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Am J Physiol Heart Circ Physiol 288: H1417-H1424, 2005. First published November 11, 2004; doi:10.1152/ajpheart.00559.2004
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Redox regulation of Ito remodeling in diabetic rat heart

Xun Li,1,2 Zhi Xu,1 Shumin Li,1 and George J. Rozanski1

1Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, Nebraska; and 2Department of Cardiology, The First Affiliated Hospital, Soochow University, Soochow, Jiangsu, People's Republic of China

Submitted 15 June 2004 ; accepted in final form 3 November 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Oxidative stress and the resulting change in cell redox state are proposed to contribute to pathogenic alterations in ion channels that underlie electrical remodeling of the diseased heart. The present study examined whether K+ channel remodeling is controlled by endogenous oxidoreductase systems that regulate redox-sensitive cell functions. Diabetes was induced in rats by streptozotocin, and experiments were conducted after 3–5 wk of hyperglycemia. Spectrophotometric assays of ventricular tissue extracts from diabetic rat hearts revealed divergent changes in two major oxidoreductase systems. The thioredoxin (TRX) system in diabetic rat heart was characterized by a 52% decrease in TRX reductase (TRXR) activity from control heart (P < 0.05), whereas TRX activity was 1.7-fold greater than control heart (P < 0.05). Diabetes elicited similar changes in the glutaredoxin (GRX) system: glutathione reductase was decreased 35% from control level (P < 0.05), and GRX activity was 2.5-fold greater than in control heart (P < 0.05). The basal activity of glucose-6-phosphate dehydrogenase, which generates NADPH required by the TRX and GRX systems, was not altered by diabetes. Voltage-clamp studies showed that the characteristically decreased density of the transient outward K+ current (Ito) in isolated diabetic rat myocytes was normalized by in vitro treatment with insulin (0.1 µM) or the metabolic activator dichloroacetate (1.5 mM). The effect of these agonists on Ito was blocked by inhibitors of glucose-6-phosphate dehydrogenase. Moreover, inhibitors of TRXR, which controls the reducing activity of TRX, also blocked upregulation of Ito by insulin and dichloroacetate. These data suggest that K+ channels underlying Ito are regulated in a redox-sensitive manner by the TRX system and the remodeling of Ito that occurs in diabetes may be due to decreased TRXR activity. We propose that oxidoreductase systems are an important repair mechanism that protects ion channels and associated regulatory proteins from irreversible oxidative damage.

insulin; transient outward current; potassium; thioredoxin; glutaredoxin

CARDIOVASCULAR COMPLICATIONS associated with diabetes mellitus have a significant impact on the clinical prognosis of patients with this disorder (7). In addition to vascular dysfunction, a distinct cardiomyopathy has been attributed to diabetes that includes profound electrical remodeling of the ventricle (5, 26, 32, 35, 36). This pathogenic change in electrophysiological phenotype likely contributes to the increased incidence of cardiac arrhythmias and sudden death in this patient population (7, 9) and is associated with marked changes in the ECG such as T-wave abnormalities and QT-interval prolongation (9). Experimental models of diabetes have shown consistent changes in action potential configuration that may help explain the clinically observed electrocardiographic changes, particularly a longer action potential duration (APD; Refs. 5, 17, 31). Voltage-clamp experiments suggest that diabetes-induced APD prolongation is due partly to a decrease in outward repolarizing current such as the Ca2+-independent transient outward K+ current (Ito) and a quasi-steady-state current (5, 26, 31, 32, 35, 36), which also control APD in human ventricular myocytes (20). Some of these changes in ion channel activity have been correlated with significant decreases in mRNA of specific K+ channel genes and abundance of their encoded proteins (26).

Results from clinical and experimental studies suggest that diabetic cardiomyopathy is linked to a multitude of metabolic disturbances and increased production of reactive oxygen species that together underlie pathophysiological changes in myocyte function (18, 28). Recent data from our laboratory (29, 35, 36) and others' work (32) support the hypothesis that depressed glucose metabolism is a key factor underlying changes in K+ channel function in myocytes of hearts from streptozotocin (STZ)-induced diabetic rats. In particular, treatment of isolated ventricular myocytes from diabetic rats with insulin or related activators of glucose utilization has been shown to upregulate K+ current density to control levels and may involve synthesis of new channel protein (32). Furthermore, our most recent studies suggest that the consequences of diabetes-related hyperglycemia and impaired glucose metabolism on cardiac K+ channel function involve alterations in cell redox state (29, 34). Consistent with these findings are studies in a variety of tissues that demonstrate that diabetic conditions are associated with a more oxidized cellular environment and damage to key cell proteins (2, 6).

