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Diphenylhydantoin Suppresses Glucose-Induced Insulin Release by Decreasing Cytoplasmic H⁺ Concentration in Pancreatic Islets

Department of Diabetes and Clinical Nutrition, Graduate School of Medicine, Kyoto University (K.N., S.F., M.S., R.K., Y.N., S.F., E.M., Y.Y., N.I.), Kyoto 606-8507, Japan; and Kansai-Denryoku Hospital (Y.S.), Osaka 553-0003, Japan

Address all correspondence and requests for reprints to: Dr. Shimpei Fujimoto, 54 Shogoin Kawahara-cho, Sakyo-ku, Kyoto 606-8507, Japan. E-mail: fujimoto{at}metab.kuhp.kyoto-u.ac.jp.

	Abstract

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Diphenylhydantoin (DPH), which is clinically used in the treatment of epilepsy, inhibits glucose-induced insulin release from pancreatic islets by a mechanism that remains unknown. In the present study, DPH is shown to suppress glucose-induced insulin release concentration-dependently. In dynamic experiments, 20 µM DPH suppressed 16.7 mM glucose-induced biphasic insulin release. DPH also suppressed insulin release in the presence of 16.7 mM glucose, 200 µM diazoxide, and 30 mM K⁺ without affecting the intracellular Ca²⁺ concentration. DPH suppressed ATP content and mitochondrial membrane hyperpolarization in the presence of 16.7 mM glucose without affecting glucose utilization, glucose oxidation, and reduced nicotinamide adenine dinucleotide phosphate fluorescence. DPH increased cytoplasmic pH in the presence of high glucose, but the increase was abolished under Na⁺-deprived conditions and HCO₃^–-deprived conditions, suggesting that Na⁺ and HCO₃^– transport across the plasma membrane are involved in the increase in cytoplasmic pH by DPH. Alkalization by adding NH₄⁺ to the extracellular medium also suppressed insulin release, ATP content, and mitochondrial membrane hyperpolarization. Because ATP production from the mitochondrial fraction in the presence of substrates was decreased by increased pH in the medium, DPH suppresses mitochondrial ATP production by reducing the H⁺ gradient across mitochondrial membrane. Using permeabilized islets, the increase in pH was shown to decrease Ca²⁺ efficacy at a clamped concentration of ATP in the exocytotic system. Taken together, DPH inhibits glucose-induced insulin secretion not only by inhibiting mitochondrial ATP production, but also by reducing Ca²⁺ efficacy in the exocytotic system through its alkalizing effect on cytoplasm.

	Introduction

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DIPHENYLHYDANTOIN (DPH) is widely used clinically in treatment of epilepsy. Oral administration of DPH results in a higher blood glucose response and diminution of the early and late insulin responses in man (1). Patients with hyperglycemic hyperosmolar nonketotic coma after administration of DPH have been reported (2). Patients with hypoglycemia caused by inslinoma have been treated with the agent to reduce hyperinsulinemia (3, 4). These phenomena may originate from the suppressive effect of DPH on insulin release from pancreatic islets. In perfused pancreas (5) and isolated pancreatic islets (6), glucose-induced insulin release is suppressed by the agent.

The mechanism of glucose-stimulated insulin release from pancreatic ß-cells has recently been well documented. Glucose stimulates insulin secretion by both triggering and amplifying signals in pancreatic ß-cells (7). The triggering pathway includes entry of glucose into ß-cells, acceleration of glycolysis in cytosol and glucose oxidation in mitochondria, increases in ATP content and ATP/ADP ratio, closure of ATP-sensitive K⁺ channels (K_ATP channels), membrane depolarization, opening of voltage-dependent Ca²⁺ channels (VDCCs), increase in Ca²⁺ influx through VDCCs, elevated intracellular Ca²⁺ concentration ([Ca²⁺]_i), and exocytosis of insulin granules. It has been reported that glucose also enhances insulin secretion K_ATP channel-independently. The K_ATP channel-independent, amplifying action of glucose has been confirmed by treatment of ß-cells with diazoxide, which prevents K_ATP channels from closing, and with a depolarizing concentration of extracellular K⁺, restores Ca²⁺ influx (7). Because glucose does not increase [Ca²⁺]_i, but, nevertheless, augments insulin release under these conditions, glucose may well exert its effects by increasing Ca²⁺ efficacy in stimulation-secretion coupling due at least in part to the direct effect of increased ATP derived from glucose metabolism on exocytosis (7).

