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Reversal of hyperglycemic preconditioning by angiotensin II: role of calcium transport

¹Department of Pharmacology, University of South Alabama College of Medicine, Mobile, Alabama; and ²Department of Oral Biology/Maxcillofacial Pathology, Medical College of Georgia, Augusta, Georgia

Submitted 20 August 2004 ; accepted in final form 8 December 2004

	ABSTRACT

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Myocardial cell death is an important contributor to the development of diabetic cardiomyopathy. It has been proposed that diabetes-mediated upregulation of the renin-angiotensin system leads to oxidative stress, the trigger for cardiomyocyte death and contractile dysfunction. However, the adverse effect of ANG II on the diabetic heart may extend beyond the development of the cardiomyopathy. ANG II also alters specific modulators of ischemic injury, such as PKC and calcium transport. Therefore, the present study examined the effect of ANG II on hyperglycemic preconditioning, a glucose-mediated condition associated with the elevation of PKC activity and alterations in calcium transport that render the cell resistant to hypoxia. Exposure of the glucose-treated cell to ANG II during the prehypoxic period blocked glucose-mediated cardioprotection. The reversal of hyperglycemic preconditioning was associated with enhanced accumulation of Ca²⁺ during hypoxia, an effect prevented by inhibition of the Na⁺/ H⁺ exchanger and the T-type Ca²⁺ channel. The inhibitors of hypoxia-mediated Ca²⁺ accumulation also blocked the reversal of hyperglycemic preconditioning by ANG II. Thus ANG II and glucose treatment exert opposite actions on the Na⁺/ H⁺ exchanger and the T-type Ca²⁺ channel. Because those transporters are involved in hypoxia-mediated apoptosis, they are logical candidates for the beneficial effects of high glucose and the adverse effects of ANG II on the hypoxic cardiomyocyte.

sodium-hydrogen exchanger; T-type calcium channel; protein kinase C; hyperglycemia; hypoxia; apoptosis; calcium overload

Both oxidative stress and impaired calcium transport play key roles in cardiomyocyte loss and impaired excitation-contraction coupling of the diabetic heart. In 1996, Kaul et al. (21) reported that the antioxidant probucol improved contractile function in a rat model of streptozotocin-dependent diabetes. In a related study, Esberg and Ren (6) found that scavengers of nitric oxide, superoxide, and peroxynitrite prevented contractile defects of cardiomyocytes cultured for 24 h in medium containing 25 mM glucose. Because the antioxidants also improved Ca²⁺ homeostasis, oxidative stress appeared to affect the activity of Ca²⁺-dependent transporters, such as the Na⁺/Ca²⁺ exchanger and sarco(endo)plasmic reticulum Ca²⁺-ATPase (SERCA). However, Fiordaliso et al. (10) implicated the diabetes-mediated upregulation of ANG II in the production of reactive oxygen species, the loss of cardiomyocytes, and the development of cardiomyopathy. In support of their hypothesis, they found that components of the renin-angiotensin system were upregulated in the diabetic heart. Moreover, both the antioxidant Tiron and the AT₁ receptor antagonist losartan blocked diabetes-mediated apoptosis and hydroxyl radical production (10, 18).

Upregulation of the renin-angiotensin system may also account for the increased susceptibility of the diabetic heart to ischemia. It is widely recognized that ANG II stimulates the activity of the Na⁺/H⁺ exchanger, the Na⁺/Ca²⁺ exchanger, and the T-type Ca²⁺ channel (3, 8, 15), transporters implicated in ischemia-mediated Ca²⁺ accumulation. Moreover, ANG II promotes the translocation and activation of PKC-, an isoform thought by some investigators to adversely affect the ischemic heart (28). Furthermore, several investigators have shown that AT₁ receptor blockade and angiotensin-converting enzyme inhibition protect the ischemic heart (17). However, the effects of ANG II on the ischemic myocardium remain controversial. ANG II has also been found to promote the translocation and activation of PKC-, an isoform implicated in ischemic preconditioning (14, 23, 34). According to Ford et al. (11), the interaction of ANG II with the AT₁ receptor mediates cardioprotection. Moreover, transgenic mice overexpressing human renin and angiotensinogen transgenes show enhanced recovery of contractile function after a hypoxia-reoxygenation insult (37). In view of these controversies and the importance of ANG II in diabetes, further studies on the effect of ANG II on the diabetic cardiomyocyte seem warranted. The present study examined the effect of ANG II on the glucose-treated cardiomyocyte, a model that focuses on the effects of chronic hyperglycemia and mimics many of the changes seen in the diabetic cardiomyocyte (5, 30, 31). Isolated cells were preferred for the present study because interpretation of the ANG II effect in the isolated heart would be complicated by the vasoactivity of the peptide.

