a critical public health issue (Remuzzi et al., 2006). Despite the prominence of glomerular changes in diabetic nephropathy, renal function correlates better with tubular and interstitial changes than with glomerular changes (Gilbert and Cooper, 1999). Tubular epithelial cells, in particular those of the proximal tubule, are direct targets for enhanced glucose levels present in diabetes. For example, renal proximal tubular (PT) cells exposed to high concentrations of glucose produce higher levels of reactive oxygen species (ROS), peroxynitrite, and transforming growth factor-β, have enhanced activation of the JAK/STAT signaling pathway and accumulate more calcium ions than PT cells exposed to normal glucose concentrations (Allen et al., 2003; Huang et al., 2007; Yamagishi et al., 2003). These findings and the protective effects of antioxidants indicate that the mechanism of cellular injury in PT cells exposed to chronic hyperglycemia involves various manifestations of oxidative and nitrosative stress (Alderson et al., 2004; Beisswenger et al., 2005; Brownlee, 2001; Chander et al., 2004; Rolo and Palmeira, 2006). Within the renal PT cell, mitochondrial function is significantly altered by chronic hyperglycemia (Rolo and Palmeira, 2006; Rosca et al., 2005). The alterations in cellular redox status and energetics that result from hyperglycemia also result in changes in both total cellular and

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Fig. 1. Morphology of PT cells from control and diabetic rats. PT cells were grown on 35-mm tissue culture dishes for 4 days in complete DMEM:F12 medium. Images were taken on a Zeiss LSM 510 confocal microscope. Magnification was 630×. Scale bar = 5 μm.

mitochondrial glutathione (GSH) status (Ghosh et al., 2005; Kakkar et al., 1997; Obrosova et al., 2003; Ueno et al., 2002; Winiarska et al., 2004; Yue et al., 2005). Consistent with these findings, we found that renal mitochondria, isolated from rats 30 days after treatment with streptozotocin (STZ) to induce diabetes, exhibited higher rates of coupled respiration but a greater extent of inhibition of respiration from exposure to oxidants or a mitochondrial toxicant, and alterations in GSH status as compared to renal mitochondria from age-matched control rats (Zhong and Lash, 2007). Rather than observing GSH depletion in renal mitochondria from diabetic rats, however, we observed elevated concentrations of GSH and rates of transport of GSH into mitochondria, suggesting that some antioxidant processes are upregulated in the diabetic state. Based on the alterations in renal cellular GSH status and mitochondrial function that we and others have observed in the diabetic state, we hypothesize that PT cells from STZ-treated, diabetic rats will be more susceptible to injury from oxidants and mitochondrial toxicants than PT cells from normal or control rats. To test this hypothesis, we prepared suspensions of PT cells from STZ-treated diabetic (isolated 30 days after induction of diabetes) and agematched control rats and grew them in primary cell culture, to assess

their mitochondrial function, cellular and mitochondrial GSH status, and sensitivity to injury from exposure to two model oxidants and a mitochondrial toxicant. When using a primary cell culture model, one question that arises is whether the cells retain the biochemical properties that were inherent at the time of isolation. In this case, the key question is whether the PT cell primary cultures from diabetic rats, as they grow to confluence over a period of 4 to 6 days, retain the changes that exist in vivo as a consequence of the diabetic state. One precedence for these PT cell primary cultures being a valid model is our previous work on PT cell primary cultures from the remnant kidney of rats that have undergone uninephrectomy and compensatory renal growth (Lash et al., 2001, 2006). In those studies, the PT cells retained the cellular hypertrophy and increased expression of numerous enzymes and rates of numerous physiological processes that occur in vivo. By extrapolation, we hypothesize that the PT cell primary cultures isolated from STZ-treated diabetic rats will also retain the biochemical and physiological changes that occur as a consequence of the diabetic state. Results from the present study demonstrate that PT cell primary cultures from diabetic rats exhibit enhanced levels of ROS, elevated mitochondrial membrane potential

Fig. 2. Basal levels of ROS in PT cells from control and diabetic rats. (A) Measurement of basal ROS by confocal microscopy. PT cells were incubated for 15 min with either 5 μM DCFH or 10 μM HE in complete medium. Green fluorescence was measured at 488-nm excitation and 530-nm emission, and red fluorescence was measured at 488-nm excitation and 600nm emission on a Zeiss LSM 510 confocal microscope. Scale bar = 5 μm. (B) Measurement of basal ROS by flow cytometry. PT cells were incubated for 15 min with 5 μM DCFH. Fluorescence-positive cells were counted by flow cytometry with approximately 10,000 events per sample. Results are means ± SE of measurements from 4 separate cell preparations each from control and diabetic rats. ⁎Significantly different (P b 0.05) between control and diabetic PT cells.

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IN). All procedures with rats were conducted in accordance with the Guiding Principles in the Use of Animals in Toxicology, as adopted by the Society of Toxicology in 1989, and all applicable federal and state regulations, and were approved by the Institutional Animal Care and Use Committee. After an acclimation period of 5 days, rats were made diabetic by an ip injection of STZ (60 mg/kg body weight). Rats developed polyuria within 24–48 h. A blood glucose level N250 mg/dl was considered indicative of the diabetic state. Rats with blood glucose N550 mg/dl were given daily injections of insulin (Humulin; 1–2 units IH, Eli Lilly Co., Indianapolis, IN) to maintain them in a ketoacidosis-free, but hyperglycemic state.

