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Published in final edited form as: Hypertension. 2009 Dec 28;55(2):468–473. doi: 10.1161/HYPERTENSIONAHA.109.145714

Protein Kinase C-Dependent NAD(P)H Oxidase Activation Induced by Type 1 Diabetes in Renal Medullary Thick Ascending Limb

Jing Yang 1, Pascale H Lane 1, Jennifer S Pollock 1, Pamela K Carmines 1
PMCID: PMC4586163  NIHMSID: NIHMS724025  PMID: 20038746

Abstract

Type 1 diabetes provokes a protein kinase C (PKC)-dependent accumulation of superoxide anion in the renal medullary thick ascending limb (mTAL). We hypothesized that this phenomenon involves PKC-dependent NAD(P)H oxidase activation. The validity of this hypothesis was explored using mTAL suspensions prepared from rats with streptozotocin-induced diabetes and from sham (vehicle-treated) rats. Superoxide production was 5-fold higher in mTAL suspensions from diabetic rats compared with suspensions from sham rats. The NAD(P)H oxidase inhibitor apocynin caused an 80% decrease in superoxide production by mTALs from diabetic rats (P<0.05 vs untreated) without altering superoxide production by sham mTALs. NAD(P)H oxidase activity was more than two-fold higher in mTALs from diabetic rats than in sham mTALs (P<0.05). Pretreatment with calphostin C (broad-spectrum PKC inhibitor) or rottlerin (PKCδ inhibitor) reduced NAD(P)H oxidase activity by ~80% in both groups; however, PKCα/β or PKCβ inhibition did not alter NAD(P)H oxidase activity in either group. Protein levels of Nox2, Nox4 and p47phox were significantly higher in diabetic mTALs than in mTALs from sham rats. In summary, elevated superoxide production by mTALs from diabetic rats was normalized by NAD(P)H oxidase inhibition. PKC-dependent, PKCδ-dependent, and total NAD(P)H oxidase activity was greater in mTALs from diabetic rats compared with sham. Protein levels of Nox2, Nox4 and p47phox were increased in mTALs from diabetic rats. We conclude that increased superoxide production by the mTAL during diabetes involves a PKCδ-dependent increase in NAD(P)H oxidase activity, in concert with increased protein levels of catalytic and regulatory subunits of the enzyme.

Keywords: Protein kinase C, NAD(P)H oxidase, thick ascending limb, type 1 diabetes

Oxidative stress is involved in several diseases including hypertension, atherosclerosis, heart failure and diabetes. During type 1 diabetes (T1D), oxidative stress is evident in endothelial cells, fibroblasts, vascular smooth muscle cells, renal mesangial cells and renal tubular epithelial cells.1 Each of these cell types can be found in the kidney and, indeed, oxidative stress is a pathogenic cofactor in the development of the renal complications of T1D, contributing to the glomerulosclerosis and tubulointerstitial fibrosis that ultimately lead to diabetic nephropathy.2 Although oxidative stress can result from a decrease in local antioxidant capacity,3 it seems that the more typical scenario involves excess production of superoxide anion (O2•−), which initiates a series of reactions that generate other reactive oxygen species (ROS) including peroxynitrite, hydrogen peroxide, hydroxyl radical and hypochlorous acid. The primary site of O2•− production within the normal rat kidney is the thick ascending limb,4 and we recently provided evidence that T1D causes a protein kinase C (PKC)-dependent increase in O2•− production by the medullary thick ascending limb (mTAL);5 however, the mechanism underlying this phenomenon has not yet been determined.

The major renal sources of O2•− are mitochondrial respiratory chain enzymes and nonphagocytic NAD(P)H oxidase.6,7 NAD(P)H oxidase is similar to the NADPH oxidase responsible for the neutrophil respiratory burst; however, it can use either NADH or NADPH as substrate and can generate O2•− for longer periods but at rates somewhat lower than the phagocytic NADPH oxidase. NAD(P)H oxidase consists of multiple subunits: a Nox family member (the catalytic subunit), p22phox, p47phox, p67phox, p40phox and rac1. Seven Nox family members (Nox1–5; Duox1–2) have been detected in nonphagocytic cells, and have 27–58% sequence similarity to Nox2 (gp91phox; expressed in phagocytic cells and a variety of nonphagocytic cells).1,8 The Nox family member(s) expressed by specific cell types and the functional relevance of different Nox proteins are still being elaborated. It is at least clear that NAD(P)H oxidase activity can be regulated by expression of the subunits, as well as by p47phox phosphorylation (generally triggered by agonist stimulation). Evidence available to date implicates NAD(P)H oxidase as a source of excess renal and vascular O2•− production during T1D,1,8,9 a process that may result from increased expression of various NAD(P)H oxidase subunits9–11 and/or the ability of T1D to induce activation of various PKC isoforms.5,12–15 Indeed, PKC activation is required for translocation of p47phox and p67phox, and subsequent O2•− production by NAD(P)H oxidase in glomeruli from diabetic rats.10 However, at present, little is known with regard to T1D-induced changes in the expression and activity of NAD(P)H oxidase in the mTAL or the role of this enzyme in the PKC-dependent acceleration of O2•− production by this nephron segment during T1D. Therefore, we hypothesized that increased O2•− production in the mTAL during T1D results from PKC-dependent NAD(P)H oxidase activation.