An important functional outcome of oxidative stress in the myocardium is a shift in the redox state of cellular proteins, which is characterized by the formation of several molecular intermediates of the sulfhydryl group of cysteine residues (24). Indeed, sulfhydryl biochemistry has been shown to play an important role in regulation of cell function, because the redox state of cysteine sulfhydryls often determines the structure and activity of enzymes, transcription factors, and transport proteins required for cell viability. Cellular protection against oxidative damage of proteins involves the activities of thiol-disulfide oxidoreductase systems that belong to the thioredoxin (TRX) superfamily (10, 15, 24, 37). TRX and glutaredoxin (GRX) are members of this superfamily that function within the cell to maintain proteins in the reduced state under physiological conditions. The TRX system, which is composed of TRX and TRX reductase (TRXR), predominantly catalyzes the reduction of protein disulfides, whereas the GRX system, made up of GRX, reduced glutathione (GSH), and glutathione reductase (GR), mainly catalyzes the reduction of protein-mixed disulfides (10, 15, 24, 37). Together, these oxidoreductase systems act as sulfhydryl repair mechanisms to control the physiological function of proteins susceptible to oxidation.

The purpose of the present study was to examine the function of endogenous oxidoreductase systems in regulating the electrophysiology of diabetic heart and to identify the components of these systems that underlie diabetes-induced changes in protein redox state. Our results suggest that K+ channels underlying Ito are regulated in a redox-sensitive manner by the TRX system and that the remodeling of Ito that occurs in diabetes is caused by decreased TRXR activity. We propose that the cardiac TRX and GRX systems comprise a functionally important repair mechanism that protects ion channels and associated cellular proteins from irreversible oxidative damage.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The institutional review committee at the University of Nebraska Medical Center approved the animal research protocol used in this study, and the investigation conformed with the Guide for the Care and Use of Laboratory Animals published by National Institutes of Health (NIH Publication No. 85-23, Revised 1996).

Induction of diabetes and isolation of cardiac myocytes. Male Sprague-Dawley rats (body wt, 150–180 g) were made diabetic by a single injection of STZ at a dose of 65 mg/kg ip (34–36). Blood glucose levels were measured 3 days after STZ injection to confirm hyperglycemia. Normal rats of similar age and weight used as controls were injected with vehicle only (1 mM citrate buffer, pH 4.5). Diabetic rats in our study exhibited a nearly fourfold increase in blood glucose concentration compared with control rats [20.4 ± 0.9 (n = 23) vs. 4.9 ± 0.2 mM (n = 38); P < 0.05]. As in our previous studies, body weight and heart weight in the diabetic group were significantly less than for control animals, but the mean heart weight-to-body weight ratio was not different between groups of rats (36).

Three to five weeks after STZ or vehicle injection, rats were given an overdose of pentobarbital sodium (100 mg/kg ip), and single ventricular myocytes were dissociated from excised, perfused hearts by a collagenase digestion procedure that has been described previously (34–36). Dissociated myocytes from both ventricles were suspended in DMEM plus Ham's F-12 (glucose concentration, 18 mM) and stored in an incubator at 37°C until used, which was usually within 6 h of isolation. For the study of ionic currents (see below), aliquots of myocytes were transferred to a cell chamber mounted on the stage of an inverted microscope and superfused with a standard external solution that contained (in mM) 138 NaCl, 4.0 KCl, 1.2 MgCl2, 1.8 CaCl2, 18 glucose, 5 HEPES, pH 7.4 adjusted with NaOH. It should be noted that isolated myocytes from diabetic and control rats were studied at the same concentration of extracellular glucose (18 mM), which approximated the mean blood glucose level measured in diabetic rats (20 mM).