	MATERIALS AND METHODS

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Cardiomyocyte preparation and incubation conditions. Neonatal cardiomyocytes were prepared as described previously (35). The cells were suspended in minimal essential medium containing 10% newborn calf serum and 0.1 mM 5'-bromo 2'-deoxyuridine and allowed to plate on either glass coverslips or polystyrene-treated petri dishes (Cat. no. 430167, Corning) at a density of 10 x 10⁶ cells/dish (10-cm diameter). They were then placed in serum-free medium containing minimal essential medium supplemented with either no addition (control) or 25 mM glucose (glucose treated). In some experiments, either 1 nM ANG II or varying concentrations of the PKC inhibitor chelerythrine were added to the incubation medium after 48 h. All experiments were initiated after a 3-day incubation at 37°C under a 5% CO₂-20% O₂ environment. To induce chemical hypoxia, the cells were transferred to medium lacking glucose and ANG II but containing 10 mM deoxyglucose and 3 mM amobarbital. In some experiments, the T-type Ca²⁺ channel antagonist mibefradil (0.3 µM), the Na⁺/H⁺ exchange antagonist cariporide (10 µM), or the combination of mibefradil and cariporide were added to the hypoxic medium.

TdT-mediated dUTP nick-end labeling procedure. The Klenow Frag EL DNA fragmentation detection kit was used to monitor end-labeled DNA fragments of apoptotic nuclei. Cells were assayed for positive TdT-mediated dUTP nick-end labeling (TUNEL) staining before and after 1 h of chemical hypoxia. In each case, the cells were fixed for 15 min in 4% formaldehyde and then resuspended in 80% ethanol for 20 min. After rehydration in PBS, the samples were incubated for 5 min with 20 µg/ml proteinase K in 10 mM Tris (pH 8.0). The slides were then rinsed in PBS. Endogenous peroxidases were inactivated by a 5-min incubation with 3% H₂O₂. After being rinsed again, the slides were placed in 1x Klenow buffer for 20 min. The Klenow labeling reaction mixture was then added to the samples, and the samples were incubated for 1.5 h at 37°C. The reaction was terminated by a 5-min incubation with buffer containing 0.5 mM EDTA (pH 8.0) followed by a 10-min exposure to blocking buffer. The samples were then treated for 30 min with peroxidase-streptavidin conjugate. After a 15-min incubation with 0.7 mg 3,3-diaminobenzidien and 0.6 mg H₂O₂-urea, the samples were rinsed and then counterstained with hematoxylin. To fix the color, the glass slides were first immersed in 100% ethanol and then xylene. Four separate fields in the light microscope were examined for dark brown (apoptotic) and purple (normal) nuclei.

Western blot analyses. The cytosolic and particulate content of various PKC isoforms were determined by Western blot analysis. After the cells were scraped from the bottom of the petri dishes, they were suspended in ice-cold Tris buffer (pH 7.4) containing 5 mM EGTA, 2 mM EDTA, 100 mM NaF, 5 mM dithiothreitol, a 1/100 dilution of protease inhibitor cocktail (Calbiochem), and 1% solutions of leupeptin and PMSF. The samples were then centrifuged at 100,000 g for 60 min. The pellet constituted the membrane-particulate fraction, and the particulate-free supernatant fraction was defined as the cytosolic fraction. Each sample containing the particulate fraction was resuspended in homogenizing buffer containing 0.5% Triton X-100 and centrifuged at 100,000 g for 60 min. The resulting detergent-treated supernatant was used in the Western blot analysis. The protein concentration of each sample was determined by the Bradford assay. Cytosolic and membrane proteins were analyzed for PKC isoform content by one-dimensional electrophoresis using 8% SDS-polyacrylamide gels. The samples were then transferred to nitrocellulose membranes, where they were blocked. After incubation with the appropriate antibody, the membranes were washed and then incubated with a secondary antibody, goat anti-rabbit IgG. The Western blots were detected by the enhanced chemiluminescence reaction. All data were analyzed by a computer program, which detected the area under the curve. Gels were also analyzed by Ponceau's solution to ensure equal protein loading.

Cellular Ca²⁺ content. Cardiomyocytes were loaded for 20 min with fura-2 using loading buffer (2 ml Krebs buffer with 25 mM HEPES plus 3 mM fura-2 AM and 6 µl Pluronic acid, 37°C). The loading and subsequent rinsing steps took place before the onset of the hypoxic insult. Fura-2 fluorescence at 340 and 380 nm (emission at 510 nm) was measured using an Olympus IX70 inverted microscope at x400 with a xenon arc lamp photomultiplier system (Photon Technologies; Monmouth Junction, NJ). Data obtained during the course of the hypoxic insult were expressed as the fluorescence ratio (F_340/380), a measure of intracellular Ca²⁺ concentration ([Ca²⁺]_i). To evaluate the basis for alterations in hypoxia-mediated Ca²⁺ accumulation, some data were expressed as the difference in fura-2 fluorescence between two experimental conditions (F_340/380 or F'_340/380), as described further in RESULTS.