Methods

Induction of type 1 diabetes

All animal procedures were approved by the University of Nebraska Medical Center Institutional Animal Care and Use Committee and conducted in accord with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Male Sprague-Dawley rats weighing ~300 g (Harlan) were anesthetized with methohexital sodium (50 mg/kg IP) to facilitate IV injection of 65 mg/kg streptozotocin (STZ rats) or vehicle (ice-cold PBS; pH 4; Sham rats). The following day, the rats were anesthetized again for SC insertion of a 2.3×2.0 mm sustained-release insulin pellet (Linplant®, Linshin Canada; STZ rats) or vehicle pellet (Sham rats) via a 16 gauge needle. The rats were provided ad libitum food and water for the ensuing 3–4 wks while housed in a temperature controlled room with a 12:12 h light:dark cycle. During this period, STZ rats gained less weight (33±10 g) than Sham rats (90±5 g). Blood glucose levels were measured with an Accu-Chek® Advantage® kit (Roche Diagnostics) prior to STZ or vehicle injection and twice weekly thereafter, with values averaging 21.0±0.7 mmol/L in STZ rats (n=24) and 5.0±0.1 mmol/L in Sham rats (n=28).

Preparation of mTAL suspensions

Fresh mTAL suspensions were prepared from the inner stripe of the outer medulla according to the method of Garvin and colleagues,16,17 with slight modification as previously described.5 The mTALs were suspended in cold HBSS containing either 5.5 or 20 mmol/L D-glucose (for Sham or STZ rats, respectively, to maintain the chronic in vivo condition of the donor rats), equilibrated with 95% O2-5% CO2 and adjusted to pH 7.4. Suspensions were kept on ice until initiating any in vitro treatment (30 min incubation at 37°C in the absence or presence of a pharmacological agent) or tissue homogenization for biochemical assays.

Measurement of O2•− production in mTAL suspensions

mTAL suspensions in polystyrene tubes were placed in the chamber of a Berthold Sirius tube luminometer to obtain a blank value. Lucigenin (final concentration=5 μmol/L) was added to the sample and, after a 2 min dark adaptation period, the luminescence signal (relative light units/sec; RLU/sec) was averaged in 30 sec blocks for the ensuing 5 min. Blank-corrected values obtained during the final 90 sec of this period, normalized per mg protein, were used for comparisons. We previously demonstrated the tempol sensitivity of this parameter,5 thus indicating that the data primarily reflect O2•− production.

NAD(P)H oxidase activity assay

mTALs were pelleted by gentle centrifugation, the supernatant decanted, and 100 μl homogenizing buffer was mixed with the pellet. The homogenizing buffer consisted of 20 mmol/L KH2PO4 and 1 mmol/L EGTA (pH 7.8) with EDTA-free protease inhibitor cocktail (1:100; Thermo Scientific). After homogenization on ice (50 strokes with a Kontes PELLET PESTLE® Micro Grinder), homogenates were centrifuged at 1,000 g for 15 min to remove unbroken cells and debris. An aliquot of the supernatant (50 μg protein) was added to a polystyrene tube containing HBSS with lucigenin and NADPH provided to achieve final concentrations of 5 and 100 μmol/L, respectively. After a dark adaptation period, emitted light (RLU/sec) was detected in the luminometer, averaged during the final 90 sec of a 5 min measurement sequence. Lucigenin chemiluminescence in this assay required exogenous NADPH, with minimal signal evident if NADH was substituted for NADPH. Hence, we report NAD(P)H oxidase activity as RLU/sec/μg protein measured in the presence of 100 μmol/L NADPH.

Western blot

Homogenates prepared from mTAL suspensions were prepared and utilized for western blot analysis according to methods previously detailed,5 except that we used Immobilon-P PVDF transfer membranes (Millipore) for p47phox detection and nitrocellulose membranes (Thermo/Pierce) for Nox2 and Nox4 detection. The following primary antibodies were utilized: 1:200 anti-Nox2/gp91phox (BD Transduction; #611414), 1:200 anti-Nox4 (Santa Cruz; sc-21860), 1:200 anti-p47phox (Santa Cruz; sc-7660-R) and 1:5000 anti-β-actin (Abcam; ab8226 or ab8227). We used secondary antibodies conjugated to infrared dyes, allowing signal detection using a LI-COR Odyssey® Imager. For quantification, band intensities were normalized to β-actin and expressed as a % of averaged sham values run on the same gel.