Fig. 3. Basal mitochondrial membrane potential in PT cells from control and diabetic rats. (A) Measurement of basal mitochondrial membrane potential by confocal microscopy. PT cells were incubated with 5 μg JC-1/ml for 20 min. JC-1 fluorescence was measured by confocal microscopy assessing the emission of the punctate red (600 nm) in polarized mitochondria using 488-nm excitation on a Zeiss LSM 510 confocal microscope. Magnification = 630×. Scale bar = 5 μm. (B) Measurement of basal mitochondrial membrane potential by flow cytometry. PT cells were stained with JC-1 and populations emitting green fluorescence (monomeric form predominantly in cytoplasm; 488-nm excitation, 530-nm emission) and those emitting red fluorescence (aggregated form predominantly in mitochondria; 488-nm excitation, 600-nm emission) were separated by flow cytometry. Results are the red:green fluorescence ratio and are means ± SE of measurements from 4 separate cell cultures each from control and diabetic rats. ⁎Significantly different (P b 0.05) from control cells.

and GSH concentrations, but greater susceptibility to oxidant- and mitochondrial toxicant-induced cell injury as compared with cells isolated from age-matched control rats. Thus, PT cell primary cultures are an appropriate and useful model in which to study the underlying biochemistry of hyperglycemia-induced nephropathy and to develop potential therapeutic approaches. Materials and methods Chemicals and materials. Percoll, collagenase (type I, EC 3.4.24.3), type I collagen, bovine serum albumin (fraction V), penicillin G, streptomycin, amphotericin B, insulin (from bovine pancreas), human transferrin, sodium selenite, hydrocortisone, epidermal growth factor (EGF; from mouse submaxillary gland), and 3,3′5triiodo-DL-thyronine (T3) were purchased from Sigma Chemical Co. (St. Louis, MO). Dulbecco's Modified Eagle's Medium:Ham's F12 (DMEM:F12, 1:1) was purchased from Mediatech (Herndon, VA). Polystyrene tissue culture dishes were purchased from Corning Inc. (Corning, NY). Polypropylene micromesh (210-μm pore size) was purchased from SpectraPor Inc. (Rancho Dominguez, CA). Animals and establishment of the diabetic state. Male SpragueDawley rats (150–175 g) were purchased from Harlan (Indianapolis,

Isolation and primary culture of renal PT cells. Renal PT cells were prepared from diabetic rats at approximately 30 days after establishment of the diabetic state and from age-matched control rats by collagenase perfusion (Jones et al., 1979) and Percoll density-gradient centrifugation (Lash and Tokarz, 1989). Briefly, kidneys were perfused first with EGTA-containing, Ca2+-free Hanks' buffer at a flow rate of 8 ml/min for 10 min, followed by perfusion with Hanks' buffer containing 0.15% (w/v) collagenase (type I) and 2 mM CaCl2 for 13–18 min at a flow rate of 5 ml/min. All buffers were continuously bubbled with 95% O2/5% CO2 and maintained at 37 °C. At the conclusion of collagenase perfusion, cells were released into KrebsHenseleit buffer, pH 7.4, supplemented with 2.55 mM CaCl2, 10 mM HEPES, and 2% (w/v) bovine serum albumin. Cell count and cell viability were estimated by mixing 0.1 ml of cells with 0.4 ml of 0.2% (w/v) trypan blue in saline and counting the total number of cells and the number of cells that took up the dye on a hemacytometer. The percentage of viable cells was calculated by subtracting the number of cells that took up the dye from the total number of cells, dividing this by the total number of cells and multiplying by 100%. Typically, 85– 95% of the cells excluded the dye. When necessary, cell concentration was adjusted to between 5 × 106 and 8 × 106 cells/ml by dilution with Krebs-Henseleit buffer. To obtain enriched fractions of renal PT cells, cortical cells (5 ml, 5–8 × 106 cells/ml) were layered on 35 ml of an isosmotic 45% (v/v) Percoll solution in 50-ml polycarbonate centrifuge tubes and centrifuged for 30 min at 20,000 × g. The PT cells (upper layer) were estimated to have a purity of 97%, based on marker enzyme activities and cell-type-specific respiratory responses (Lash, 1990; Lash and Tokarz, 1989). Based on enzymology and morphology (Lash, 1990; Lash and Tokarz, 1989), the renal PT cell preparation contains cells derived from both convoluted and straight segments of the proximal tubule. Cell count and cell viability were estimated with trypan blue on a hemacytometer as described above. Cell viability (i.e., fraction of cells that excluded trypan blue) of the PT cells obtained after the Percoll separation was typically 90%–95%. Cell counts were not adjusted for viability before cells were plated for culture. For cell culture, freshly isolated PT cells were suspended in 2 ml of Krebs-Henseleit buffer and diluted with an appropriate amount of culture medium before plating. Basal medium was a 1:1 mixture of DMEM:F12. Supplementation included 15 mM HEPES, pH 7.4, 20 mM NaHCO3, 5 μg insulin/ml (=0.87 μM), 5 μg human transferrin/ml (=66 nM), 100 ng hydrocortisone/ml (=0.28 μM), 100 ng epidermal growth factor/ml (=17 nM), 30 nM sodium selenite, 7.5 pg T3/ml (=111 nM), and an antibiotic mixture containing 192 IU penicillin G/ml, 200 μg streptomycin sulfate/ml, and 2.5 μg amphotericin B/ml (Lash et al., 1995). PT cells were seeded onto collagen-coated, 25-cm flasks or 35-mm polystyrene tissue culture dishes at a density of 2–4 × 106 cells/ml. Cultures were grown at 37 °C in a humidified incubator under an atmosphere of 95% air/5% CO2. Fresh medium was added to the dishes after 24 h (Day 1) and every 48 h thereafter. Cells were processed for the various incubations and assays on Day 4 or Day 5,

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when growth reached ~80% confluence. Although normal, mature PT cells are quiescent, we chose to perform all studies on cells just prior to confluence to prevent over-growth or senescence of cells, which typically occur by Day 6 or Day 7 of culture, but can begin as early as Day 5 (Lash et al., 1995). Use of cells at ~80% confluence should provide better uniformity in responses. Isolation of mitochondria from PT cells. On Day 4 of primary culture, mitochondria and cytoplasm were isolated using the mitochondrial isolation kit from Thermo (Rockford, IL).