Current-voltage relationship. Conventional whole cell voltage-clamp configurations were used as described previously (39). Briefly, to measure transmembrane currents in single rat cardiomyocytes by the standard gigaseal patch-clamp technique, whole cell recordings were performed. Recording pipettes were made of hemo capillaries (Warner Instruments; Hamden, CT) pulled by a two-stage puller (PC-10, Narishige; Tokyo, Japan) and heat polished with a microforge (model MF-200, World Precision Instruments; Sarasota, FL) before use. Pipette resistance was in the range of 2–5 M when filled with intracellular solution. All experiments were performed at room temperature (22–25°C). An EPC-9 patch-clamp amplifier (HEKA Elektronik, Lambrecht; Pfalz, Germany) was used to acquire data with Pulse/PulseFit software (HEKA) and filtered at 2.9 kHz. Voltage-dependent currents were corrected for linear leak and residual capacitance using an on-line P/n subtraction paradigm; the definition of the so-called P/n leak correction protocol is as follows: in a voltage range, where voltage-dependent channels are not active, a scaled-down version of the pulse protocol is applied n times, and the resulting current is averaged, scaled, and subtracted from that elicited by the main test pulse. The extracellular (bath) solution contained (in mmol/l) 2 CaCl₂, 110 tetraethylammonium chloride, 10 CsCl, and 10 HEPES 10 (pH 7.4, adjusted with tetraethylammonium hydroxide). The intracellular (pipette) solution contained (in mmol/l) 130 N-methyl-D-glucamine, 10 EGTA, 5 BAPTA, 10 HEPES, 6 MgCl₂, 4 CaCl₂, and 2 Mg-ATP (pH 7.2, adjusted with methane sulfonic acid). All solutions were adjusted to 290 to 300 mosM with sucrose.

Statistical analysis. The statistical significance of the data was determined using the Student's t-test for comparison within groups and ANOVA combined with Tukey's post hoc test for comparison between groups. Values of P < 0.05 were considered statistically significant.

	RESULTS

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Figure 1A shows that chemical hypoxia triggers a time-dependent increase in apoptosis among isolated cardiomyocytes in culture. Before hypoxia, 11.9 ± 1.3% of control and 14.2 ± 1.6% of glucose-treated cardiomyocytes were apoptotic, where control and glucose treatment refer to cells incubated for 3 days with medium containing 5 and 25 mM glucose, respectively. Two hours of chemical hypoxia carried out in the absence of glucose increased the percentage of apoptotic cells in the control and glucose-treated groups to 54.5 ± 5.1% and 33.6 ± 5.5%, respectively. Taking into consideration the number of apoptotic cells found in culture before chemical hypoxia, the 2-h hypoxic insult caused 42.6% of the control cells and 19.4% of the glucose-treated cells to undergo apoptosis. The difference between the two values (23.2%) represents the degree of hyperglycemic preconditioning, a term used to describe protection arising from prehypoxic (not hypoxic) exposure of cardiomyocytes to medium containing 25 mM glucose. As seen in Fig. 1B, the maximum degree of glucose protection (23.2 ± 1.1%) was achieved after 2 h of chemical hypoxia. However, the addition of ANG II (1.0 nM) to the prehypoxic incubation medium completely blocked the cardioprotective effect of high glucose; note that ANG II was also excluded from the hypoxic medium. This adverse effect of ANG II appears to be largely unrelated to its proapoptotic activity, because ANG II-mediated apoptosis disappears after prolonged periods of hypoxia. While the addition of ANG II to the prehypoxic incubation medium increased the number of apoptotic cells in culture by 8.5% before hypoxia, after 120 min of hypoxia, the extent of hypoxia-induced death was virtually identical in control cells incubated with medium containing ANG II (54.8 ± 6.3%) or those that had been exposed to ANG II during the prehypoxic period (54.5 ± 5.1%).

View larger version (20K): [in this window] [in a new window]   Fig. 1. Effect of ANG II on hyperglycemic preconditioning. Isolated neonatal cardiomyocytes were incubated for 3 days with medium containing either 5 (control) or 25 mM glucose (glucose). Some of the cells were treated during the last 24 h with medium containing 1 nM ANG II. The cells were then exposed for 45–150 min to medium lacking glucose and ANG II but containing 10 mM deoxyglucose and 3 mM amobarbital. The presence of nuclear end-labeled DNA fragments in these cells after various periods of chemical hypoxia was detected using the TdT-mediated dUTP nick-end labeling (TUNEL) technique. Data are expressed as the percentage of cells undergoing positive brown TUNEL staining, with 15–20 cells examined per field and 4–5 fields examined per preparation. A: time course of hypoxia-mediated increase in TUNEL staining of glucose-treated and control cells. Values shown represent means ± SE of 4–5 different cell preparations. *Significant difference between control and glucose-treated cells (P < 0.05). B: effect of ANG II on hyperglycemic preconditioning. TUNEL data are expressed as the percentage of glucose-treated cells protected against hypoxia-mediated apoptosis. Exposure of the glucose-treated cells to medium containing ANG II abolished hyperglycemic preconditioning. Values shown represent means ± SE of 4–5 preparations. *Significant difference between the ANG II-treated and untreated cells (P < 0.05).