The mechanism of the inhibitory effect of DPH on glucose-induced insulin release has been described. DPH causes hyperpolarization of the plasma membrane in islet cells (8), which also can be brought about by the stimulatory effect of the agent on Na⁺,K⁺-adenosine triphosphatase (Na⁺,K⁺-ATPase) activity in plasma membrane (9) or the inhibitory effect on Na⁺ accumulation in isolated islets (6). However, the role of the stimulatory effect of DPH on Na⁺,K⁺-ATPase activity in the inhibition of glucose-induced insulin release is unclear, because DPH fails to affect ⁸⁶Rb net uptake in the presence of high glucose, which reflects K⁺ influx in islets (10). The effect of DPH on glycolysis in islets is controversial. In one study, DPH reduced glucose utilization in islets in the presence of high glucose (11), but in another study, it was not affected by the agent (12). DPH reduces both ⁴⁵Ca uptake in islets and ⁴⁵Ca outflow from islets in the presence of high glucose (10), suggesting that DPH inhibits glucose-induced insulin release by interfering with Ca²⁺ uptake via VDCCs (12).

In the present study we have examined the mechanism of the inhibition of glucose-induced insulin release from pancreatic islets by DPH.

	Materials and Methods

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Animals
Male Wistar rats, weighing 180–230 g, were obtained from Shimizu Co. (Kyoto, Japan). The animals were fed standard laboratory chow ad libitum and allowed free access to water in an air-conditioned room with a 12-h light, 12-h dark cycle until they were used in the experiments. All experiments were carried out with rats 8–12 wk of age. The animals were maintained and used in accordance with the Guidelines for Animal Experiments of Kyoto University.

Islet isolation and culture
Islets of Langerhans were isolated from Wistar rats by collagenase digestion as described previously (13). Isolated islets were cultured for 12 h in RPMI 1640 medium containing 10% fetal calf serum, 100 U/ml penicillin, 100 µg/ml streptomycin, and 5.5 mM glucose at 37 C in humidified air containing 5% CO₂.

Measurement of insulin release from isolated rat pancreatic islets
Insulin release from intact islets was monitored using either batch incubation or a perifusion system using Krebs-Ringer bicarbonate buffer (KRBB) supplemented with 0.2% BSA (fraction V) and 10 mM HEPES adjusted to pH 7.4 (KRBB medium). The detailed method of batch incubation and perifusion of islets to monitor insulin release was described previously (13). For batch incubation experiments, islets were preincubated at 37 C for 30 min in KRBB medium supplemented with 2.8 mM glucose. Groups of five islets then were batch-incubated for 30 min in 0.7 ml KRBB medium with test materials. At the end of the incubation period, islets were pelleted by centrifugation, and aliquots of the buffer were sampled. For perifusion experiments, groups of 20 islets were placed in parallel chambers (400 µl each) of a perifusion apparatus and perifused with the same medium at a rate of 0.7 ml/min at 37 C. The medium was continuously gassed with 95% O₂ and 5% CO₂. Islets were perifused for 30 min to establish a stable insulin secretory rate at the basal level of glucose. DPH and the stimulating dose of glucose were added to the medium according to each experimental protocol. The samples were collected at the times indicated in the figures. The amount of immunoreactive insulin was determined by RIA, using rat insulin as a standard.

Measurement of [Ca²⁺]_i
The method of measurement of [Ca²⁺]_i using fura-2/acetoxymethyl ester (AM)-loaded dispersed cells was described previously (13). In brief, cultured islets were washed in PBS and incubated with 0.25% trypsin and 1 mM EDTA solution (Invitrogen Life Technologies, Inc., Grand Island, NY) for 2 min at 37 C. Digestion was terminated by rinsing the cells in cold PBS. They were washed in PBS again, placed on small glass coverslips (15 min; 4 mm), coated with Cell-Tak (Collaborative Biomedical Products, Bedford, MA) to accelerate cell adhesion, and incubated in KRBB medium containing 2.8 mM glucose in humidified air containing 5% CO₂ at 37 C for at least 15 min. Fura-2/AM (1 µM; Molecular Probes, Eugene, OR) was then loaded with dispersed islet cells for 30 min at 37 C. A heat-controlled chamber on the stage of an inverted microscope kept at 36 ± 1 C was superfused with KRBB medium containing 2.8 mM glucose. The cells were exited successively at 340 and 380 nm, and the fluorescence emitted at 510 nm was captured by CCD camera (Micro Max 5 MHz System; Roper Industries, Trenton, NJ). One second was used to acquire a 340/380 image pair, including the duration for filter exchange. The images were analyzed with Meta Fluor image analyzing system (Universal Imaging Corp., West Chester, PA). The 340-nm (F340) and 380-nm (F380) fluorescence signals were detected every 10 sec, and the ratios (F340/F380) were calculated. In vitro calibration was performed using a fura-2 calcium imaging calibration kit (Molecular Probes), and F340/F380 was converted into calibrated values of [Ca²⁺]i.