Statistical analysis

All data are reported as means±SEM, with n values representing the number of rats. Statistical comparisons utilized the unpaired t-test or ANOVA with post-hoc comparisons made using the Student-Newman-Keuls method. If the data were not normally distributed, the Kruskal-Wallis ANOVA on Ranks was employed, with post-hoc comparisons utilizing the Holm-Sidek method. P values <0.05 were considered significant.

Results

NAD(P)H oxidase inhibition normalizes mTAL O2•− production during T1D

Consistent with our previous study,5 O2•− production measured by lucigenin chemiluminescence in mTAL suspensions from Sham rats averaged 430±31 RLU/sec/mg protein and was accelerated 5-fold in mTALs from STZ rats (P<0.05; Figure 1). Preincubation with the NAD(P)H oxidase inhibitor apocynin (100 μmol/L; Calbiochem) decreased O2•− production by 80% in STZ mTALs, but had no effect on Sham mTALs. Thus, NAD(P)H oxidase inhibition reversed the increase in O2•− production otherwise apparent in mTAL suspensions from STZ rats.

Figure 1.

Figure 1

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O2•− production by mTAL suspensions under untreated conditions (n=12 per group) and after 30 min pre-treatment with 100 μmol/L apocynin (NAD(P)H oxidase inhibitor; n=8 per group). *P<0.05 vs Sham; †P<0.05 vs Untreated.

PKC-dependent NAD(P)H oxidase activation in the mTAL during T1D

As shown in Figure 2, NAD(P)H oxidase activity in STZ mTALs was more than 2-fold higher than that in Sham mTALs (P<0.05). Pretreatment of mTAL suspensions with 1 μmol/L calphostin C (a broad-spectrum PKC inhibitor with an IC50 of 0.05 μmol/L)18 prior to tissue homogenization reduced NAD(P)H oxidase by ~80% in both groups (P<0.05 vs Untreated). Calphostin C-sensitive (PKC-dependent) NAD(P)H oxidase activity calculated from these data was significantly greater in STZ mTALs (45.8±9.6 RLU/sec/μg protein) than in Sham mTALs (19.0±2.0 RLU/sec/μg protein). These data reveal that NAD(P)H oxidase activity is increased in the rat mTAL during T1D and this phenomenon is PKC-dependent.

Figure 2.

Figure 2

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Effects of T1D and PKC inhibition on NAD(P)H oxidase activity in the mTAL. mTAL suspensions from Sham (n=7) and STZ rats (n=8) were left untreated or pretreated 30 min with 1 μmol/L calphostin C (broad-spectrum PKC inhibitor), 1 μmol/L Gö6976 (PKCα/β inhibitor), 50 nmol/L indolylmalemide-1 (PKCβ inhibitor) or 10 μmol/L rottlerin (PKCδ inhibitor) prior to measurement of NAD(P)H oxidase activity. *P<0.05 vs Sham; †P<0.05 vs Untreated.

To determine which PKC isoform is involved in the accelerated NAD(P)H oxidase activity accompanying T1D, NAD(P)H oxidase activity was assayed after pretreatment of mTAL suspensions with PKC inhibitors having relative isoform selectivity: 1 μmol/L Gö6976 (PKCα/β inhibitor, IC50=1.3–6 nmol/L),19 50 nmol/L indolylmaleimide-1 (PKCβ inhibitor marketed by Calbiochem; IC50=5–21 nmol/L),20 or 10 μmol/L rottlerin (PKCδ inhibitor; IC50=3.6 μmol/L).21 We previously confirmed that none of these agents adversely alter mTAL viability under these experimental conditions.5 As shown in Figure 2, Gö6976 and indolylmaleimide-1 had no effect on NAD(P)H oxidase activity in mTALs from Sham or STZ rats. However, rottlerin significantly reduced NAD(P)H oxidase activity by ~80% in both Sham and STZ mTALs (P<0.05 vs Untreated for both groups). Rottlerin-sensitive (PKCδ-dependent) NAD(P)H oxidase activity calculated from these data was significantly higher in STZ mTALs (45.6±10.8 RLU/sec/μg protein) than in Sham mTALs (18.1±2.1 RLU/sec/μg protein). Thus, the T1D-induced increase in PKCδ-dependent NAD(P)H oxidase activity is virtually identical in magnitude to the increase in PKC-dependent NAD(P)H oxidase activity, and appears to underlie the increase in total NAD(P)H oxidase activity evident under these conditions.

T1D increases Nox2, Nox4 and p47phox levels in the mTAL

Western blot was used to detect NAD(P)H oxidase subunit protein levels in mTALs from Sham and STZ rats. As shown in Figure 3, Nox2, Nox4 and p47phox protein levels were significantly increased in mTALs from STZ rats, achieving values averaging 161±21, 134±14 and 130±9% of Sham, respectively. Thus, T1D triggers upregulation of these NAD(P)H oxidase subunits in the rat mTAL.

Figure 3.