Recording techniques. Ionic currents were recorded using the whole cell configuration of the patch-clamp technique. Briefly, borosilicate glass capillaries were pulled (model P-87, Sutter Instruments), heat polished to an internal tip diameter of 1–2 µm, and filled with a pipette solution that contained (in mM) 135 KCl, 3 MgCl2, 10 HEPES, 3 Na2-ATP, 10 EGTA, and 0.5 Na-GTP, with pH 7.2 adjusted with KOH. Filled pipettes with resistance of 2–4 M{Omega} were coupled to a patch-clamp amplifier (Axopatch 200B; Axon Instruments), and after correction of the liquid junction potential and creation of a gigaohm seal, the external solution was switched to one that contained 0.5 mM CdCl2 to block Ca2+ channels. During the exchange of external solutions, the membrane within the pipette was ruptured, and at least 5 min were allowed for the contents of the pipette and cytoplasm to equilibrate. A computer program (pClamp; Axon Instruments) controlled command potentials and acquired current signals, which were filtered at 2 kHz using a four-pole, low-pass Bessel filter. Currents were sampled at 4 kHz by a 12-bit resolution analog-to-digital converter and were stored on the hard disk of a computer. All electrophysiological experiments were done at room temperature (22–24°C).

Ito was evoked in each cell by 500-ms depolarizing pulses to test potentials between –40 and +60 mV (0.2 Hz). The holding potential in all experiments was –80 mV, and a 100-ms prepulse was applied to –60 mV to inactivate the fast Na+ current. For each test pulse, Ito amplitude was measured as the difference between the peak outward current and the steady-state current level at the end of the depolarizing pulse. Although the steady-state current was decreased in our model of diabetes as in other studies (32), the changes were relatively small compared with Ito. We found that the same results were obtained when Ito was measured as the peak current as for the difference current. Moreover, alterations in Ito due to diabetes were compared with the inward rectifier current, IK1, which was recorded with 100-ms test pulses applied to potentials of –120 to –40 mV from a holding potential of –80 mV. All electrophysiological data were normalized as current densities (in pA/pF) by dividing measured current amplitude by whole cell capacitance.

Enzyme assays. Total activity of TRX was measured by the insulin disulfide reduction assay (11). This assay is based on the ability of TRX to reduce insulin in the presence of NADPH and an excess of TRXR. Briefly, aliquots of extracts from ventricular tissue samples were added to a reaction mixture that contained 47 mM potassium phosphate buffer, 0.1 mM EDTA, 0.2 mM NADPH, and 0.5 mg insulin. The reaction was started by adding 1 U bovine TRXR, and absorbance was read at 340 nm for 5 min at 37°C in a spectrophotometer (ThermoSpectronic). Measured activity was expressed in milliunits (mU) per milligram of protein with 1 mU TRX activity defined as 1 nmol NADPH oxidized per minute.

GRX activity was measured by the hydroxyethyl disulfide reduction assay (27). Aliquots of cell lysate from ventricular tissue samples were added to a reaction mixture that contained 0.2 mM NADPH, 0.5 mM GSH, 0.1 M potassium phosphate buffer (pH 7.4), and 0.4 U GR. The reaction was initiated by adding 2 mM hydroxyethyl disulfide, and the NADPH absorbance at 340 nm was monitored over 5 min. Activity was expressed in milliunits per milligram of protein with 1 mU GRX activity defined as 1 nmol NADPH oxidized per minute. It should be noted that the TRX and GRX assays used in the present study measured total reducing activity and did not differentiate between the active (reduced) and inactive (oxidized) forms of these redox-active enzymes. Moreover, the assays did not distinguish between the different isoforms of TRX and GRX (15, 21, 25, 37).

TRXR activity was measured by a modification of the insulin disulfide reduction assay where excess TRX is added to a reaction mixture together with sample (11). Aliquots of cell lysate from ventricular tissue samples were added to a reaction mixture that contained 80 mM HEPES (pH 7.5), 6 mM EDTA, 0.9 mg/ml NADPH, and 2 mg/ml insulin. The reaction was started by adding 5 µM Escherichia coli TRX, and the samples were heated to 37°C for 20 min. The reaction was stopped by adding 500 µl of a solution that contained 0.4 mg/ml DTNB and 6 M guanidine hydrochloride in 0.2 M Tris·HCl, pH 8.0, and the absorbance was read at 412 nm. Measured absorbance values from tissue samples were compared with standard curves generated with known amounts of rat liver TRXR.