Measurement of ATP content
After groups of 10 islets were preincubated with 2.8 mM glucose for 30 min, they were incubated in tubes for 30 min in 0.5 ml KRBB medium with 16.7 or 2.8 mM glucose with test materials. The reaction was stopped by immediate addition and mixing of 0.1 ml 2 M HClO₄ to the tubes, which were immediately mixed by vortex and sonicated in ice-cold water for 3 min. They were then centrifuged (3000 x g, 3 min), and a fraction (0.4 ml) of the supernatant was mixed with 100 µl 2 M HEPES and 100 µl 1 M Na₂CO₃.

The ATP content in islets was determined by luminometric method as previously described (14). A fraction of the extracts (0.1 ml) was diluted with 0.1 ml 20 mM HEPES solution (pH 7.4; with NaOH). The ATP concentration in the solutions was measured by adding 0.1 ml luciferin-luciferase solution in a bioluminometer (Luminometer model 20e, Turner Designs, Sunnyvale, CA). To draw a standard curve, blanks and ATP standards were run through the entire procedure, including the extraction steps.

Measurement of glucose utilization and oxidation
Glucose utilization and oxidation were measured as previously described (15). For utilization, cultured islets were preincubated in KRBB medium with 2.8 mM glucose at 37 C for 30 min. Batches of 30 islets for each condition were incubated at 37 C for 90 min in 150 µl medium containing 1.5 µCi [5-³H]glucose (Amersham Biosciences, Little Chalfont, UK). Aliquots of the incubation medium (100 µl) and 20 µl 1 M HCl were transferred into small tubes and placed into a glass vial containing 0.5 ml water. The capped vials were incubated overnight at 34 C to allow ³H₂O in the inner tubes to equilibrate with the water in the outer vials. Afterward, the inner tube was lifted out, and the disintegrations per minute of ³H₂O in the water were counted. In parallel incubation, the recovery ratio of ³H₂O (Amersham Biosciences) was measured. After subtracting blank disintegrations per minute from sample disintegrations per minute, glucose utilization was calculated using the disintegrations per minute, specific radioactivity of [5-³H]glucose, and recovery ratio of ³H₂O. For oxidation, all procedures were the same as those for utilization, except for the use of [U-¹⁴C]glucose (0.5 µCi/tube) and 0.5 ml hydroxide of hyamine 10-X (Packard, Meriden, CT) in place of [5-³H]glucose and 0.5 ml water, respectively. After subtracting blank disintegrations per minute from sample disintegrations per minute, glucose oxidation was calculated using the disintegrations per minute and specific radioactivity of [U-¹⁴C]glucose.

Fluorescence measurement of mitochondrial membrane potential
Mitochondrial membrane potential (m) was measured by 5,5', 6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1) fluorescence as previously reported (16). Isolated islets were incubated for 15 min in KRBB medium containing 2.8 mM glucose at room temperature in the dark with 10 µg/ml JC-1. The islets were washed in PBS and incubated with 0.25% trypsin and 1 mM EDTA solution (Invitrogen Life Technologies, Inc., Grand Island, NY) for 3 min at 37 C, diluted by 20 ml cold PBS, and dispersed by pipetting. The dispersed islet cells were applied to glass cuvettes. After preincubation in KRBB medium with 2.8 mM glucose for 20 min at 37 C, fluorescence was determined using a spectrofluorophotometer (RF 5000, Shimadzu, Kyoto, Japan) with excitation wavelength at 490 nm and emission wavelength at 590 nm and with stirring medium containing dispersed cells in cuvettes at 37 C. At time zero, basal fluorescence was determined, when glucose and DPH or NH₄Cl, at final concentrations of 16.7 mM and 20 µM or 3 mM, respectively, were added. Cuvettes were incubated in humidified air containing 5% CO₂ at 37 C, and determinations were performed at the time indicated in the figures. Fluorescence was corrected by subtracting parallel blanks in which islet cells were not loaded with JC-1.