View larger version (26K): [in this window] [in a new window]   Fig. 2. Effect of glucose treatment and ANG II exposure on the distribution of various PKC isoforms. Neonatal rat cardiomyocytes were incubated for 3 days with medium containing 5 (control) or 25 mM glucose. Some of the cells were exposed to medium containing 1 nM ANG II during the last 24 h. On the third day, cells were harvested and separated into cytosolic and membrane fractions by centrifugation. PKC isoform contents of the two fractions were measured by Western blot analysis, with bands detected by the enhanced chemiluminescence reaction after exposure to goat anti-rabbit IgG. A: representative Western blots of PKC isoforms (PKC-, PKC-2, PKC-, PKC-, and PKC-) from cytosolic and membrane fractions. B: relative levels of PKC-, PKC-2, PKC-, PKC-, and PKC- in control and glucose-treated cells. Values of 1 were assigned to bands corresponding to control PKC-, control PKC-2, control PKC-, control PKC-, and control PKC-. Data shown are means ± SE of 5–6 different preparations. *Significant differences between control and glucose-treated cells (P < 0.05). C: relative levels of PKC-, PKC-2, PKC-, PKC-, and PKC- in control and glucose-treated cells exposed to ANG II. Values of 1 were assigned to bands corresponding to control PKC-, control PKC-2, control PKC-, control PKC-, and control PKC-. Data shown are means ± SE of 5–6 different preparations. *Significant differences between control and glucose-treated cells exposed to ANG II (P < 0.05).

The translocation of PKC- is considered a key step in ischemic preconditioning and other types of cardioprotection (4, 14, 23, 29, 34). However, its role in hyperglycemic preconditioning remains unclear. One concern is that despite the elevation in particulate levels of PKC-, glucose treatment did not promote the translocation of the isoform. Moreover, ANG II caused some translocation of PKC- while reversing hyperglycemic preconditioning. Therefore, the potential role of cardioprotective PKC isoforms in glucose-mediated cardioprotection was further examined by evaluating the effectiveness of the PKC inhibitor chelerythrine in blocking hyperglycemic preconditioning. As expected, chelerythrine triggered a concentration-dependent increase in apoptosis of normoxic cells, with the degree of chelerythrine-mediated apoptosis being identical in control and glucose-treated cardiomyocytes during normoxia but not hypoxia. Figure 3 shows the effect of chelerythrine on hypoxia-mediated apoptosis in control and glucose-treated cells after hypoxic data were corrected for the effect of chelerythrine during normoxia. A similar pattern was observed using another PKC inhibitor, calphostin C. In the absence of chelerythrine, 90 min of hypoxia caused 29% of the control cells and 11% of the glucose-treated cells to undergo apoptosis, with the difference between the two groups representing the degree of hyperglycemic preconditioning. Chelerythrine had no effect on hypoxia-mediated apoptosis in the control cells. However, it caused a concentration-dependent decrease in the degree of hyperglycemic preconditioning. The K_i for chelerythrine-mediated inhibition of hyperglycemic preconditioning was 1.0 µM, whereas the K_i for PKC inhibition is reportedly 0.7–0.8 µM (16), suggesting a role for PKC in hyperglycemic preconditioning.

View larger version (15K): [in this window] [in a new window]   Fig. 3. Effect of chelerythrine on hyperglycemic preconditioning. Neonatal rat cardiomyocytes were incubated for 3 days with medium containing either 5 (control) or 25 mM glucose. After 48 h, cells from each group were exposed to 0, 0.2, 1.0, 3.5, or 10 µM chelerythrine for 24 h. Cells were then subjected to either 1 h of normoxia or chemical hypoxia, the latter with medium containing 10 mM deoxyglucose and 3 mM amobarbital but no glucose or chelerythrine. The extent of apoptosis was assessed from the number of cells exhibiting positive TUNEL staining. The data were corrected for the effects of chelerythrine during normoxia. Values shown represent means ± SE of 4 preparations.

View larger version (18K): [in this window] [in a new window]   Fig. 4. Effect of glucose treatment and ANG II exposure on hypoxia-induced rise in intracellular Ca2+ concentration ([Ca2+]i). Neonatal rat cardiomyocytes were incubated for 3 days with medium containing 5 (control) or 25 mM glucose before being loaded with fura-2. Some of the cells were exposed to medium containing 1 nM ANG II during the last 24 h. The cells were then loaded with fura-2, and fluorescence at 340 and 380 nm was measured. After the fura-2 fluorescence was obtained for the normoxic cells, the cells were exposed to 1 h of chemical hypoxia. Shown is the time course of the hypoxia-induced rise in [Ca2+]i of representative cells. F340/380, fluorescence ratio.