GR activity was measured by the technique of Carlberg and Mannervik (3). For this assay, tissue samples were homogenized in ice-cold Tris buffer (0.1 M, pH 8.0 with 2 mM EDTA) and centrifuged at 4°C (6,000 g) for 30 min. A 200-µl aliquot of the supernatant was added to a cuvette that contained KH2PO4 buffer (0.2 M, pH 7.0) plus 2 mM EDTA, 20 mM oxidized glutathione, and 2 mM NADPH. The change in absorbance at 340 nm was monitored for 5 min at 30°C. GR activity was expressed in milliunits, which were defined as the amount of enzyme catalyzing the reduction of 1 nmol NADPH per minute.

Glucose-6-phosphate dehydrogenase (G6PD) activity was measured using a commercial kit (OxisResearch). Ventricular tissue samples were homogenized in supplied diluent and centrifuged at 4°C (6,000 g) for 30 min, and a 50-µl aliquot of supernatant was added to a reaction mixture that contained NADP, glucose-6-phosphate, and 6-phosphogluconic acid. The change in absorbance at 340 nm in the presence of both substrates was measured at 37°C for 5 min. A second 50-µl aliquot of supernatant was added to a separate reaction mixture that contained NADP plus 6-phosphogluconic acid alone, and absorbance was again measured at 340 nm. G6PD activity was calculated by subtracting the rate of change of absorbance with 6-phosphogluconic acid alone from that measured with the combined substrates. The rationale for this procedure is to eliminate the contribution of 6-phosphogluconate dehydrogenase to total NADPH production and thus yield a more accurate estimation of rate-limiting G6PD activity. The net change in absorbance was expressed in milliunits, which were defined as the enzyme activity to produce 1 nM of NADPH per minute. Activities for TRX, GRX, TRXR, and G6PD were normalized per milligram of protein as measured using a commercial kit (Pierce).

Statistical analysis. All results are expressed as means ± SE. Statistical comparisons of two groups were made using Student's t-test, whereas comparison of more than two groups was carried out by ANOVA. When a significant difference among groups was indicated by the initial analysis, individual paired comparisons were made using Dunnett's t-test. Differences were considered significant at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Diabetes-induced changes in oxidoreductase systems. Cellular protection from irreversible oxidative damage in most mammalian cell types involves the activities of thiol-disulfide oxidoreductase systems that control the redox state of proteins and participate in antioxidant defense mechanisms (10, 15, 24, 37). These endogenous pathways undergo dynamic changes in response to environmental stress conditions such as oxidative stress (10, 19, 22, 33, 37). To examine the status of cardiac oxidoreductase systems after 3–5 wk of diabetes, tissue samples from the left ventricle were assayed for TRX, GRX, TRXR, and GR. Figure 1 (A and C) shows mean data of the basal activities of TRX and GRX, which are the terminal components of the major oxidoreductase systems. Compared with values from control animals, the activities of TRX (Fig. 1A) and GRX (Fig. 1C) were significantly increased in diabetic rat hearts. In particular, TRX activity was increased 1.7-fold, whereas GRX was increased 2.5-fold from control heart (P < 0.05). A second series of assays was conducted to compare the activities of TRXR and GR in diabetic and control rat hearts. Figure 1(B and D) illustrates that in contrast to the respective downstream effectors TRX and GRX, the activities of TRXR (Fig. 1B) and GR (Fig. 1D) in the left ventricle were significantly decreased in diabetic rat hearts compared with control hearts. Specifically, the basal activity of TRXR in diabetic rat heart was 52% less than in control heart (P < 0.05), which paralleled a change in GR (Fig. 1D) that was decreased by 35% from control heart (P < 0.05). In the present study, we did not analyze regional differences in TRXR activity in control or diabetic rat hearts. Recent data from our laboratory showed no significant regional differences in GR activity in diabetic or control rat hearts (S. Li, X. Li, and G. J. Rozanski, unpublished observations).



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  		Fig. 1. Oxidoreductase systems in diabetic rat heart. Tissue samples from left ventricle of control and diabetic rat hearts were assayed for thioredoxin (TRX, A), thioredoxin reductase (TRXR, B), glutaredoxin (GRX, C), and glutathione reductase (GR, D) activity. Numbers in parentheses represent n, no. of hearts assayed. *P < 0.05 compared with control.