Catalase treatment. In addition to N-acetyl-L-cysteine (NAC) (see below), we employed a targeted antioxidant enzyme to quench cellular ROS. The molecule, a genetically engineered version of the enzyme catalase, contains a modified type 1 peroxisome targeting signal at its carboxy terminus (specifically, the three amino acids serine–lysine–leucine or, in single amino acid abbreviations, SKL), and a cell-penetrating polyarginine domain near its amino terminus. Referred to herein as catalase-SKL, the enzyme also contains an amino-terminal hexahistidine tag for purification via nickel-affinity chromatography. Catalase-SKL was expressed,

Fig. 4. NAC and catalase-SKL prevent the increase in ROS formation induced by AA. PT cells received either no pretreatment or were pretreated for 1 h with 5 mM NAC or for 12 h with 1 μM catalase-SKL. PT cells were then treated for 2 h with 2 μM AA. After treatments, cells were incubated for 15 min with 5 μM DCFH and fluorescence was measured by flow cytometry. (A) Fluorescence peak shifts. Diagram of representative fluorescence peak shifts elicited by toxicants with and without pretreatments. The proportion of cells emitting high fluorescence (= high level of ROS) are denoted by the percentages in each panel. (B) Statistical analysis of ROS peak shift data. Quantitation of the proportion of cells emitting high fluorescence. Results are means ± SE of measurements from 3 separate cell cultures each from control and diabetic rats. ⁎Significantly different (P b 0.05) from basal level within each group. #Significantly different (P b 0.05) from values for the corresponding treatment group.

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purified, characterized, and employed (at 1 μM) as described by Young et al. (2008). Phase contrast microscopy. On Day 4, cells were incubated for 4 h with either medium or various concentrations of antimycin A (AA), methyl vinyl ketone (MVK), or tert-butyl hydroperoxide (tBH), with

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or without a 1-h preincubation with 5 mM NAC, and morphology was assessed with a Nikon TM S microscope and attached Olympus DP 12 digital camera. Confocal microscopy. PT cells were grown on collagen-coated, 35mm culture dishes and were viewed with a Zeiss Triple-Laser Scanning

Fig. 5. NAC prevents the increase in ROS formation induced by AA, MVK, and tBH. PT cells received either no pretreatment or were pretreated for 1 h with 5 mM NAC. PT cells were then treated for 2 h with either 10 μM AA, 25 μM MVK, or 100 μM tBH. After treatments, cells were incubated for 15 min with 5 μM DCFH and fluorescence-positive cells were measured by flow cytometry. (A) Fluorescence peak shifts induced by toxicants and NAC. Shown is a representative set of incubations with overlays of fluorescence cell distributions with no addition (green tracing), addition of the indicated toxicant (blue tracing), and cells preincubated with NAC and incubated with the indicated toxicant (pink tracing). (B) Statistical analysis of ROS peak shift data. Besides the incubations with AA, MVK, and tBH, some cells were also incubated for 2 h in a simulated, hyperglycemic environment (additional 300 mg% glucose in medium; total glucose content = 615 mg%). Results are the relative fluorescence intensity and are means ± SE of measurements from 4 separate cell cultures each from control and diabetic rats. ⁎Significantly different (P b 0.05) from the basal level within the same group. #Significantly different (P b 0.05) from values of the corresponding treatment group.

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Confocal Microscope (LSM 510) with integrated workstation at the Confocal Imaging Core Facility in the School of Medicine at Wayne State University (Detroit, MI). This is a core facility of the NIEHS Center

for Molecular Toxicology with Human Applications at Wayne State. Initial magnification was 196×. Cells were stained with 5 μg/ml of 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-benzoimidazolylcarbocya-

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nine iodide (JC-1) for 20 min for measurement of mitochondrial membrane potential or with either 5 μM 2,7-dichlorofluorescein (DCFH) or 10 μM dihydroethidium (HE) for 15 min for measurement of ROS levels. These incubation times for the fluorescent dyes were sufficient to achieve complete equilibration in PT cells from both control and diabetic rats. Determination of ROS levels by flow cytometry. PT cells were incubated with either 10 μM AA, 100 μM tBH, or 25 μM MVK for 2 h with or without a 1-h preincubation with 5 mM NAC. Cells were then incubated with 5 μM DCFH for 15 min, media were withdrawn, cells were washed with PBS, removed by brief trypsinization, cell pellets were resuspended in 0.5 ml of complete DMEM:F12 medium, and samples were covered with foil and kept on ice until analysis. Cells that were green fluorescence-positive were counted by flow cytometry using a FACSCalibur Flow Cytometer (BD Biosciences, San Jose, CA), with analysis of approximately 20,000 events per sample. Determination of mitochondrial membrane potential by flow cytometry. PT cells were incubated as in the ROS measurements and then incubated for 20 min with 5 μg of JC-1/ml. After washing with PBS and trypsinization, cell pellets were resuspended in 0.5 ml of complete medium, covered by foil, and kept on ice until analysis. Green (=monomeric form of dye found in cytoplasm) and red (=aggregated form of dye accumulated in mitochondria) fluorescent-positive cells were counted by flow cytometry using a FACSCalibur Flow Cytometer (BD Biosciences, San Jose, CA) with approximately 20,000 events per sample. Assessment of cell death by LDH release assay. Cells grown on 35-mm dishes were treated with either AA (2, 8, or 16 μM), tBH (50, 100, or 200 μM), or MVK (25, 50, or 100 μM) for 4 h with or without a 1h preincubation with 5 mM NAC. LDH activity in supernatants and total cell extracts was measured spectrophotometrically as NADH oxidation at 340 nm after addition of pyruvate and NADH. The fraction of LDH release was calculated by the formula: %LDH release = [LDH activity in medium/(LDH activity in medium + LDH activity in total cells)] × 100%. Assessment of apoptosis by FACS and flow cytometry. Cells were dislodged from culture dishes by brief tryspin/EDTA treatment, fixed in 70% (v/v) ethanol, stained with propidium iodide (50 μg/ml) containing RNAse A (100 U/ml), and were analyzed by flow cytometry using a FACSCalibur Flow Cytometer (BD Biosciences, San Jose, CA). Analysis was performed with 20,000 events per sample using ModFit LT v. 3 for Macintosh data acquisition software package. Propidium iodide was detected by the FL-2 photomultiplier tube. Fractions of apoptotic cells were quantified by analysis of the sub-G1 (subdiploid) peak. Cell aggregates were discarded in the flow cytometry analysis by postfixation aggregate discrimination; cells outside the box in each inset (distribution of cells according to fluorescence intensity) were those excluded from analysis. Percentage values in each panel indicate the percentage of subdiploid cells. Measurement of GSH contents. GSH contents in mitochondria and cytoplasm from PT cells were measured with the GSH-Glo kit from