View larger version (20K): [in this window] [in a new window]   Fig. 5. Effect of PKC and Na+/Ca2+ exchanger inhibitors on hypoxia-mediated Ca2+ accumulation. Fura-2-loaded cardiomyocytes were subjected to a hypoxic insult in medium supplemented with either no addition (control), PKC inhibitors [chelerythrine (1.0 µM) or calphostin C (1.0 µM)], or the Na+/Ca2+ exchange inhibitor KB-R7943 (0.3 µM). The effect of each inhibitor on hypoxia-mediated Ca2+ accumulation, represented by F340/380, of control cells was assessed after the initiation of chemical hypoxia. Shown are tracings from representative cells.

View larger version (17K): [in this window] [in a new window]   Fig. 6. Effect of hyperglycemic preconditioning on hypoxia-mediated Ca2+ influx via Na+/H+ and Na+/Ca2+ exchangers. Cardiomyocytes were incubated for 3 days with medium containing either 5 or 25 mM glucose. The fura-2-loaded cells were then subjected to a hypoxic insult in medium supplemented with either no addition (control) or the Na+/H+ exchange inhibitor cariporide (10 µM). The effect of cariporide on hypoxia-mediated Ca2+ accumulation, represented by F340/380, of control and glucose-treated cells was assessed after 1 h of chemical hypoxia. A: effect of cariporide on hypoxia-mediated Ca2+ accumulation (F340/380) by control cells. B: effect of cariporide on hypoxia-mediated Ca2+ accumulation (F340/380) by glucose-treated cells. C: effect of cariporide on reduction of hypoxia-mediated Ca2+ accumulation by glucose treatment. The change in hypoxia-mediated Ca2+ accumulation (defined as F340/380) was assessed by subtracting the F340/380 value of the glucose-treated cells after 1 h of chemical hypoxia from the F340/380 value of the control cells after 1 h of hypoxia. Cariporide reduced F340/380, indicating that flux through the Na+/H+ and Na+/Ca2+ exchangers is reduced by glucose treatment. Values shown represent means ± SE of 3–14 different cellular preparations, with 3 cells examined for each preparation. *Significant difference between the cariporide-treated and untreated cells (P < 0.05).

View larger version (17K): [in this window] [in a new window]   Fig. 7. Contribution of T-type Ca2+ current to hypoxia-mediated Ca2+ accumulation. Neonatal rat cardiomyocytes were loaded with fura-2, and fluorescence was monitored at 340 and 380 nm. After a steady-state fura-2 fluorescence was obtained for cells incubated under normoxic conditions, the cells were subjected to chemical hypoxia with medium containing no additions (control), cariporide (10 µM), or the combination of cariporide and the T-type Ca2+ channel antagonist mibefradil (0.3 µM). Shown is the time course of the hypoxia-induced rise in [Ca2+]i of representative cells.

View larger version (23K): [in this window] [in a new window]   Fig. 8. Effect of hyperglycemic preconditioning on hypoxia-mediated Ca2+ influx via the T-type Ca2+ channel. Glucose-treated and control cells were loaded with fura-2 and then subjected to chemical hypoxia in medium supplemented with either the Na+/H+ exchange inhibitor cariporide (10 µM) or the combination of cariporide and the T-type Ca2+ channel antagonist mibefradil (0.3 µM). The effect of mibefradil on Ca2+ accumulation of control and glucose-treated cells after 1 h of chemical hypoxia in medium containing cariporide was assessed. A: effect of mibefradil on hypoxia-mediated Ca2+ accumulation (F340/380) by control cells exposed to cariporide. B: effect of mibefradil on hypoxia-mediated Ca2+ accumulation (F340/380) by glucose-treated cells exposed to cariporide. C: contribution of the T-type Ca2+ channel toward reduction of hypoxia-mediated Ca2+ accumulation by glucose treatment. F340/380 was assessed by subtracting F340/380 of the glucose-treated cell after 1 h of chemical hypoxia from the F340/380 value of the control cell after 1 h of hypoxia. Mibefradil reduced F340/380 in cells exposed to cariporide, indicating that flux through the T-type Ca2+ channel is reduced by glucose treatment. Values shown represent means ± SE of 3–14 different cellular preparations, with 3 cells examined for each preparation. #Significant difference between the cariporide-treated and cariporide + mibefradil-treated cells (P < 0.05).