Figure 3

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Effect of T1D on NAD(P)H oxidase subunit protein levels in the mTAL. Shown are representative western blots and densitometric analysis of Nox2 (Sham n=5; STZ n=7), Nox4 (n=8 per group), and p47phox (n=16 per group) levels in mTALs from Sham and STZ rats. *P<0.05 vs Sham.

Discussion

In humans, systemic oxidative stress is evident at onset of T1D in children and adolescents, and is increased by early adulthood.22 Thus, the mechanisms underlying renal oxidative stress in the early stage of T1D likely have significant pathophysiological relevance. Accordingly, our study focused on events evident 3–4 weeks after induction of T1D in rats by STZ injection, with moderate hyperglycemia achieved by partial insulin replacement. These animals display renal hypertrophy and hyperfiltration, as well as increased urinary excretion of TGF-β (a profibrotic molecule), H2O2, thiobarbituric acid reactive substances, and 8-hydroxy-2′deoxyguanosine (indicators of oxidative stress); however, only moderate microalbuminuria is evident at this time point.23 Insulin replacement or pancreatic beta cell transplantation to achieve euglycemia prevents the multiple renal functional changes evident in STZ rats,24,25 thus ruling out the possibility that these phenomena represent a direct renal toxic effect of STZ. We recently reported increased O2•− production in the mTAL of STZ rats, and that this phenomenon is PKC-dependent.5 The increased O2•− production by STZ mTALs seems to be initiated by the hyperglycemic milieu, as acute (30 min) exposure of mTALs from normal rat kidney to 20 mM glucose is sufficient to accelerate O2•− production 2-fold.5 The results of the present study reveal that apocynin reverses excess O2•− production in mTAL suspensions from STZ rats. Moreover, increased expression of NAD(P)H oxidase subunits (Nox2, Nox4, and p47phox) was evident in STZ mTALs, in concert with increased NAD(P)H oxidase activity. Furthermore, PKC inhibition (either broad-spectrum or PKCδ-specific) abolished the increase in NAD(P)H oxidase activity. These data implicate PKC-dependent NAD(P)H oxidase activation, together with increased subunit expression, in the accelerated O2•− production that arises in the mTAL during the early stage of T1D.

The present study employed apocynin (100 μmol/L) to determine the contribution of NAD(P)H oxidase to O2•− production by mTAL suspensions. In addition to its ability to inhibit NADPH oxidase in phagocytic cells, apocynin can act as an antioxidant (scavenging either H2O2 or •OH), but only at concentrations exceeding that used in the present study.26 Moreover, apocynin can interfere with O2•− detection when using high lucigenin concentrations known to undergo redox cycling; however, this phenomenon is not evident at the low (5 μmol/L) lucigenin concentration used in the present study.26 Finally, activation of apocynin requires H2O2 and myeloperoxidase,26,27 which is expressed primarily in leukocytes. We are unaware of any evidence that this enzyme is expressed in the mTAL; however, myeloperoxidase uptake by mTAL cells may occur subsequent to its release by infiltrating inflammatory cells, similar to processes described in pulmonary epithelial cells,28 and remain available in freshly-prepared mTAL suspensions. Alternatively, as apocynin can be activated in vitro by co-incubation with H2O2 and horseradish peroxidase,29 other peroxidases expressed by the mTAL might be capable of activating apocynin. Thus, reasonable scenarios exist to provide for apocynin activation in mTALs under the conditions of our study. Thus, while we cannot absolutely rule out the possibility that apocynin exerted non-specific effects in our experimental setting, the ability of apocynin to normalize lucigenin chemiluminescence in mTAL suspensions from STZ rats supports the contention that NAD(P)H oxidase is a primary source of increased O2•− production by the mTAL during T1D.

In contrast with the effects observed in STZ mTALs, apocynin had no effect on O2•− production in mTALs from Sham rats. This observation differs somewhat from the results of Li and coworkers,4 who concluded that NAD(P)H oxidase is a major enzyme responsible for O2•− production by the thick ascending limb from normal rat kidney. Their conclusion was based on dihydroethidium fluorescence responses to diphenyleneiodonium (DPI) in the cortical thick ascending limb. It is possible that cortical and medullary portions of the thick ascending limb have different enzymatic sources of O2•−, or that the effects of DPI in this nephron segment reflect its ability to inhibit not only NAD(P)H oxidase but also other flavin-containing enzymes that also have the capacity to generate O2•− (i.e. NO synthase). In contrast, apocynin acts by binding to p47phox, preventing the assembly of NAD(P)H subunits that is required for enzymatic activity. The lack of effect of apocynin on mTAL suspensions from Sham rats suggests that enzymes other than NAD(P)H oxidase may be responsible for O2•− production in the absence of agonist stimulation or a pathophysiological processes. It is also possible that O2•− production by mTALs from Sham rats reflects the functional impact of Nox4, which is capable of constitutively generating O2•− in the absence of the various regulatory subunits of NAD(P)H oxidase30 and, thus, is unlikely to be influenced by apocynin.