Measurement of cytoplasmic pH and reduced nicotinamide adenine dinucleotide (phosphate) [NAD(P)H] fluorescence
Cytoplasmic pH was measured using the previously described method (17) with slight modifications. 2',7'-Bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein AM (1 µM) was loaded with cultured islets for 40 min at 37 C. The islets were immediately placed in a heat-controlled chamber on the stage of an inverted microscope kept at 36 ± 1 C and superfused with KRBB medium with 2.8 mM glucose for 30 min. The islets were excited successively at 440 and 490 nm, and the fluorescence emitted at 535 nm was captured by CCD camera (Micro Max 5 MHz System, Roper Industries). The 440 nm (F440) and 490 nm (F490) fluorescence signals were detected every 10 sec, and the ratios (F490/F440) were calculated. Calibration of cytoplasmic pH was performed using the K⁺/H⁺ ionophore nigericin (18), which equilibrates proton and potassium gradients over the cell membranes according to the formula: [H⁺]_i = [H⁺]_ox([K⁺]_i/[K⁺]_o) (19). Islets were exposed to a solution containing 10 µM nigericin and 115 mM K⁺, which was reported to be identical with [K⁺]_i in pancreatic ß-cells (20). A three-point calibration was performed by changing pH₀ from 7.5 to 7.0 and 6.5. In the nigericin-permeabilized islets, the fluorescence values reached a new steady state within 5 min, and pH_i was assumed to be identical with pH_o. The fluorescence ratio (F490/F440) was plotted against pH_i and fitted by a linear regression. The observed pH_i values in the presence of 2.8 mM glucose and those in the presence of 16.7 mM were nearly equal to reported values in islets (17). In some experiments, NaCl and NaHCO₃ were replaced by choline chloride and choline bicarbonate, respectively, and 5 µM atropine sulfate was added to prevent cholinergic effects.

For NAD(P)H measurement, the islets without dye were excited successively at 360 nm, and the fluorescence emitted at 470 nm was imaged as previously described (21). Changes in NADPH fluorescence signal are expressed as a percentage of control values by dividing the signal at a given time by the average signal at 2.8 mM glucose during the last 1 min before stimulation. Images of cytoplasmic pH and NAD(P)H were analyzed with Meta Fluor image analyzing system (Universal Imaging, West Chester, PA).

Measurement of mitochondrial ATP production
The mitochondrial suspension from isolated islets was prepared by repeated centrifugation, as previously reported (16). First, isolated islets were homogenized in solution A consisting of 50 mM HEPES, 100 mM KCl, 1.8 mM ATP, 1 mM EGTA, 2 mM MgCl₂, and 0.5 mg/ml BSA (electrophoretically homogeneous). After precipitation of cell debris and nuclei by centrifugation, the supernatant was more rapidly centrifuged (10,000 x g) to obtain a pellet containing the mitochondrial fraction. Afterward, the precipitation diluted by 200 µl solution A was centrifuged again and finally rinsed three times in the solution consisting of 20 mM HEPES, 3 mM KH₂PO₄, 1 mM EGTA, 20 mM gluconate sodium, 0.3 mM MgCl₂, 130 mM gluconate potassium, and 0.5 mg/ml BSA (electrophoretically homogeneous; medium B) adjusted to pH 7.0 with KOH. The mitochondrial fraction in 500 µl medium B was kept on ice until use. The reaction was started by adding 5 µl mitochondrial suspension to 495 µl prewarmed medium B (adjusted to each pH at 37 C with gluconate or KOH) supplemented with 0.5 mM succinate plus 0.5 mM pyruvate or 1 mM glycerol 3-phosphate, 50 µM ADP, and 1 µM diadenosine pentaphosphate (DAPP). DAPP is a specific inhibitor of adenylate kinase used to measure ATP production exclusively by oxidative phosphorylation. After 30-min incubation at 37 C, the reaction was stopped by the addition of 0.5 µM antimycin A. The samples were cooled to room temperature, and the ATP concentration in the solutions was measured by adding luciferin-luciferase solution to each sample with a bioluminometer (Luminometer model 20e, Turner Designs). To draw a standard curve, blank and ATP standards were added to parallel samples containing the complete incubation mixture, except for the mitochondrial suspension.