View larger version (18K): [in this window] [in a new window]   Fig. 9. Glucose treatment decreases T-type Ca2+ channel activity in cardiomyocytes. Cardiomyocytes were incubated for 3 days with medium containing either 5 (control) or 25 mM glucose (glucose). A: representative families of Ca2+ currents in cardiomyocytes. Voltage steps from a negative holding potential (VH = –90 mV) reveal a total inward Ca2+ current (top) that comprises both low- and high-voltage-activated (LVA and HVA) components (i.e., T- and L-type Ca2+ currents). The HVA component (L type) was taken (bottom) directly from the recordings, with VH = –40 mV and LVA (T type) channels being inactivated. The LVA component (T type) was obtained by taking the difference between the current measured from VH of –90 and –40 mV. B: current traces corresponding to the LVA [VH(–90) – VH(–40)] component at –30 mV and HVA [VH(–40)] at 0 mV from normal () and 25 mM glucose-treated () cells. C: current-voltage curves (of peak currents) representing the measurements made at a VH of –40 mV (top) and the difference between the measurements made at –90 and –40 mV (bottom) from control () and glucose-treated () cardiomyocytes (n = 15 and 12, respectively), showing that glucose treatment decreases whole cell LVA (T type) Ca2+ current but not HVA (L type) Ca2+ current. Ca2+ (2 mM) was used as a charge carrier. Membrane potential (Vm) is in mV, and total current is in pA. Temperature was 22–25°C.

View larger version (18K): [in this window] [in a new window]   Fig. 10. Effect of ANG II on hypoxia-mediated Ca2+ influx via Na+/H+ and Na+/Ca2+ exchangers and the T-type Ca2+ channel. Cardiomyocytes were incubated for 3 days, with the last 24 h with medium containing either no addition (control) or 1.0 nM ANG II. The cells were loaded with fura-2 and then subjected to a hypoxic insult in medium lacking ANG II but supplemented with no further additions (control), the Na+/H+ exchange inhibitor cariporide (10 µM), or the combination of cariporide and the T-type Ca2+ channel antagonist mibefradil (0.3 µM). The effect of cariporide and the combination of cariporide plus mibefradil on Ca2+ accumulation of control and ANG II-treated cells was assessed after 1 h of chemical hypoxia. A: effect of cariporide and cariporide plus mibefradil on hypoxia-mediated Ca2+ accumulation (F340/380) by control cells. B: effect of cariporide and cariporide plus mibefradil on hypoxia-mediated Ca2+ accumulation (F'340/380) by ANG II-pretreated cells. C: contribution of flux through the Na+/H+-Na+/Ca2+ exchanger combination and the T-type Ca2+ channel toward enhanced hypoxia-mediated Ca2+ accumulation by ANG II pretreatment. The elevation in hypoxia-mediated Ca2+ accumulation (defined as ’F340/380) was assessed by subtracting the F340/380 value of the control cell after 1 h of chemical hypoxia from the F'340/380 value of the ANG II-pretreated cell after 1 h of hypoxia. Cariporide reduced F'340/380, indicating that flux through the Na+/H+-Na+/Ca2+ exchanger combination was enhanced by ANG II. Similarly, mibefradil reduced F'340/380 in cells exposed to cariporide, indicating that enhanced T-type Ca2+ current contributes to elevated hypoxia-mediated Ca2+ accumulation in ANG II-treated cells. Values shown represent means ± SE of 3–14 different cellular preparations, with 3 cells examined for each preparation. *Significant difference between the cariporide-treated and untreated cells (P < 0.05); #significant difference between the cariporide-treated and cariporide + mibefradil-treated cells (P < 0.05).

View larger version (16K): [in this window] [in a new window]   Fig. 11. Effect of cariporide and mibefradil on the combined actions of ANG II and glucose treatment. Cardiomyocytes were incubated for 3 days with medium containing 25 mM glucose, with some of the cells exposed to 1 nM ANG II during the last 24 h. The glucose + ANG II-treated cells were loaded with fura-2 and then subjected to a hypoxic insult in medium lacking glucose and ANG II but supplemented with no further additions (glucose + ANG II), the Na+/H+ exchange inhibitor cariporide (10 µM), or the combination of cariporide and the T-type Ca2+ channel antagonist mibefradil (0.3 µM). Cariporide and cariporide plus mibefradil reduced hypoxia-mediated Ca2+ accumulation in glucose + ANG II-treated cells. The effects of ANG II and glucose treatment on Ca2+ homeostasis were largely additive. *Significant difference between the glucose and glucose + cariporide cells (P < 0.05).

View larger version (16K): [in this window] [in a new window]   Fig. 12. Effect of cariporide and mibefradil in the reversal of hyperglycemic preconditioning by ANG II. Cardiomyocytes were incubated for 3 days with medium containing either 5 or 25 mM glucose, with the last 24 h being with medium containing either no further addition (control) or 1.0 nM ANG II. The cells were then subjected to a hypoxic insult in medium lacking ANG II but supplemented with no further additions (control) or the combination of the Na+/H+ exchange inhibitor cariporide (10 µM) and the T-type Ca2+ channel antagonist mibefradil (0.3 µM). After 1 h of chemical hypoxia, cell death was assessed by TUNEL staining. Values shown represent means ± SE of 4 preparations, with 3–5 fields examined (100 cells examined/field). *Significant difference between cells treated with cariporide plus mibefradil and those lacking the inhibitors (P < 0.05).