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Promega (Madison, WI) and were quantified by chemiluminescence in a SpectraMax 2 plate reader using GSH as a standard. Data analysis. All values are expressed as means ± SE of measurements from the indicated number of individual cell cultures. Fisher's protected least significance difference t-test was performed to determine significant differences between pairs with P values b 0.05 considered significant. Results Morphologic similarity between control and diabetic PT cells PT cells from 1-month control and diabetic rats, obtained by collagenase perfusion and Percoll density-gradient centrifugation, were assessed by phase-contrast microscopy after 4 days in culture (Fig. 1). Cells from both control and diabetic rats appeared polygonal with no obvious cellular hypertrophy in the PT cells from the diabetic rats. Thus, epithelial morphology and growth patterns of primary cell cultures were similar in cells from both the control and diabetic rats. PT cells from diabetic rats have higher basal levels of ROS than those from control rats PT cells from control and diabetic rats on Day 4 of culture were incubated with 5 μM DCFH or 10 μM HE for 15 min, and fluorescence was analyzed by confocal microscopy to obtain a qualitative assessment of basal levels of ROS (Fig. 2A). Intensity of green fluorescence due to DCFH and red fluorescence due to HE were both notably higher in PT cells from diabetic rats. DCFH-stained PT cells were then subjected to flow cytometry analysis to obtain a more quantitative measure of basal ROS levels (Fig. 2B). Basal levels of ROS, as indicated by the fraction of cells possessing a high level of green fluorescence, were approximately 2-fold higher in PT cells from diabetic rats. PT cells from diabetic rats have higher basal mitochondrial membrane potential than those from control rats PT cells from control and diabetic rats on Day 4 of culture were incubated with 5 μM JC-1 for 20 min, and fluorescence was analyzed by confocal microscopy to obtain a qualitative assessment of basal mitochondrial membrane potential (Fig. 3A). JC-1 is a mitochondrial membrane potential-sensitive dye that forms red aggregated punctate dots in cells with hyperpolarized mitochondria, whereas the dye is viewed as diffuse, green fluorescence in cells containing depolarized mitochondria. Confocal microscopy showed that PT cells from diabetic rats contained a markedly higher degree of hyperpolarized mitochondria. The ratio of red to green fluorescence intensity from JC-1 was then quantified by flow cytometry to obtain a more quantitative assessment of basal mitochondrial membrane potential (Fig. 3B). As with ROS measurement by flow cytometry, JC-1 red-to-green fluorescence ratios in PT cells from diabetic rats were nearly 2-fold higher than those from control rats, consistent with the confocal microscopy photomicrographs.

Fig. 6. NAC diminishes the decreases in mitochondrial membrane potential induced by AA, MVK, and tBH. PT cells received either no pretreatment or were pretreated for 1 h with 5 mM NAC. PT cells were then treated for 2 h with either 10 μM AA, 25 μM MVK, or 100 μM or 200 μM tBH. After treatments, cells were incubated for 20 min with 5 μg JC-1/ml and fluorescence distribution of cells was measured by flow cytometry. (A) Green fluorescence peak shifts induced by toxicants and NAC. Shown is a representative set of incubations with overlays of fluorescence cell distributions with no addition (green tracing), addition of the indicated toxicant (blue tracing), and cells preincubated with NAC and incubated with the indicated toxicant (pink tracing). (B) Statistical analysis of JC-1 green peak shift data. Besides the incubations with AA, MVK, and tBH, some cells were also incubated for 2 h in a simulated, hyperglycemic environment (additional 300 mg% glucose in medium; total glucose content = 615 mg%). Results are the relative, green fluorescence intensity, indicating cells with low mitochondrial membrane potential, and are means ± SE of measurements from 4 separate cell cultures each from control and diabetic rats. ⁎Significantly different (P b 0.05) from the basal level within the same group. #Significantly different (P b 0.05) from values of the corresponding treatment group. (C) JC-1 red:green ratio. Ratios of red:green fluorescence for the incubation groups shown in panel B are calculated here. ⁎Significantly different (P b 0.05) from the basal level within the same group. #Significantly different (P b 0.05) from values of the corresponding treatment group.