Measurement of insulin release from permeabilized islets
Measurement of insulin release from permeabilized islets was performed using the method previously described (22). Cultured islets were preincubated with KRBB medium containing 2.8 mM glucose for 30 min. They were washed twice in cold potassium gluconate buffer (KG buffer) containing 134 mM gluconate potassium, 4 mM gluconate sodium, 6 mM NaH₂PO₄, 3 mM MgCl₂, 2.5 mM EGTA, 30 mM HEPES, and 0.5% BSA (adjusted to each pH with gluconate or KOH), with CaCl₂ added to a Ca²⁺ concentration of 30 nM. The islets were permeabilized by high voltage discharge (four exposures each of 450-µsec duration to an electrical field of 4.0 kV/cm) in KG buffer and washed once with the same buffer. Groups of electrically permeabilized islets then were batch-incubated for 30 min at 37 C in 0.4 ml KG buffer with 5 mM ATP and various concentrations of Ca²⁺ and H⁺. The Ca²⁺ concentrations at pH 7.0 were determined as previously described (13). The amount of Ca²⁺ added to the medium at pH other than 7.0 was determined by monitoring the free Ca²⁺ concentration using a Ca²⁺ electrode (Horiba, Kyoto, Japan) and titrating the CaCl₂ solution to each medium to achieve the same voltage value as the pH 7.0 medium at 37 C. At the end of the incubation period, permeabilized islets were pelleted by centrifugation (15,000 x g, 180 sec), and aliquots of the buffer were sampled for insulin determination.

Materials
RPMI 1640 medium, diazoxide, carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), ADP, nigericin sodium, DAPP, gluconate, antimycin A, and glycerol 3-phosphate were purchased from Sigma-Aldrich Corp. (St. Louis, MO). Succinate was purchased from Sigma-Aldrich Corp. (Steimheim, Germany). Luciferin-luciferase was obtained from Turner Designs. Fura-2/AM, JC-1, and 2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein/AM were purchased from Molecular Probes. All other agents, including 5,5-diphenylhydantoin sodium salt, were obtained from Nacalai Tesque (Kyoto, Japan).

Statistical analysis
Results are expressed as the mean ± SE. Statistical significance was evaluated by unpaired Student’s t test. P < 0.05 was considered significant.

	Results

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Effect of DPH on insulin release from pancreatic islets
DPH inhibited 16.7 mM glucose-stimulated insulin release from islets in a concentration-dependent manner, but did not affect insulin release in the presence of 2.8 mM glucose (Table 1A). Significant inhibition of insulin release was detected at 5 µM. Insulin release at 16.7 mM glucose was inhibited to the level of basal release by 80 µM DPH. The K_ATP channel-independent, amplifying action of glucose on insulin release has been confirmed by treatment of ß-cells with diazoxide, which prevents K_ATP channels from closing, and with a depolarizing concentration of extracellular K⁺ that restores Ca²⁺ influx. To evaluate the inhibitory effect of DPH on the amplifying action of glucose on insulin release, insulin release was measured in the presence of diazoxide and a depolarizing concentration of K⁺. DPH also suppressed insulin secretion in the presence of high of glucose, diazoxide, and high K⁺ (Table 1B). In the perifusion experiments, insulin release in the presence of 2.8 mM glucose was not altered by 20 µM DPH. However, 20 µM DPH suppressed the 16.7 mM glucose-induced biphasic insulin release during both the first and second phases (Fig. 1A). Insulin release in the presence of 16.7 mM glucose, 200 µM diazoxide, and 30 mM K⁺ was significantly suppressed about 5 min after application of 20 µM DPH, and the effect was reversed about 15 min after withdrawal of the agent (Fig. 1B). DPH (20 µM) also suppressed 30 mM K⁺-induced monophasic insulin release in the presence of 2.8 mM (Fig. 1C).

View larger version (21K): [in this window] [in a new window]   FIG. 1. Inhibitory effect of DPH on insulin release from intact islets. Values represent the mean ± SE. A, Time course of the effect of 20 µM DPH on biphasic 16.7 mM glucose-induced insulin release. Two groups of islets were perifused with 2.8 mM glucose (G) for 30 min and stimulated with 16.7 mM glucose at 0 min for 30 min in the presence (; n = 5) or absence (; n = 5) of DPH. DPH was introduced 15 min before 16.7 mM glucose administration. Values measured from 4–30 min are significantly less in DPH-treated islets than the corresponding values in control islets (P < 0.01). B, Time course of the effect of 20 µM DPH on insulin release in the presence of 16.7 mM glucose (G), 200 µM diazoxide (Dz), and a depolarizing concentration (30 mM) of K+. After two groups of islets were perifused for 15 min (–15 to 0 min) in the presence of 16.7 mM glucose, 200 µM diazoxide, and 5 mM K+, they were stimulated with 30 mM K+ at time zero in the presence of 16.7 mM glucose and 200 µM diazoxide for 70 min (0–70 min). One group of islets was exposed to 20 µM DPH from 10–40 min (; n = 7), and the other group continued to be perifused without DPH (; n = 7). The withdrawal effect of DPH for 30 min (from 40–70 min) is also shown. Values measured from 12–50 min are significantly less in DPH-treated islets than the corresponding values in control islets (P < 0.01). C, Time course of the effect of 20 µM DPH on monophasic 30 mM K+-induced insulin release in the presence of 2.8 mM glucose. Two groups of islets were perifused with 2.8 mM glucose for 30 min and stimulated with 30 mM K+ at 0 min in the presence of 2.8 mM glucose for 30 min in the presence (; n = 5) or absence (; n = 5) of DPH. DPH was introduced 10 min before 30 mM K+ administration. Values measured at 3, 4, 5, 7, 9, 12, 18, 22, and 26 min are significantly less in DPH-treated islets than the corresponding values in control islets (P < 0.05). Average values from 0–30 min were significantly less in DPH-treated islets than the corresponding values in control islets (control, 26.9 ± 2.3; DPH, 17.8 ± 1.3 pg/islet·min, P < 0.01).