 
The overall activities of the TRX and GRX systems rely on the supply of reducing equivalents in the form of NADPH, which is produced mainly by the pentose pathway in most cell types. The rate-limiting enzyme of this pathway, G6PD, converts NADP+ to NADPH during the conversion of glucose-6-phosphate to 6-phosphogluconate (12, 13). Because this key enzymatic step has been shown to significantly affect cell redox state in heart (12, 13), we conducted additional assays to determine whether the pentose pathway could have contributed to electrical remodeling in our diabetic rats. However, as summarized in Fig. 2, left ventricular G6PD activity values at 3–5 wk of experimental diabetes were not different from control values.



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  		Fig. 2. Basal activity of glucose-6-phosphate dehydrogenase (G6PD). Left ventricular tissue samples from control and diabetic rat hearts were assayed for G6PD. Numbers in parentheses represent n, no. of hearts assayed.

 
Reversal of K+ channel remodeling and role of TRX system. Voltage-clamp studies from our laboratory (29, 34–36) and others (5, 17, 26, 32) have consistently shown significant decreases in the basal density of specific K+ currents in myocytes from diabetic rat hearts compared with control animals. Consistent with these findings, the present study conducted at 3–5 wk of diabetes found that the maximum Ito density (+60 mV) in diabetic rat myocytes under basal conditions was ~35% less than in control myocytes (P < 0.05; Fig. 3) with no significant changes in voltage- or time-dependent properties. Also consistent with previous studies, Ito was sensitive to metabolic stimuli augmenting glucose utilization in cardiac myocytes. In particular, treatment of diabetic rat myocytes with insulin or the pyruvate dehydrogenase activator dichloroacetate (DCA) normalized Ito density after a delay of several hours. Figure 3A compares raw current traces from a control rat myocyte and diabetic rat myocytes treated with 0.1 µM insulin or 1.5 mM DCA for 3–4 h. Ito density was clearly increased in the insulin- and DCA-treated myocytes compared with the untreated cell from the same group. The mean current-voltage relations of several myocytes from each group are compared in Fig. 3B and show that insulin and DCA significantly increased Ito density in diabetic rat myocytes to control levels. In the DCA-treated group, Ito density was slightly greater than the control level at more negative test potentials (–20 to 10 mV). Similar treatment of myocytes from control rats with 0.1 µM insulin or 1.5 mM DCA for 3–4 h did not alter maximum Ito density (data not shown). As in previous studies (34–36), the IK1 was not altered by diabetic conditions.



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  		Fig. 3. Upregulation of transient outward K+ current (Ito) by insulin and dichloroacetate (DCA) in diabetic rat myocytes. Superimposed current traces elicited by voltage-clamp depolarizations from –40 to +60 mV (20-mV increments) are shown for control and diabetic rat myocytes pretreated with 0.1 µM insulin or 1.5 mM DCA for 3–4 h (A). Mean current-voltage relations for Ito in control and indicated experimental groups are shown (B). *P < 0.05 compared with control.

 
To examine the functional impact of G6PD on redox control of K+ channels, the responses of Ito to insulin and DCA were retested in diabetic rat myocytes in the presence of one of two structurally distinct G6PD inhibitors, dehydroepiandrosterone (DHEA) or 6-aminonicotinamide (6-AN). In these and subsequent experiments to test the effects of different blockers, myocytes were pretreated with inhibitor for 30 min before 0.1 µM insulin or 1.5 mM DCA was added for an additional 3–4 h. Figure 4 illustrates that the effects of insulin and DCA to upregulate Ito were inhibited by 100 µM DHEA and 1 mM 6-AN. Neither blocker alone significantly altered maximum Ito density when examined in diabetic rat myocytes treated for 3–4 h (6-AN treated: 16.4 ± 2.5, n = 4; DHEA treated: 14.3 ± 4.2, n = 4; untreated: 16.5 ± 1.9 pA/pF, n = 14; P > 0.05) nor control myocytes treated for the same duration (6-AN treated: 27.4 ± 3.1, n = 10; DHEA treated: 32.1 ± 4.3, n = 7; untreated: 28.4 ± 1.0 pA/pF, n = 30; P > 0.05). Although these data suggest that G6PD plays a prominent role in the regulation of K+ channels, we found that the activity of this enzyme in our model of diabetes was not different from control (see Fig. 2).