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NAC and catalase-SKL both prevent AA-induced ROS increase Besides the basal differences in ROS levels and mitochondrial membrane potential, an important question is how PT cells from control and diabetic rats respond to chemical challenges. We first examined the effects of AA, a mitochondrial complex III inhibitor that is known to increase formation of ROS, on ROS formation. A relatively low concentration of AA (2 μM) was used to elicit a modest amount of ROS formation. PT cells from control and diabetic rats were incubated for 2 h with 2 μM AA, with or without either a 1-h preincubation with 5 mM NAC or a 12-h preincubation with 1 μM catalase-SKL, and DCFH fluorescence was quantified by flow cytometry. Incubation with AA caused a peak shift to the right, indicating that the proportion of cells with higher DCFH fluorescence intensity increased (Fig. 4A). The proportion of high-fluorescence intensity cells after AA treatment was greater in PT cells from diabetic rats than in those from control rats (22.6% vs. 53.8%, respectively). Preincubation with either NAC or catalase-SKL almost completely prevented the peak shift in PT cells from both control and diabetic rats. Analysis of the proportion of high-fluorescence intensity cells further illustrated the greater response to AA in PT cells from diabetic rats and the equivalent ability in cells from both control and diabetic rats of NAC and catalase-SKL to prevent the increase (Fig. 4B). NAC prevents increases in ROS induced by AA, tBH, or MVK The oxidative response of PT cells from control and diabetic rats was further studied by assessing effects of two model oxidants, tBH and MVK, comparing this to the effect of AA, and assessing the ability of NAC to prevent the increases in ROS levels. A higher concentration of AA (10 μM) than that used above was used in these experiments to elicit production of ROS in amounts comparable to those produced by tBH and MVK. PT cells from control or diabetic rats were incubated for 2 h with 10 μM AA, 100 μM tBH, or 25 μM MVK, with or without a 1h pretreatment with 5 mM NAC, and ROS were quantified by incubation with 5 μM DCFH and flow cytometry (Fig. 5). All three toxicants shifted the fluorescence peak to the right (blue tracing vs. green tracing), demonstrating an increase in ROS formation (Fig. 5A). The extent of the shift was generally greater in PT cells from diabetic rats, indicating more toxicant-induced ROS formation than occurs in PT cells from control rats. Preincubation with NAC (pink tracing) resulted in comparable decreases in ROS formation in toxicant-treated PT cells from both control and diabetic rats. These responses were quantified according to the proportion of cells exhibiting high fluorescence (Fig. 5B) and showed that both basal and toxicantstimulated ROS levels are higher in PT cells from diabetic rats but that preincubation with NAC provides equivalent and nearly complete protection in PT cells from both control and diabetic rats. In addition, cells were incubated in a simulated hyperglycemic environment (addition of 300 mg% glucose to normal medium, which already contains 315 mg% glucose). This treatment, however, had no effect on ROS levels in PT cells from either group. NAC prevents decreases of mitochondrial membrane potential induced by AA, tBH, or MVK Similar to the experiments described above for ROS levels, effects of AA, tBH, and MVK on mitochondrial membrane potential were determined in PT cells by flow cytometry (Fig. 6). Treatment of PT cells with each of the chemicals caused a shift in the green fluorescence peak (blue tracing for toxicant treated vs. green tracing

Fig. 8. NAC protects PT cells from increased LDH release induced by AA, tBH, and MVK. PT cells were incubated for 4 h with the indicated concentrations of medium (= Control), AA, tBH, and MVK, with or without a 1-h preincubation with 5 mM NAC. LDH activities in supernatants and cells were measured spectrophotometrically as NADH oxidation at 340 nm. Results are means ± SE of measurements from 4 separate cell cultures each for cells from control and diabetic rats. ⁎Significantly different (P b 0.05) from the corresponding control incubation.

for untreated controls) to the right, indicating increases in the proportion of cells possessing lower mitochondrial membrane potentials and, therefore, less aggregation and mitochondrial accumulation of JC-1 (Fig. 6A). The rightward shifts were greater in the PT cells from diabetic rats, again indicating a larger mitochondrial toxicity response in cells from the diabetic rats. Preincubation with 5 mM NAC almost completely (and equivalently in PT cells from both control and diabetic rats) prevented the shifts in fluorescence peaks (pink tracing). Quantitation of the population of JC-1-treated cells with a high amount of green fluorescence (i.e., low membrane potential) similarly showed that PT cells from diabetic rats generally experienced greater increases in the low-membrane potential subpopulation of cells after toxicant exposure than do PT cells from control rats (Fig. 6B). There was a striking concentration dependence with tBH as the toxicant, with 100 μM tBH producing a 1.3fold and 2.4-fold increase in control and diabetic PT cells, respectively, and 200 μM tBH producing a 5.2-fold and 52.3-fold increase in control and diabetic PT cells, respectively. Unlike the assessment of ROS (cf. Fig. 5B), incubation of cells with an additional 300 mg% glucose (hyperglycemia) did cause a significant increase in the population of low-membrane potential cells, but only in PT cells from diabetic rats. Preincubation with NAC prevented the shift to low membrane potential but was only partially effective in PT cells from diabetic rats with 200 μM tBH as the toxicant. We also show the red-to-green fluorescence ratio for the various treatment groups of PT cells from control and diabetic rats as a direct indicator of the proportion of cells with high mitochondrial membrane potential (Fig. 6C). The pattern observed here is virtually the inverse of that for the green fluorescence intensity shown in Fig. 6B. Thus, tBH caused concentration-dependent decreases in the red-to-green ratio, and these were more pronounced in PT cells from diabetic rats. MVK was similarly very

Fig. 7. NAC protects PT cells from morphological damage due to AA (A), tBH (B), and MVK (C). PT cells were incubated for 4 h with medium (= Control) or the indicated concentrations of AA, tBH, and MVK, with or without a 1-h preincubation with 5 mM NAC. Cellular morphology was assessed by phase contrast microscopy, using a Nikon TM S microscope. Magnification = 150×.