View larger version (19K): [in this window] [in a new window]   FIG. 2. Time course of the effect of 20 µM DPH on [Ca2+]i in dispersed islet cells. Cells that responded to 16.7 mM glucose were regarded as ß-cells. Values represent the mean ± SE. A, Effect of DPH in the presence of 16.7 mM glucose (G). The glucose concentration was raised from 2.8 to 16.7 mM at time zero. DPH was introduced to DPH-treated cells (; n = 11) and was not administrated to control cells (; n = 12) 10 min before 16.7 mM glucose administration. The values after 5 min were significantly less in DPH-treated islets than the corresponding values in control islets (P < 0.01). Average [Ca2+]i of DPH-treated cells from –5 to 0 min was similar to that of control cells (control, 84.7 ± 3.6; DPH, 83.3 ± 2.5 nM). The average [Ca2+]i of DPH-treated cells from 0–30 min was significantly less than that of control cells (control, 150.4 ± 5.7; DPH, 118.6 ± 3.3 nM; P < 0.01). B, Effect of DPH in the presence of 16.7 mM glucose (G), 200 µM diazoxide (Dz), and a depolarizing concentration (30 mM) of K+. The glucose concentration was raised from 2.8 to 16.7 mM at time zero. Diazoxide and a depolarizing concentration of K+ were introduced from 10 min. DPH was administrated to DPH-treated cells (; n = 25) from 15–30 min and was not administrated to control cells (; n = 17). None of the values indicated was significantly different in the two groups. The average [Ca2+]i of DPH-treated cells was similar to that of control cells [from –5 to 0 min: control, 73.9 ± 4.6; DPH, 75.1 ± 2.9 (not significant); from 0 to 10 min: control, 143.2 ± 5.7; DPH 140.3 ± 4.1 (not significant); from 10 to 15 min: control, 157.1 ± 7.2; DPH 153.5 ± 4.0 (not significant); from 15–30 min: control, 175.4 ± 6.4; DPH 167.1 ± 4.0 nM (not significant)]. C, Effect of DPH in the presence of 2.8 mM glucose (G) and a depolarizing concentration of (30 mM) K+. The K+ concentration was raised from 5 to 30 mM at time zero in the presence of 2.8 mM glucose (G). DPH was introduced to DPH-treated cells (; n = 17) and not to control cells (; n = 15) 10 min before 30 mM K+ administration. All values indicated were similar in the two groups. Average [Ca2+]i of DPH-treated cells from –5 to 0 min and from 0–30 min were similar to those of control cells (from –5 to 0 min: control, 82.5 ± 4.4; DPH, 86.2 ± 3.6; from 0–30 min: control,143.1 ± 4.6; DPH, 138.4 ± 3.6 nM).