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  		Fig. 4. Effects of G6PD inhibitors on response of Ito to insulin and DCA. Myocytes from diabetic rat hearts were treated with 0.1 µM insulin (A) or 1.5 mM DCA (B) for 3–4 h with or without the G6PD inhibitors dehydroepiandrosterone (DHEA, 100 µM) or 6-aminonicotinamide (6-AN, 1 mM). Maximum Ito density (+60 mV) for each cell is expressed relative to the mean value of control myocytes (28.4 pA/pF; n = 30). Numbers in parentheses represent n, no. of myocytes examined from 2 or 3 diabetic rats. Data for untreated myocytes in B are replotted from A. *P < 0.05 compared with untreated diabetic rat myocytes.

 
We previously reported that the electrophysiological effect of insulin on diabetic rat myocytes is blocked by inhibitors of glutathione metabolism, particularly 1,3-bis-(2-chloroethyl)-1-nitrosourea (BCNU), which is a widely used inhibitor of GR (34). In the present study, we also found that 100 µM BCNU blocked upregulation of Ito density by insulin in diabetic rat myocytes as it did for DCA-treated cells (data not shown). However, studies on other cell types suggest that BCNU also inhibits TRXR, which shares a high degree of homology with GR (30). Thus to further probe the role of the TRX system in controlling K+ channel remodeling, we examined the effects of two structurally different compounds with relatively high selectivity profiles for TRXR: 13-cis-retinoic acid (RA, 1 µM) and auranofin (AF, 10 nM). The latter is one of several gold-containing compounds that display high selectivity for proteins that contain selenocysteine such as TRXR (25). Figure 5(A and B) illustrates that both compounds blocked the electrophysiological effects of insulin and DCA on Ito in diabetic rat myocytes. Neither blocker alone altered Ito density in diabetic rat myocytes when treated for 3–4 h (RA treated: 16.9 ± 1.8, n = 5; AF treated: 15.7 ± 2.1, n = 4; untreated: 16.5 ± 1.9 pA/pF, n = 14; P > 0.05) nor in control myocytes treated for the same duration (RA treated: 27.9 ± 4.0, n = 11; AF treated: 31.0 ± 4.7, n = 10; untreated: 28.4 ± 1.0 pA/pF, n = 30; P > 0.05). The selectivity of RA and AF was also examined in isolated myocytes treated with inhibitor for 3–4 h and assayed for TRXR and GR. These data, which are summarized in Fig. 5C, indicate that RA and AF at the concentrations used in the present experiments significantly decreased TRXR activity with little change in GR.



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  		Fig. 5. Effects of TRXR inhibitors on response of Ito to insulin and DCA. Myocytes from diabetic rat hearts were treated with 0.1 µM insulin (A) or 1.5 mM DCA (B) for 3–4 h with or without 13-cis-retinoic acid (RA, 1 µM) or auranofin (AF, 10 nM) to inhibit TRXR. Maximum Ito density (+60 mV) for each cell is expressed relative to the mean value of control myocytes. Numbers in parentheses represent n, no. of myocytes examined from 2 or 3 diabetic rats. Data for untreated myocytes are replotted from Fig. 4. *P < 0.05 compared with untreated diabetic rat myocytes. Inhibitory effects of AF and RA are shown (C). TRXR and GR activities were assayed in suspensions of isolated myocytes from control rat hearts treated for 3–4 h with 10 nM AF or 1 µM RA. Data are expressed relative to untreated control samples. Numbers in parentheses represent n, no. of hearts examined. *P < 0.05 compared with the hypothetical value for no inhibition, 1.0.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Redox regulation of protein function by oxidoreductase systems. The etiology of diabetic cardiomyopathy is postulated to involve oxidative stress resulting from increased production of reactive oxygen species and deficits in antioxidant defense mechanisms (6, 18). A major consequence of excess reactive oxygen species production is oxidative modification of proteins that leads to alterations in cell function. Reversible protein oxidation mainly involves the reactivity of the free sulfhydryl (-SH) group of cysteine residues. This side chain can exist in a number of different molecular states that can trigger the activation or inactivation of protein activity (24). Accumulating evidence indicates that the redox status of cysteine sulfhydryl groups controls the structure and activity of many enzymes, receptors, transcription factors, and transport proteins required for normal cell function (2, 14, 21, 24, 25, 27, 37). Studies on heart and other tissues demonstrate that diabetic conditions are associated with elevated levels of oxidized cellular proteins (2, 6) including increased disulfide bond formation on the cardiac ryanodine receptor that may contribute to systolic dysfunction of the ventricle (2).