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effective in decreasing the red-to-green ratio in PT cells from both control and diabetic rats. Acute hyperglycemia produced no significant effect on the red-to-green ratios. NAC was protective, producing complete protection from AA- and MVK-induced depolarization, but only partial protection from tBH-induced depolarization, particularly at the higher concentration of tBH in PT cells from diabetic rats. Morphological damage and acute cytotoxicity of AA, tBH, and MVK On Day 4 of primary culture, when PT cells were approximately 80% confluent, cells were incubated with a high and low concentration of each toxicant for 4 h, with some cells incubated with the higher concentration of toxicant also preincubated for 1 h with 5 mM NAC. Cellular morphology was then assessed by phase contrast microscopy and sample photomicrographs are shown only for the controls and cells treated with the high concentration of toxicant ± NAC (Fig. 7). AA produced modest cell damage in PT cells from both control and diabetic rats, and preincubation with 5 mM NAC largely prevented this damage. tBH produced noticeably

greater morphological damage in PT cells from diabetic rats, and preincubation with 5 mM NAC largely prevented this damage, although protection appeared less complete in PT cells from diabetic rats. In contrast, the extent of morphological damage from MVK appeared to be similar in PT cells from both control and diabetic rats, but again this damage was largely prevented by preincubation with NAC. Acute cytotoxicity was assessed in a more quantitative manner by measuring LDH release after incubating cells for 4 h with either MVK, tBH, or AA, with or without a 1-h preincubation with 5 mM NAC (Fig. 8). Although the concentrations of toxicants used were not always the same as those used in previous experiments, the intention was to expose cells to toxicants at 2 to 3 concentrations that would produce a range of LDH release. PT cells incubated with AA only exhibited significant LDH release with 16 μM AA; no difference was observed in the response of cells from control and diabetic rats and preincubation with NAC completely prevented the increase in LDH release. For cells incubated with MVK, a concentration-dependent increase in LDH release was observed, but no differences were observed between control and diabetic PT cells. Similar to results

Fig. 9. NAC prevents apoptosis in PT cells induced by MVK. PT cells were incubated for 4 h with 100 μM MVK with or without a 1-h preincubation with 5 mM NAC. Cells were processed for FACS analysis by flow cytometry, as described in Materials and methods, using 20,000 events/sample and ModFit LT v. 3 Macintosh data acquisition software package. Fractions of apoptotic cells were quantified as the sub-G1 peaks and are indicated by the percentages in each panel. Cells outside the boxes in each inset were those excluded from analyses.

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with AA, preincubation with NAC completely prevented the increases in LDH release due to MVK. In contrast to results with AA and MVK, incubation of PT cells from diabetic rats with tBH exhibited significantly more LDH release than PT cells from control rats at the highest concentration tested. Preincubation with NAC completely prevented the increases in LDH release with 100 μM tBH, only partially prevented the increase in LDH release with 200 μM tBH in PT cells from diabetic rats, and had no significant effect on the LDH release induced by 200 μM tBH in PT cells from control rats. Besides LDH release, which is considered a measure of cellular necrosis, we also compared the induction of apoptosis by all three toxicants, as assessed by propidium iodide staining, FACS analysis, and flow cytometry. Neither AA nor tBH produced significant increases in the fraction of subdiploid cells from either control or diabetic rats (data not shown). In contrast, incubation for 4 h with 100 μM MVK produced a small increase in the fraction of apoptotic cells in PT cells from control rats (4.66% subdiploid cells) but a much larger increase in the fraction of apoptotic cells in PT cells from diabetic rats (44.0% subdiploid cells) (Fig. 9). Preincubation for 1 h with 5 mM NAC completely prevented MVK-induced apoptosis in PT cells from both control and diabetic rats. GSH content is higher in cytoplasm and mitochondria of PT cells from diabetic rats Nearly confluent PT cells from control and diabetic rats were fractionated into cytoplasmic and mitochondrial compartments to analyze the contents of GSH (Fig. 10). GSH concentrations were 2.6-fold and 6.7-fold higher in cytoplasm and mitochondria from PT cells of diabetic rats, respectively, as compared to corresponding fractions from PT cells of control rats. Discussion Diabetes is considered to be a metabolic disorder in which alterations in mitochondrial function play a central role in the mechanism of disease progression in various affected organs (Brownlee, 2001; Rolo and Palmeira, 2006; Wallace, 2005). It is also a general risk factor for many forms of drug-induced nephrotoxicity in humans (Naughton, 2008), making an improved understanding of the biochemical basis of diabetic nephropathy important for improving human health. Some specific forms of drug-induced nephrotoxicity that have been shown in epidemiological studies to exhibit increases in populations of diabetic patients or otherwise exposed individuals include radiocontrast agents (Andrew and Berg, 2004; Badero et al., 2008), envi-

Fig. 10. Contents of GSH in cytoplasm and mitochondrial from PT cells. After 4 days in primary culture, mitochondrial and cytoplasmic fractions were isolated from PT cells from control and diabetic rats. GSH contents were measured using the Promega GSHGlo kit and assayed by luminescence in a plate reader. Results are means ± SE of measurements from 5 separate cell cultures each from control and diabetic rats. ⁎Significantly different (P b 0.05) from corresponding fraction from control rats.