View larger version (37K): [in this window] [in a new window]   FIG. 3. Time course of the effect of 20 µM DPH (A–E) and 3 mM NH4+ (F) on cytoplasmic pH in intact islets. Values represent the mean ± SE. After islets were perifused with 2.8 mM glucose (G) for 30 min, exposure to 16.7 mM glucose was begun 15 min before application of DPH and NH4+ (A–D and F). A, In the presence of 16.7 mM glucose, one group of islets was exposed to 20 µM DPH from 0–6 min (; n = 16), and the other group continued to be perifused without DPH (; n = 11). The effect of withdrawal of DPH for 4 min (from 6–10 min) is also shown. Values of DPH-treated islets at 1, 2, 3, 4, 5, 6, and 7 min are significantly greater than the corresponding values in control islets (P < 0.01). B, In the presence of 16.7 mM glucose, 200 µM diazoxide, and 30 mM K+, which were applied 15 min before application of DPH, one group of islets was exposed to 20 µM DPH from 0–8 min (; n = 20), and the other group continued to be perifused without DPH (; n = 17). The effect of withdrawal of DPH for 5 min (from 8–13 min) is also shown. Values of DPH-treated islets at 1, 2, 3, 4, 5, 6, 7, 8, and 9 min are significantly greater than the corresponding values in control islets (P < 0.01). C, In the presence of 16.7 mM glucose without HCO3–, one group of islets was exposed to 2 µM DPH from 0 min (; n = 16), and the other group continued to be perifused without DPH (; n = 18). NH4+ (10 mM) was applied to both groups of islets from 8 min. Values of DPH-treated islets indicated every 1 min are not significantly greater than the corresponding values in control islets. D, In the presence of 16.7 mM glucose without Na+, one group of islets was exposed to 20 µM DPH from 0–8 min (; n = 44), and the other group continued to be perifused without DPH (; n = 32). NH4+ (10 mM) was applied to both groups of islets from 14 min. Values of DPH-treated islets indicated every 1 min are not significantly greater than the corresponding values in control islets. E, Effect of DPH in the presence of 2.8 mM glucose. After islets were perifused with 2.8 mM glucose for 20 min, one group of islets was exposed to 20 µM DPH from 0 min (; n = 12), and the other group continued to be perifused without DPH (; n = 10). The K+ concentration were increased from 5–30 mM from 10 min. The indicated values for DPH-treated islets after 2.5 min were significantly greater than the corresponding values for control islets (P < 0.05). F, Effect of 3 mM NH4+ in the presence of 16.7 mM glucose. In the presence of 16.7 mM glucose, one group of islets was exposed to 3 mM NH4+ from 0 min (; n = 5), and the other group continued to be perifused without NH4+ (; n = 6). Values of NH4+-treated islets after 2.5 min were significantly greater than the corresponding values in control islets (P < 0.01).

View larger version (19K): [in this window] [in a new window]   FIG. 4. Time course of the effect of 20 µM DPH on NAD(P)H fluorescence in the presence of 16.7 mM glucose (G) in intact islets. After perifusion with 2.8 mM glucose for 30 min, the islets were exposed to 16.7 mM glucose at time zero. One group of islets was exposed to 20 µM DPH from 5 min (; n = 13), and the other group continued to be perifused without DPH (; n = 13). Changes in NAD(P)H fluorescence signal were calculated as a percentage of control values by dividing the signal at a given time by the average signal at 2.8 mM glucose during the last 1 min before stimulation. Values represent the mean ± SE. Values in DPH-treated islets indicted in the figure are not significantly different from the corresponding values in control islets.

Effects of DPH and NH₄⁺ on m

View larger version (21K): [in this window] [in a new window]   FIG. 5. Time course of the effects of DPH and NH4+ on m in the presence of 16.7 mM glucose. JC-1-loaded, dispersed islet cells were preincubated in KRBB medium with 2.8 mM glucose for 30 min. At time zero, basal fluorescence was determined, and cells were incubated in the presence of 16.7 mM glucose with () or without () 20 µM DPH and 3 mM NH4+. , Incubation with 2.8 mM glucose throughout. At 60 min, FCCP (final concentration, 1 µM) was added to the medium. Results are expressed as the mean change in fluorescence (arbitrary units) compared with that at time zero. *, P < 0.01, 16.7 mM glucose vs. 16.7 mM glucose plus DPH (A) or 16.7 mM glucose plus NH4+ (B). , P < 0.01, 16.7 mM glucose vs. 2.8 mM glucose. A, Effect of 20 µM DPH. Values are expressed as the mean ± SE of nine determinations. B, Effect of 3 mM NH4+. Values are expressed as the mean ± SE of eight determinations.

	Discussion

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In the present study, 20 µM DPH, a lower concentration than that used in previous studies (5, 8, 10, 11, 12) (80 µM), was used, because even 5 µM DPH suppressed high glucose-induced insulin release. DPH inhibited glucose-induced insulin release in the presence of high glucose, diazoxide, and a depolarizing concentration of K⁺, which demonstrates that the inhibitory effect of DPH does not necessarily require reduced [Ca²⁺]_i.