NAD(P)H oxidase activation has been demonstrated in renal cortex31,32 and glomeruli10 of STZ rats. Consistent with these findings, we found that NAD(P)H oxidase activity was increased significantly in STZ mTALs. Depending on the Nox family member serving as the catalytic subunit, NAD(P)H oxidase activity can be regulated by subunit expression and/or cytosolic activator proteins. Both of these possibilities were explored in the present study. As PKC activity in the mTAL is increased during T1D, and PKC inhibition markedly attenuates O2•− production in mTAL suspensions from STZ rats,5 we assessed the involvement of PKC in the T1D-induced NAD(P)H oxidase activation using a pharmacological approach. The effect of calphostin C to markedly reduce NAD(P)H oxidase activity in mTALs from Sham and STZ rats confirms the key involvement of PKC as a determinant of NAD(P)H oxidase activity in these cells. We also found that calphostin C-sensitive NAD(P)H oxidase activity was increased in STZ mTALs compared with Sham, indicating a role for PKC in the NAD(P)H oxidase activation accompanying T1D. These observations are consistent with evidence that high glucose levels can stimulate ROS production through PKC-dependent activation of NAD(P)H oxidase in vascular smooth muscle and endothelial cells.33

Glucose- or diabetes-induced NAD(P)H oxidase activation has been shown to be reliant on PKCα, PKCβ, PKCδ, or PKCζ, depending on the tissue studied.8 In the mTAL, T1D increases expression of PKCα and PKCβ, and evidence suggests that both are activated in STZ rats.5 Moreover, PKCδ activity (based on cytosol vs. membrane distribution) is substantial in mTALs under normal and diabetic conditions.5 To determine which isoform is involved in the NAD(P)H oxidase activation in STZ mTALs, we utilized PKC inhibitors with relative isoform specificity. The results revealed the ability of PKCδ inhibition to decrease NAD(P)H oxidase activity in both STZ and Sham mTALs, while inhibition of PKCα and/or β had no effect. Moreover, PKCδ-dependent NAD(P)H oxidase activity was much higher in STZ mTALs than in Sham mTALs. These data indicate that PKCδ (but not PKCα or PKCβ) is critical for both basal and T1D-stimulated NAD(P)H oxidase activity in mTAL. Interestingly, our previous investigation provided evidence that increased O2•− production by intact STZ mTALs was dependent on PKCα and PKCδ;5 however, results of the present study implicate PKCδ (but not PKCα) in the T1D-induced NAD(P)H oxidase activation measured in mTAL homogenates. The O2•− production assay performed in intact cells relies solely on endogenous cofactors and substrate availability, while the NAD(P)H oxidase activity assay utilizes cell homogenates with the provision of excess substrate (NADPH), so it is difficult to make a direct comparison of the results. However, it seems that PKCα influences O2•− production during T1D through an NAD(P)H oxidase-independent mechanism, perhaps through effect on NO synthase, xanthine oxidase, or the mitochondrial respiratory chain enzymes.

Based on our accumulated observations, it is likely that the tonically high level of PKCδ activity in the mTAL contributes to O2•− production via effects on NAD(P)H oxidase activation. In the setting of T1D, in which there is no apparent change in PKCδ protein level or translocation (activity) in the mTAL,5 increased PKCδ-dependent NAD(P)H oxidase activity most likely results from an increase in the availability of PKCδ substrate. This scenario is consistent with our observation that mTALs from STZ rats express elevated protein levels of p47phox, a known substrate of PKCδ.34 Given that p47phox phosphorylation is key to subunit assembly, and that this process requires PKCδ,34 we postulate that the combination of high basal PKCδ activity and increased p47phox protein levels promotes increased NAD(P)H oxidase activation in the mTAL during T1D. The impact on enzyme activity is likely facilitated by the concomitant increase in protein levels of the catalytic subunit Nox2. The observation of increased mTAL expression of Nox2 and p47phox is consistent with reported changes evident at the whole kidney level that were prevented by chronic apocynin treatment.9 This observation suggests that oxidative stress in T1D provokes a positive-feedback increase in NAD(P)H oxidase subunit expression.

Interestingly, mTALs express at least two different Nox family members, Nox2 and Nox4, and protein levels of both are elevated in T1D. An increase in Nox4 protein levels in the mTAL during T1D is consistent with reports of similar changes in kidney cortex and isolated glomeruli.32 The T1D-induced increase in Nox4 protein levels should contribute to increased NAD(P)H oxidase activity and O2•− production, independent of changes in expression or phosphorylation of regulatory subunits. Nox4 expression is subject to regulation by PKCα,35 which is upregulated in the mTAL during diabetes.5 Thus, PKC-dependent regulation of NAD(P)H oxidase activity and O2•− production during diabetes may involve both pre- and post-translational mechanisms.