Protection of cellular proteins from oxidative modification mainly involves ubiquitous thiol-disulfide oxidoreductase systems. TRX and GRX belong to the thioredoxin superfamily of oxidoreductases that function within the cytoplasm in antioxidant reactions and in maintaining proteins in a reduced state. A functionally related set of enzymes is also localized to the mitochondria (15, 21, 25, 37). TRX and GRX are ~12-kDa proteins that contain a signature -Cys-X-X-Cys- motif at their active sites. For the cytosolic isoform of TRX, the active site contains a -Cys-Gly-Pro-Cys- motif, whereas the active site for GRX has a -Cys-Pro-Tyr-Cys- motif (10, 21). In addition to differences in their active sites, the two enzymes display unique substrate specificities. TRX operates with NADPH and TRXR (TRX system) to catalyze the reduction of intra- or intermolecular protein disulfides (4, 11, 15, 21, 25, 33, 37). In contrast, GRX catalyzes the reduction of protein-mixed disulfides as part of the GRX system that includes GSH and GR (4, 10, 14, 15, 21, 27). Together, these oxidoreductase systems act as sulfhydryl repair mechanisms to control the physiological function of proteins susceptible to oxidation.

Most if not all of the functions of TRX and GRX depend on the activities of TRXR and GR, respectively. Both reductases are ~110–120-kDa homodimers, and TRXR contains a unique COOH-terminal selenocysteine residue that is essential for enzyme activity (3, 11, 15, 21, 25, 37). The selenocysteine residue of TRXR makes this enzyme as well as other selenoenzymes sensitive to inhibition by gold-containing compounds such as aurothioglucose and auranofin (25). In the present study, we tested the efficacy of auranofin and RA based on their reported high selectivity profiles for TRXR (25). Our data using these blockers (see Fig. 5) suggest that the TRX system plays a key role in redox regulation of K+ channels that underlie Ito. However, the GRX system is also an important redox regulator of cell function (10, 14, 15, 22, 27), and we found this system to be altered by diabetes in a qualitatively similar manner as the TRX system (see Fig. 1, B and D). Despite evidence obtained in the present study, the precise electrophysiological functions of the cardiac TRX and GRX systems are not well defined. In particular, we previously reported that upregulation of Ito density by insulin in diabetic rat myocytes is inhibited by buthionine sulfoximine, which is a blocker of GSH synthesis, and is mimicked by exogenous GSH (34). These previous findings and the results of the present study suggest that the cardiac TRX and GRX systems may act synergistically to regulate K+ channel activity.

The expression of TRX and GRX in many cell types is sensitive to environmental stress factors such as inflammation, ultraviolet radiation, or H2O2 exposure (15, 19, 22, 37). It is postulated that stress-induced upregulation of oxidoreductase activity is an important cellular compensatory mechanism to protect vital proteins from irreversible oxidative damage. This mechanism has not been well characterized in heart, but recent studies in rat and mouse models of myocarditis have shown TRX expression to be increased at early stages of the immune response (19, 33). In our study, done at 3–5 wk of diabetes, measured TRX and GRX activities in extracts from left ventricle were markedly greater than those of control animals (see Fig. 1, A and C), which is consistent with (oxidative) stress-induced upregulation. It should be noted, however, that TRX and GRX assays measure total activity and do not differentiate between the active (reduced) and inactive (oxidized) forms of the enzymes. It is therefore likely that decreased TRXR and GR activities in diabetic heart (see Fig. 1, B and D) result in correspondingly decreased levels of active TRX and GRX. This condition would imply that the functional activity of the TRX and GRX systems can be significantly impaired by decreases in one or more of its components.