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

The present study supports a role for both PKC and improved Ca²⁺ handling in hyperglycemic preconditioning. PKC-₂ is a potential candidate for the dependence of hyperglycemic preconditioning on PKC. Not only are the levels of PKC-₂ elevated in the diabetic heart (13, 24), but PKC-₂ has been implicated in the development of the diabetic cardiomyopathy (38). Recent studies suggest that PKC-₂ enhances the expression of both the angiotensin-converting enzyme (40) and the L-type calcium channel (1). Therefore, PKC-₂ overexpression mediates two events that adversely impact hyperglycemic preconditioning: the activation of the renin-angiotensin system and enhanced calcium influx. In light of these actions, it is difficult to attribute hyperglycemic preconditioning to the upregulation of PKC-₂.

There is substantial evidence that the translocation of one of the PKC isoforms, PKC-, from the cytosol to the particulate fraction contributes to ischemic preconditioning and other forms of cardioprotection (4, 14, 23, 29, 34). Not only does PKC- translocate to the membrane in response to ischemic preconditioning (29), but downregulation of PKC- through gene disruption or use of a selective PKC- antagonist blocks ischemic/hypoxic preconditioning (14, 34). Moreover, overexpression of PKC- protects the ischemic heart while leaving H⁺ generation unchanged (4). However, a role for PKC- in hyperglycemic preconditioning is less clear. Levels of PKC- in the particulate fraction of the glucose-treated cardiomyocyte were elevated relative to those of the control cell, but the 24% increase in particulate PKC- content occurred without the translocation of the isoform from the cytosol to the membrane fraction. Moreover, ANG II modestly promoted the translocation of PKC- from the cytosol to the particulate fraction in the glucose-treated cell despite reversing hyperglycemic preconditioning.

Cell signaling initiated by ANG II is dependent on the activation of PKC (32). Therefore, the substantial translocation of PKC- from the cytosol to the particulate fraction is likely an important event. Indeed, both glucose treatment and ANG II increase particulate levels of PKC-; however, the increase in particulate PKC- by ANG II is associated with a decrease in cytosolic levels of the isoform, whereas glucose treatment elevates both the particulate and cytosolic content of PKC-. Thus ANG II promotes the translocation of PKC- from the cytosol to the particulate fraction, whereas glucose treatment primarily enhances the expression of PKC-, causing only a modest shift in the localization of PKC- to the particulate fraction. The significance of elevated particulate PKC- content on the hypoxic cardiomyocyte remains controversial. According to Zhao and Kukreja (41) and Miyawaki et al. (26), PKC- exerts cardioprotective activity, whereas Murriel and Mochly-Rosen (28) feel that PKC- promotes cell damage. Nonetheless, the study by Murriel and Mochly-Rosen (28) using state-of-the-art techniques provides compelling evidence linking PKC- to cardiac injury. Assuming PKC- worsens ischemic injury, as reported by Murriel and Mochly-Rosen (28), it is possible that glucose-mediated elevations in PKC- levels act to minimize the extent of hyperglycemic preconditioning. Moreover, it would be attractive to suggest that a downstream effector of PKC- might be involved in the reversal of hyperglycemic preconditioning by ANG II.

One of the unique and extremely important cardioprotective mechanisms in the diabetic heart is the improvement in Ca²⁺ homeostasis. Tani and Neely (36) made the initial discovery that diabetic hearts were resistant to ischemia-mediated Ca²⁺ accumulation. This improvement was subsequently traced in part to a diabetes-linked reduction in Na⁺/H⁺ exchanger activity (22). In a related study, Schaffer et al. (35) found that the glucose-treated cardiomyocyte was also resistant to hypoxia-induced injury, an effect associated with an improvement in Na⁺ and Ca²⁺ handling. The present study provides additional lines of evidence supporting the notion that hyperglycemic preconditioning depends on transporters involved in hypoxia-mediated Ca²⁺ accumulation. First, glucose treatment renders the cell resistant to hypoxia-mediated cell death while lowering [Ca²⁺]_i. Second, glucose treatment reduces the activity of two transporters, the Na⁺/H⁺ exchanger and the T-type Ca²⁺ channel, which contribute to hypoxia-mediated Ca²⁺ accumulation. Third, inhibitors of the Na⁺/H⁺ exchanger and the T-type Ca²⁺ channel protect the control cell against hypoxia-mediated Ca²⁺ accumulation and cell death. Fourth, ANG II reverses hyperglycemic preconditioning while elevating [Ca²⁺]_i. Finally, inhibitors of the Na⁺/H⁺ exchanger and the T-type Ca²⁺ channel prevent reversal of hyperglycemic preconditioning by ANG II.