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ronmental contaminants of drinking water such as uranium (Seldén et al., 2009), aminoglycosides when used in the elderly (Baciewicz et al., 2003), and lead (Ekong et al., 2006). A recent study in a rat model of diabetes (Doi and Ishida, 2009) demonstrated a marked increase in acetaminophen-induced nephrotoxicity as compared to that in normal rats, suggesting that the increased susceptibility of diabetics in the human population applies to a very broad range of potentially nephrotoxic drugs and chemicals. Previous work of ours (Zhong and Lash, 2007) showed that isolated mitochondria from rat renal cortex of diabetic rats exhibited higher respiration rates, significantly higher GSH levels and modestly higher rates of GSH uptake into mitochondria, but higher basal and toxicant-stimulated rates of lipid peroxidation. These data suggested that rat kidneys, in particular the proximal tubules, exhibit upregulation of mitochondrial function and some antioxidants in response to the chronic hyperglycemic state but that these apparently adaptive changes are insufficient to prevent higher levels of oxidative stress. Accordingly, an oxidative challenge with tBH or MVK produced a greater extent of inhibition of state 3 respiration in isolated mitochondria from diabetic rat kidneys than in those from normal rat kidneys. This last observation implies that kidneys from diabetic rats are more susceptible to injury from exposure to toxic chemicals and that diabetes, and the ensuing diabetic nephropathy, are additional risk factors for exposure to chemical toxicants. In the present study, we sought to extend those observations by developing primary cultures of PT cells from diabetic rats as an intact cellular model in which to investigate the processes responsible for the altered biochemistry of PT cells in diabetic rats. Several lines of evidence showed that even after 30 days of diabetes, a time point at which diabetic nephropathy in the STZ-treated rat has not yet fully manifested (Obrosova et al., 2003; Ueno et al., 2002; Yue et al., 2005), significant changes were observed in mitochondrial function and redox homeostasis. The central role of oxidative stress, mitochondrial dysfunction, and perturbations in GSH homeostasis have been well established in kidneys from the STZ diabetic rat (Alderson et al., 2004; Hamada et al., 2007; Katyare and Satav, 2005; Obrosova et al., 2003; Ueno et al., 2002; Yue et al., 2005). Moreover, evidence of increased lipid peroxidation in diabetic nephropathy has been obtained in both patients and STZ-treated rats (Chang et al., 2005). Thus, there is ample precedence to investigate the role of alterations in mitochondrial function and redox homeostasis as underlying mechanisms in the pathology of diabetic nephropathy. Analysis of the morphology of PT cells from control and diabetic rats showed both cell cultures exhibited normal epithelial shape and growth patterns and revealed no significant differences in these parameters. This suggests that primary culture of PT cells from diabetic rats retain normal cellular morphology and can be used as an in vitro model for study of diabetic nephropathy. Despite the similar morphology, analysis of basal levels of ROS and mitochondrial membrane potential revealed marked differences between PT cells from normal and diabetic rats. For investigation of ROS, we used two fluorogenic probes, DCFH and HE, and found that PT cells from diabetic rats exhibited much higher intensity of fluorescence by confocal microscopy. As DCFH detects primarily H2O2 and peroxynitrite whereas HE detects primarily superoxide anions (Walrand et al., 2003), this indicates that multiple species of ROS and potentially reactive nitrogen species are involved in the biochemical changes underlying the diabetic state. PT cells from diabetic rats similarly exhibited more intense red fluorescence from staining with the mitochondrial membrane potential-sensitive dye JC-1 as compared to PT cells from control rats. Such an elevation in basal mitochondrial membrane potential is consistent with our previous finding of higher rates of state 3 respiration in isolated renal mitochondria from diabetic rats (Zhong and Lash, 2007). The mechanism by which this hypermetabolic state produces hyperpolarization of PT cell mitochondria, however, requires

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further investigation. The retention of elevated ROS and mitochondrial membrane potential in primary PT cell cultures provides further evidence that the diabetic phenotype is retained in cell culture and that this model can be used to investigate underlying mechanisms for the observed biochemical and functional changes. These alterations in ROS levels and mitochondrial membrane potential were further studied by assessing the influence of two model oxidants, tBH and MVK, and a mitochondrial toxicant, AA, on the toxic response of PT cell cultures. While all three test agents can generate ROS and an oxidative stress, the subtle differences in responses of the cells to exposure are due to differences in mechanism of action. While tBH is itself an organic peroxide that oxidizes GSH by serving as a substrate for GSH peroxidase and generates H2O2, MVK alkylates soft nucleophiles such as GSH, thereby causing GSH depletion and oxidative stress. In contrast, AA generates ROS by inhibiting the mitochondrial electron transport chain, thereby promoting partial reduction of molecular oxygen. The effectiveness of catalase-SKL in protecting PT cells from AA-induced increases in ROS as detected by DCFH suggests that H2O2 is the primary form of ROS that is involved in AA-induced toxicity. NAC, which can serve as a precursor to GSH but also act independently as an antioxidant, was similarly protective against toxicant-induced increases in ROS and decreases in mitochondrial membrane potential. The absence of a significant cytotoxic effect of a simulated, acute hyperglycemic environment (i.e., incubation with an additional 300 mg% glucose) in PT cells from control rats suggests that the effects of hyperglycemia require chronic exposure. This makes sense in light of the proposed biochemical mechanisms for cytotoxicity from hyperglycemia involving metabolic disturbances and mitochondrial dysfunction. It was interesting that the acute, hyperglycemic incubation did modestly enhance the decrease in mitochondrial membrane potential but only in PT cells from diabetic rats. This is consistent with the greater sensitivity of PT cells from diabetic rats to chemical stresses. The protective ability of catalase-SKL and NAC was roughly equivalent in PT cells from control and diabetic rats. Several examples exist in the literature of antioxidant therapy ameliorating the adverse effects of experimental diabetic nephropathy. Examples include dietary GSH (Ueno et al., 2002), lipoic acid (Obrosova et al., 2003; Prabhakar et al., 2007), and vitamins E and C (Kedziora-Kornatowska et al., 2003; Kim et al., 2000; Lee et al., 2007; Stahl and Sies, 1997). Although studies specifically focusing on the mitochondrial pool of GSH have been conducted in cardiac cells (Ghosh et al., 2005), no such studies have been conducted in the kidneys of diabetic rats. Our previous work in suspensions of isolated renal mitochondria (Zhong and Lash, 2007) and the current results showing that mitochondrial GSH contents in PT cells from diabetic rats were increased to a much greater extent than cytoplasmic GSH contents point to the importance of this specific pool of GSH in diabetic nephropathy. The mitochondrial GSH pool is determined by transport of cytoplasmic GSH into the mitochondria by the action of two organic anion transporters, the dicarboxylate carrier (Slc25a10) and 2-oxoglutarate carrier (Slc25a11) (Lash, 2006). Altered expression of one or both of these carriers may occur in diabetes (Kaplan et al., 1990; Zhong and Lash, 2007), further suggesting that these proteins could be therapeutic targets for novel approaches to ameliorating the altered mitochondrial function and redox state of PT cells in diabetes. Although it was beyond the scope of the current study to investigate the relationship between the different cellular responses, it is clear that increases in ROS formation and decreases in mitochondrial membrane potential are closely related and are quite sensitive to mitochondrial inhibitors or oxidants. Some differential ability of NAC to protect was noted with cells incubated with tBH, when assessing morphology and LDH release. For example, whereas pretreatment of PT cells from both control and diabetic rats only partially reversed the increases in LDH release induced by 200 μM tBH (cf. Fig. 8),