The reducing activity of the TRX and GRX systems requires metabolically derived NADPH, which is generated mainly by the cytosolic pentose pathway (12, 13). The rate-limiting enzyme of this pathway, G6PD, has been shown to have a significant effect on cell redox state. For example, contractile function of ventricular myocytes is inhibited by pharmacological blockers of G6PD (12). Hearts from mice deficient in cardiac G6PD exhibit depleted levels of GSH compared with controls and more severe injury after ischemia and reperfusion (13). Recent data from our laboratory also show that GSH is significantly depleted in diabetic rat ventricular myocytes and that restoration of control GSH levels by insulin and DCA is blocked by DHEA and 6-AN (S. Li, X. Li, and G. J. Rozanski, unpublished observations). These and the present studies (see Fig. 4) identify G6PD as an important component of redox-related control mechanisms. However, our data also show that cardiac G6PD activity at 3–5 wk of diabetes was not different from control (see Fig. 2), which suggests that the pentose pathway was not itself a major contributor to redox alterations in our diabetic rat model. Considering that glucose utilization is markedly impaired in diabetic heart (8, 28), it is possible that depressed metabolic steps upstream of G6PD may affect the availability of substrate to generate NADPH for the TRX and GRX systems.

Insulin regulation of Ito. Decreased insulin signaling and impaired glucose utilization are postulated to play major roles in electrical remodeling of heart in diabetes mellitus. In support of this hypothesis, we (29, 36, 34; see Figs. 3–5) and others (32) have shown that insulin treatment of diabetic rat ventricular myocytes upregulates Ito density to normal levels after a delay of several hours. Moreover, specific abnormalities of glucose utilization such as depressed pyruvate dehydrogenase activity have also been linked to diabetic cardiomyopathy (7). In this regard, we have shown that agents that directly activate this regulatory enzyme such as DCA mimic the effects of insulin on Ito in diabetic rat myocytes (36; Figs. 3–5). Nevertheless, a variety of experimental models indicate that intrinsic insulin signaling is a major factor controlling cardiac K+ channel function. For example, voltage-clamp studies of a genetic mouse model of type 2 (insulin-resistant) diabetes show that Ito density in heart is significantly decreased compared with control animals (31). Perhaps the most direct investigations of the electrophysiological importance of insulin signaling in heart are studies of mice with cardiac-specific knockout of the insulin receptor. This model is characterized by normal glucose tolerance (1) and significant but gender-dependent remodeling of Ito (31). These latter studies suggest an important contribution of intrinsic cardiomyocyte insulin signaling to cardiac ion-channel function that is independent of diabetes-related systemic factors such as hyperglycemia and hyperlipidemia. Nevertheless, these systemic factors likely accelerate the development of cardiac ion-channel remodeling in diabetes by promoting oxidative stress (6, 18).

Previous data from our laboratory and the present study (see Figs. 4 and 5) further suggest that redox-mediated mechanisms participate in regulating K+ channels that underlie Ito (29, 34), and that these mechanisms are under the control of the insulin-signaling cascade. It may be postulated that insulin and DCA increase substrate flux through the pentose pathway and thereby increase the availability of NADPH required by the TRX and GRX systems to reduce oxidized proteins involved in controlling Ito density. However, the redox-sensitive proteins regulating Ito density are not well characterized, nor is it known whether they are involved in 1) signaling pathways that control channel activity or gene expression, 2) trafficking or turnover of channel protein, or 3) direct modulation of channel protein or associated {beta}-subunits. In the context of diabetic cardiomyopathy, the insulin-sensitive control of Ito density in ventricular myocytes is blocked by inhibitors of protein synthesis and of protein trafficking via the cytoskeleton (32). These data are consistent with decreased Ito density in nondiabetic models of heart failure (20, 23, 29) and support the hypothesis that Ito channels are downregulated in chronic disease states by alterations in transcription and surface expression of channel protein (20, 23, 26). Nevertheless, the role of redox mechanisms in the control of Ito channel density and the contribution of insulin signaling necessitate further study.

In conclusion, K+ channels that underlie Ito are regulated in a redox-sensitive manner by the TRX system, although contribution by the GRX system cannot be ruled out. The remodeling of Ito that occurs in diabetes is correlated with significant decreases in TRXR and GR activities. We postulate that the TRX and GRX systems comprise a functionally important repair mechanism in heart that protects ion channels and other vital cellular proteins from irreversible oxidative damage.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
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REFERENCES
 
This work was supported by Grant HL-66446 from the National Heart, Lung, and Blood Institute and by the American Diabetes Association.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. J. Rozanski, Dept. of Cellular and Integrative Physiology, Univ. of Nebraska College of Medicine, 985850 Nebraska Medical Center, Omaha, NE 68198-5850 (E-mail: grozansk{at}unmc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

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