Perspectives

The initial renal structural alteration in T1D is glycogen deposition in the epithelial cells of the thick ascending loop of Henle,36 which leads to decreased production of Tamm-Horsfall protein (THP) by these cells. Recent evidence indicates that decreased urinary THP concentration in T1D is associated with an 8-fold increased risk of cardiovascular death and uremia.37 This observation underscores the importance of understanding the effects of T1D on mTAL function; however, surprisingly little investigation has focused on this topic. The product of NAD(P)H oxidase, O2•−, can influence Na+ transport by the normal mTAL, exerting a stimulatory effect that counteracts the inhibitory impact of NO.38,39 Our recent work indicates that NO bioavailability in the mTALs is reduced during T1D, and that O2•− scavenging with tempol unmasks increased NO production under these conditions.40 Therefore, it is likely that the mechanisms underlying NAD(P)H oxidase activation during T1D exert an impact on Na+ balance and extracellular fluid volume homeostasis, possibly favoring volume retention with an attendant link to hypertension. Moreover, antioxidant therapy that minimizes the T1D-induced increased in NAD(P)H oxidase activity may curtail the development of tubulointerstitial fibrosis, which ultimately contributes substantially to diabetic nephropathy. Further investigation will be necessary to unveil the systemic and chronic consequences of NAD(P)H oxidase activation in the mTAL during diabetes.

Acknowledgments

The authors gratefully acknowledge the skilled technical support of Rachel W. Fallet.

Sources of Funding

This work was funded by National Institutes of Health grant DK063416 (to P.K.C. and J.S.P.). J. Yang was supported by a research assistantship from the Graduate Studies Office at the University of Nebraska Medical Center.