Although the combined actions of the Na⁺/H⁺ and Na⁺/Ca²⁺ exchangers have been touted as a major source of hypoxia-mediated Ca²⁺ accumulation (20), a role for the T-type Ca²⁺ channel in Ca²⁺ overload has received little attention. Yet, in the present study, the T-type Ca²⁺ channel antagonist mibefradil was found to be a potent inhibitor of hypoxia-mediated Ca²⁺ accumulation. Some of mibefradil's effect have been attributed to its weak inhibition of the Na⁺/Ca²⁺ exchanger; nonetheless, we found that the T-type Ca²⁺ channel antagonist kurtoxin reduces hypoxia-mediated Ca²⁺ accumulation without affecting the activity of the Na⁺/Ca²⁺ exchanger (data not shown). Moreover, mibefradil significantly reduces hypoxia-mediated Ca²⁺ accumulation in cells exposed to cariporide. Therefore, in rat neonatal cardiomyocytes, the T-type Ca²⁺ channel is an important contributor to hypoxia-mediated Ca²⁺ accumulation.

Glucose-treated cells show a reduction in hypoxia-mediated Ca²⁺ accumulation that is only partially related to reduced Na⁺/Ca²⁺ exchanger activity. In the present study, the difference in the fura-2 fluorescence ratio between control and hyperglycemic-preconditioned cells (F_340/380) was about 0.9. In the presence of cariporide, which blocks hypoxia-mediated Ca²⁺ influx via the Na⁺/H⁺-Na⁺/Ca²⁺ exchanger combination, mibefradil reduced F_340/380 from 0.5 to 0.18. These data show that glucose-mediated reductions in T-type Ca²⁺ current also contribute to the reduction in [Ca²⁺]_i of the hypoxic cell.

In contrast to glucose treatment, exposure of the control cell to ANG II during the prehypoxic period promotes hypoxia-mediated Ca²⁺ accumulation. On the basis of F'_340/380, the fluorescence difference between the control and the ANG II-treated cell, ANG II elevates [Ca²⁺]_i of the hypoxic cell by enhancing Ca²⁺ influx via the Na⁺/H⁺-Na⁺/Ca²⁺ exchanger combination and the T-type Ca²⁺ channel.

ANG II also elevates hypoxia-mediated Ca²⁺ accumulation by the glucose-treated cell. Indeed, the ANG II-mediated increase in F_340/380 of the control and glucose-treated cells is comparable, indicating that glucose treatment has no effect on ANG II signaling. Therefore, it is not surprising that exposure of the glucose-treated cell to ANG II prevents hyperglycemic preconditioning. Several lines of evidence support the notion that Ca²⁺ influx via the two exchangers and the T-type Ca²⁺ channel play crucial roles in the reversal of hyperglycemic preconditioning by ANG II. First, ANG II reverses hyperglycemic preconditioning while activating transporters that enhance hypoxia-mediated elevations in [Ca²⁺]_i (Figs. 1, 10, and 11). Second, glucose treatment promotes cardioprotection while reducing hypoxia-mediated Ca²⁺ influx via the exchangers and the T-type Ca²⁺ channel (Figs. 1, 6, and 8). Third, the T-type Ca²⁺ channel antagonist mibefradil and the Na⁺/H⁺ exchange inhibitor cariporide reduce hypoxia-mediated Ca²⁺ accumulation in glucose-treated cells exposed to ANG II (Figs. 6 and 8). Finally, mibefradil and cariporide partially block the increase in hypoxia-mediated apoptosis arising from ANG II exposure (Fig. 12).

The present study may have pathological significance. Recently, Mozaffari and Schaffer (27) proposed that ANG II was a likely regulator of infarct size in the diabetic preconditioned heart. This hypothesis was based on the finding that elevations in afterload reversed prediabetic preconditioning. Because a major consequence of elevated afterload is cell stretching, which in turn causes the release of ANG II from the heart (33), the investigators proposed that ANG II was a logical candidate for the adverse effects of elevated afterload, particularly because ANG II activates Ca²⁺ transporters involved in hypoxia-mediated Ca²⁺ overload.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This study was supported by National Heart, Lung, and Blood Institute Grants HL-63723 (to S. Schaffer) and HL-74116 (to S. Wu).

	ACKNOWLEDGMENTS

 
We acknowledge Michelle Guyton for technical assistance with whole cell current recordings.

	FOOTNOTES

 

Address for reprint requests and other correspondence: S. Schaffer, Dept. of Pharmacology, Univ. of South Alabama College of Medicine, Mobile, AL 36688 (E-mail: sschaffe{at}jaguar1.usouthal.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.

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TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 

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