protection appears to be almost complete when looking at cellular morphology (cf. Fig. 7B). Two conclusions to draw from this are that LDH release is a more sensitive gauge of cell injury than microscopic examination of morphology and that some amount of LDH release can occur without significant evidence of morphological damage. A final issue is the apparent paradox that although cellular and mitochondrial concentrations of GSH are markedly higher in PT cells from diabetic rats, these cells are more sensitive to oxidative injury. This suggests two possible interpretations. First, the increase in GSH may be a compensatory response to the chronic, hyperglycemic state but is insufficient to protect the cells when they are challenged by exposure to an oxidant or oxidant-generating chemical. Second, cellular or mitochondrial GSH may not serve as an appropriate marker of anti-oxidant capacity in renal PT cells from diabetic rats; rather, other parameters, such as the thioredoxin system, for example, may be more accurate indicators of redox status. Additional studies are ongoing to assess these possibilities. In conclusion, the present work demonstrates the validity of using primary cultures of PT cells from diabetic rats to study the underlying biochemistry of diabetic nephropathy. This is the first study to show that basal differences exist in ROS and mitochondrial membrane potential in PT cells from control and diabetic rats and that the latter are more susceptible to injury from exposure to oxidants or a mitochondrial toxicant. Acknowledgments This research was supported by Department of Defense Grant PR64340 and the NIEHS Center for Molecular Toxicology with Human Applications (Grant P30-ES06639) at Wayne State University. References Alderson, N.L., Chachich, M.E., Frizzell, N., Canning, P., Metz, T.O., Januszewski, A.S., Youssef, N.N., Slitt, A.W., Baynes, J.W., Thorpe, S.R., 2004. Effect of antioxidants and ACE inhibition on chemical modification of proteins and progression of nephropathy in the streptozotocin diabetic rat. Diabetologia 47, 1385–1395. Allen, D.A., Harwood, S.M., Varagunam, M., Raftery, M.J., Yaqoob, M.M., 2003. High glucose-induced oxidative stress causes apoptosis in proximal tubular epithelial cells and is mediated by multiple caspases. FASEB J. 17, 908–910. Andrew, E., Berg, K.J., 2004. Nephrotoxic effects of X-ray contrast media. J. Toxicol. Clin. Toxicol. 42, 325–332. Baciewicz, A.M., Sokos, D.R., Cowan, R.I., 2003. Aminoglycoside-associated nephrotoxicity in the elderly. Ann. Pharmacother. 37, 182–618. Badero, O.J., Schlanger, L., Rizk, D., 2008. Gadolinium nephrotoxicity: case report of a rare entity and review of the literature. Clin. Nephrol. 70, 518–522. Beisswenger, P.J., Drummond, K.S., Nelson, R.G., Howell, S.K., Szwergold, B.S., Mauer, M., 2005. Susceptibility to diabetic nephropathy is related to dicarbonyl and oxidative stress. Diabetes 54, 3274–3281. Brownlee, M., 2001. Biochemistry and molecular cell biology of diabetic complications. Nature 414, 813–820. Chander, P.N., Gealekman, O., Brodsky, S.V., Elitok, S., Tojo, A., Crabtree, M., Gross, S.S., Goligorsky, M.S., 2004. Nephropathy in Zucker diabetic fat rat is associated with oxidative and nitrosative stress: prevention by chronic therapy with a peroxynitrite scavenger ebselen. J. Am. Soc. Nephrol. 15, 2391–2403. Chang, J.M., Kuo, M.C., Kuo, H.T., Chiu, Y.W., Chen, H.C., 2005. Increased glomerular and extracellular malondialdehyde levels in patients and rats with diabetic nephropathy. J. Lab. Clin. Med. 146, 210–215. Doi, K., Ishida, K., 2009. Diabetes and hypertriglyceridemia modify the mode of acetaminophen-induced hepatotoxicity and nephrotoxicity in rats and mice. J. Toxicol. Clin. Toxicol. 34, 1–11. Ekong, E.B., Jaar, B.G., Weaver, V.M., 2006. Lead-related nephrotoxicity: a review of the epidemiologic evidence. Kidney Int. 70, 2074–20874. Geiss, L.S., Herman, W.H., Goldschmid, M.G., DeStefano, F., Eberhardt, M.S., Ford, E.S., German, R.R., Newman, J.M., Olson, D.R., Sepe, S.J., 1993. Surveillance for diabetes mellitus: United States, 1980–1989. MMWR CDC Surveill. Summ. 42, 1–20. Ghosh, S., Pulinilkunnil, T., Yuen, G., Kewalramani, G., An, D., Qi, D., Abrahani, A., Rodrigues, B., 2005. Cardiomyocyte apoptosis induced by short-term diabetes requires mitochondrial GSH depletion. Am. J. Physiol. 289, H768–H776. Gilbert, R.E., Cooper, M.E., 1999. The tubulointerstitium in progressive diabetic kidney disease: more than an aftermath of glomerular injury? Kidney Int. 56, 1627–1637. Hamada, Y, Miyata, S., Nii-Kono, T., Kitazawa, R., Kitazawa, S., Higo, S., Fukunaga, M., Ueyama, S., Nakamura, H., Yodoi, J., Fukugawa, M., Kasuga, M., 2007. Overexpression of thioredoxin1 in transgenic mice suppresses development of diabetic nephropathy. Nephrol. Dial. Transplant. 22, 1547–1557.

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