Footnotes

Disclosures

None

References

  • 1.Li JM, Shah AM. ROS generation by nonphagocytic NADPH oxidase: potential relevance in diabetic nephropathy. J Am Soc Nephrol. 2003;14:S221–S226. doi: 10.1097/01.asn.0000077406.67663.e7. [DOI] [PubMed] [Google Scholar]
  • 2.Forbes JM, Coughlan MT, Cooper ME. Oxidative stress as a major culprit in kidney disease in diabetes. Diabetes. 2008;57:1446–1454. doi: 10.2337/db08-0057. [DOI] [PubMed] [Google Scholar]
  • 3.West IC. Radicals and oxidative stress in diabetes. Diabet Med. 2000;17:171–180. doi: 10.1046/j.1464-5491.2000.00259.x. [DOI] [PubMed] [Google Scholar]
  • 4.Li N, Yi F-X, Spurrier JL, Bobrowitz CA, Zou A-P. Production of superoxide through NADH oxidase in thick ascending limb of Henle’s loop in rat kidney. Am J Physiol Renal Physiol. 2002;282:F1111–F1119. doi: 10.1152/ajprenal.00218.2001. [DOI] [PubMed] [Google Scholar]
  • 5.Yang J, Lane PH, Pollock JS, Carmines PK. PKC-dependent superoxide production by the renal medullary thick ascending limb from diabetic rats. Am J Physiol Renal Physiol. 2009;297:F1220–F1228. doi: 10.1152/ajprenal.00314.2009. [DOI] [PubMed] [Google Scholar]
  • 6.Zou A-P, Li N, Cowley AW., Jr Production and actions of superoxide in the renal medulla. Hypertension. 2001;37:547–553. doi: 10.1161/01.hyp.37.2.547. [DOI] [PubMed] [Google Scholar]
  • 7.Vaziri ND. Roles of oxidative stress and antioxidant therapy in chronic kidney disease and hypertension. Curr Opin Nephrol Hypertens. 2004;13:93–99. doi: 10.1097/00041552-200401000-00013. [DOI] [PubMed] [Google Scholar]
  • 8.Gao L, Mann GE. Vascular NAD(P)H oxidase activation in diabetes: a double-edged sword in redox signalling. Cardiovasc Res. 2009;82:9–20. doi: 10.1093/cvr/cvp031. [DOI] [PubMed] [Google Scholar]
  • 9.Asaba K, Tojo A, Onozato ML, Goto A, Quinn MT, Fujita T, Wilcox CS. Effects of NADPH oxidase inhibitor in diabetic nephropathy. Kidney Int. 2005;67:1890–1898. doi: 10.1111/j.1523-1755.2005.00287.x. [DOI] [PubMed] [Google Scholar]
  • 10.Kitada M, Koya D, Sugimoto T, Isono M, Araki S, Kashiwagi A, Haneda M. Translocation of glomerular p47phox and p67phox by protein kinase C-β activation is required for oxidative stress in diabetic nephropathy. Diabetes. 2003;52:2603–2614. doi: 10.2337/diabetes.52.10.2603. [DOI] [PubMed] [Google Scholar]
  • 11.Etoh T, Inoguchi T, Kakimoto M, Sonoda N, Kobayashi K, Kuroda J, Sumimoto H, Nawata H. Increased expression of NAD(P)H oxidase subunits, NOX4 and p22phox, in the kidney of streptozotocin-induced diabetic rats and its reversibity by interventive insulin treatment. Diabetologia. 2003;46:1428–1437. doi: 10.1007/s00125-003-1205-6. [DOI] [PubMed] [Google Scholar]
  • 12.Igarashi M, Wakasaki H, Takahara N, Ishii H, Jiang ZY, Yamauchi T, Kuboki K, Meier M, Rhodes CJ, King GL. Glucose or diabetes activates p38 mitogen-activated protein kinase via different pathways. J Clin Invest. 1999;103:185–195. doi: 10.1172/JCI3326. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Craven PA, DeRubertis FR. Protein kinase C is activated in glomeruli from streptozotocin diabetic rats. Possible mediation by glucose. J Clin Invest. 1989;83:1667–1675. doi: 10.1172/JCI114066. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Inoguchi T, Xia P, Kunisaki M, Higashi S, Feener EP, King GL. Insulin’s effect on protein kinase C and diacylglycerol induced by diabetes and glucose in vascular tissues. Am J Physiol Endocrinol Metab. 1994;267:E369–E379. doi: 10.1152/ajpendo.1994.267.3.E369. [DOI] [PubMed] [Google Scholar]
  • 15.Koya D, Jirousek MR, Lin YW, Ishii H, Kuboki K, King GL. Characterization of protein kinase C β isoform activation on the gene expression of transforming growth factor-β, extracellular matrix components, and prostanoids in the glomeruli of diabetic rats. J Clin Invest. 1997;100:115–126. doi: 10.1172/JCI119503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Silva G, Beierwaltes WH, Garvin JL. Extracellular ATP stimulates NO production in rat thick ascending limb. Hypertension. 2006;47:563–567. doi: 10.1161/01.HYP.0000197954.93874.ef. [DOI] [PubMed] [Google Scholar]
  • 17.Varela M, Herrera M, Garvin JL. Inhibition of Na-K-ATPase in thick ascending limbs by NO depends on O2− and is diminished by a high-salt diet. Am J Physiol Renal Physiol. 2004;287:F224–F230. doi: 10.1152/ajprenal.00427.2003. [DOI] [PubMed] [Google Scholar]
  • 18.Keenan C, Goode N, Pears C. Isoform specificity of activators and inhibitors of protein kinase C γ and δ. FEBS Lett. 1997;415:101–108. doi: 10.1016/s0014-5793(97)01104-6. [DOI] [PubMed] [Google Scholar]
  • 19.Martiny-Baron G, Kazanietz MG, Mischak H, Blumberg PM, Kochs G, Hug H, Marme D, Schachtele C. Selective inhibition of protein kinase C isozymes by the indolocarbazole Gö6976. J Biol Chem. 1993;268:9194–9197. [PubMed] [Google Scholar]
  • 20.Tanaka M, Sagawa S, Hoshi Ji, Shimoma F, Matsuda I, Sakoda K, Sasase T, Shindo M, Inaba T. Synthesis of anilino-monoindolylmaleimides as potent and selective PKCβ inhibitors. Bioorg Med Chem Lett. 2004;14:5171–5174. doi: 10.1016/j.bmcl.2004.07.061. [DOI] [PubMed] [Google Scholar]
  • 21.Gschwendt M, Muller HJ, Kielbassa K, Zang R, Kittstein W, Rincke G, Marks F. Rottlerin, a novel protein kinase inhibitor. Biochem Biophys Res Commun. 1994;199:93–98. doi: 10.1006/bbrc.1994.1199. [DOI] [PubMed] [Google Scholar]
  • 22.Dominguez C, Ruiz E, Gussinye M, Carrascosa A. Oxidative stress at onset and in early stages of type 1 diabetes in children and adolescents. Diabetes Care. 1998;21:1736–1742. doi: 10.2337/diacare.21.10.1736. [DOI] [PubMed] [Google Scholar]
  • 23.Sasser JM, Sullivan JC, Hobbs JL, Yamamoto T, Pollock DM, Carmines PK, Pollock JS. Endothelin A receptor blockade reduces diabetic renal injury via an anti-inflammatory mechanism. J Am Soc Nephrol. 2007;18:143–154. doi: 10.1681/ASN.2006030208. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Lee DL, Sasser JM, Hobbs JL, Boriskie A, Pollock DM, Carmines PK, Pollock JS. Posttranslational regulation of NO synthase activity in the renal medulla of diabetic rats. Am J Physiol Renal Physiol. 2005;288:F82–F90. doi: 10.1152/ajprenal.00127.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Palm F, Ortsäter H, Hansell P, Liss P, Carlsson PO. Differentiating between effects of streptozotocin per se and subsequent hyperglycemia on renal function and metabolism in the streptozotocin-diabetic rat model. Diabetes Metab Res Rev. 2004;20:452–459. doi: 10.1002/dmrr.472. [DOI] [PubMed] [Google Scholar]
  • 26.Heumüller S, Wind S, Barbosa-Sicard E, Schmidt HH, Busse R, Schroder K, Brandes RP. Apocynin is not an inhibitor of vascular NADPH oxidases but an antioxidant. Hypertension. 2008;51:211–217. doi: 10.1161/HYPERTENSIONAHA.107.100214. [DOI] [PubMed] [Google Scholar]
  • 27.Simons JM, Hart BA, Ip Vai Ching TR, Van DH, Labadie RP. Metabolic activation of natural phenols into selective oxidative burst agonists by activated human neutrophils. Free Radic Biol Med. 1990;8:251–258. doi: 10.1016/0891-5849(90)90070-y. [DOI] [PubMed] [Google Scholar]
  • 28.Haegens A, Vernooy JHJ, Heeringa P, Mossman BT, Wouters EFM. Myeloperoxidase modulates lung epithelial responses to pro-inflammatory agents. Eur Respir J. 2008;31:252–260. doi: 10.1183/09031936.00029307. [DOI] [PubMed] [Google Scholar]
  • 29.Vejrazka M, Micek R, Stipek S. Apocynin inhibits NADPH oxidase in phagocytes but stimulates ROS production in non-phagocytic cells. Biochim Biophys Acta. 2005;1722:143–147. doi: 10.1016/j.bbagen.2004.12.008. [DOI] [PubMed] [Google Scholar]
  • 30.Brandes RP, Schroder K. Differential vascular functions of Nox family NADPH oxidases. Curr Opin Lipidol. 2008;19:513–518. doi: 10.1097/MOL.0b013e32830c91e3. [DOI] [PubMed] [Google Scholar]
  • 31.Block K, Gorin Y, Abboud HE. Subcellular localization of Nox4 and regulation in diabetes. Proc Natl Acad Sci U S A. 2009;106:14385–14390. doi: 10.1073/pnas.0906805106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Gorin Y, Block K, Hernandez J, Bhandari B, Wagner B, Barnes JL, Abboud HE. Nox4 NAD(P)H oxidase mediates hypertrophy and fibronectin expression in the diabetic kidney. J Biol Chem. 2005;280:39616–39626. doi: 10.1074/jbc.M502412200. [DOI] [PubMed] [Google Scholar]
  • 33.Inoguchi T, Li P, Umeda F, Yu HY, Kakimoto M, Imamura M, Aoki T, Etoh T, Hashimoto T, Naruse M, Sano H, Utsumi H, Nawata H. High glucose level and free fatty acid stimulate reactive oxygen species production through protein kinase C-dependent activation of NAD(P)H oxidase in cultured vascular cells. Diabetes. 2000;49:1939–1945. doi: 10.2337/diabetes.49.11.1939. [DOI] [PubMed] [Google Scholar]
  • 34.Bey EA, Xu B, Bhattacharjee A, Oldfield CM, Zhao X, Li Q, Subbulakshmi V, Feldman GM, Wientjes FB, Cathcart MK. Protein kinase Cδ is required for p47phox phosphorylation and translocation in activated human monocytes. J Immunology. 2004;174:5730–5738. doi: 10.4049/jimmunol.173.9.5730. [DOI] [PubMed] [Google Scholar]
  • 35.Xu H, Goettsch C, Xia N, Horke S, Morawietz H, Forstermann U, Li H. Differential roles of PKCα and PKCε in controlling the gene expression of Nox4 in human endothelial cells. Free Radic Biol Med. 2008;44:1656–1667. doi: 10.1016/j.freeradbiomed.2008.01.023. [DOI] [PubMed] [Google Scholar]
  • 36.Rasch R, Torffvit O, Bachmann S, Jensen PK, Jacobsen NO. Tamm-Horsfall glycoprotein in streptozotocin diabetic rats: A study of kidney in situ hybridization, immunohistochemistry, and urinary excretion. Diabetologia. 1995;38:525–535. doi: 10.1007/BF00400720. [DOI] [PubMed] [Google Scholar]
  • 37.Sejdiu I, Torffvit O. Decreased urinary concentration of Tamm-Horsfall protein is associated with development of renal failure and cardiovascular death within 20 years in type 1 but not in type 2 diabetic patients. Scand J Urol Nephrol. 2008;42:168–174. doi: 10.1080/00365590701644691. [DOI] [PubMed] [Google Scholar]
  • 38.Ortiz PA, Garvin JL. Interaction of O2− and NO in the thick ascending limb. Hypertension. 2002;39:591–596. doi: 10.1161/hy0202.103287. [DOI] [PubMed] [Google Scholar]
  • 39.Ortiz PA, Garvin JL. Superoxide stimulates NaCl absorption by the thick ascending limb. Am J Physiol Renal Physiol. 2002;283:F957–F962. doi: 10.1152/ajprenal.00102.2002. [DOI] [PubMed] [Google Scholar]
  • 40.Foster JM, Carmines PK, Pollock JS. PP2B-dependent NO production in the medullary thick ascending limb during diabetes. Am J Physiol Renal Physiol. 2009;297:F471–F480. doi: 10.1152/ajprenal.90760.2008. [DOI] [PMC free article] [PubMed] [Google Scholar]

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