In pancreatic ß-cells, intracellular Ca²⁺ and ATP are the most important regulators of insulin secretion (7). DPH decreases the ATP content in the presence of high glucose. Mitochondrial ATP production is driven by the proton-motive force that includes the m generated by the electron transport chain, and the rate of ATP synthesis in mitochondria is closely correlated with m (23). Because DPH reduces the hyperpolarizing effect of glucose on m, the compound clearly reduces the H⁺ gradient across the mitochondrial membrane.

The H⁺ gradient is brought about by transport of high-energy electrons in the respiratory chain. These electrons are derived from NADH and FADH₂, generated in the Krebs cycle in the matrix and/or transferred from cytosol by the shuttle system. A decrease in the supply of substrates to the mitochondria results in a decreased H⁺ gradient across the mitochondrial membrane. Glucose utilization reflects the velocity of glycolysis (24), and NAD(P)H autofluorescence dominantly reflects the redox state of mitochondria (25). Because DPH did not affect glucose utilization or NAD(P)H fluorescence in the presence of high glucose, DPH did not affect the supply of reduced equivalents to mitochondria. In addition, DPH did not affect ATP production from mitochondrial fraction. Because increased pH in the medium is reported to decrease the H⁺ gradient across the inner membrane of isolated rat heart mitochondria (26), we investigated the effect of DPH on the cytoplasmic H⁺ concentration. DPH increased cytoplasmic pH in the presence of high glucose immediately and reversibly. Alkalization by adding NH₄⁺ to the extracellular medium also suppressed insulin release, ATP content, and mitochondrial membrane hyperpolarization. Moreover, ATP production from the mitochondrial fraction in the presence of pyruvate and succinate and in the presence of glycerol 3-phosphate, by which the supply of mitochondrial reduced equivalents is derived mainly from the Krebs cycle and the glycerol phosphate shuttle, respectively, was decreased by increasing pH in the medium. These results indicate that DPH in the presence of high glucose increases cytoplasmic pH, which suppresses mitochondrial ATP production by reducing the H⁺ gradient across the mitochondrial membrane.

High glucose increased NAD(P)H fluorescence during the initial 5 min, which suggests that increased NADH production overwhelms NADH utilization. After 5 min of high-glucose exposure, it reaches a plateau, suggesting that increased NADH production balances increased NADH utilization, which eventually increases ATP production. During the plateau period, DPH did not affect NADH fluorescence. Thus, DPH might decrease NADH production and utilization simultaneously, resulting in unchanged NADH fluorescence and decreased ATP production. To rule out this possibility, glucose oxidation was measured. Glucose oxidation reflects the velocity of glucose-derived metabolite metabolism in the Krebs cycle that constitutes the major source of NADH production. NH₄⁺ (3 mM) did not affect glucose oxidation, which is compatible with a previous result using 10 mM NH₄⁺ (27). Because DPH also did not affect glucose oxidation, both NADH production and utilization were unaffected by cellular alkalization, suggesting that cellular alkalization reduces the efficacy of NADH utilization in producing ATP. This may result from the reduction of H⁺-gradient generated by NADH utilization due to diffusion of H⁺ near the cytoplasmic side of the mitochondrial innermembrane to a cytoplasmic milieu in which the H⁺ concentration is reduced.

The effects of intracellular pH on insulin release in the presence of a basal level of glucose were also reported (28). In the presence of a basal level of glucose, insulin release was not changed by alteration of intracellular pH when the intracellular pH was less than 7.5. In the presence of 2.8 mM glucose, 20 µM DPH also elevated cytoplasmic pH. Consistent with these results, in the present study, insulin release in the presence of a basal level (2.8 mM) of glucose was unaltered by 20 µM DPH, which increased intracellular pH to approximately 7.2.

The effects of increasing cytoplasmic pH on insulin release from islets and pancreatic ß-cells have been examined. Glucose-induced, time-dependent potentiation of insulin release is inhibited by increasing intracellular pH (29). The addition of more than 10 mM NH₄⁺ to the extracellular medium decreases glucose-induced insulin release (30, 31, 32), increases cytoplasmic pH (30, 31, 33), hyperpolarizes the plasma membrane (31, 34), and decreases [Ca²⁺]_i (31). Hyperpolarization due to cytoplasmic alkalization has been shown to be derived from augmentation of K_ATP channel activity in experiments using intact ß-cells (31, 35). However, in experiments using excised patch, alkalization decreases K_ATP channel activity at clamped ATP levels (35, 36). This discrepancy might be explained by our present results: cytoplasmic alkalization suppresses mitochondrial ATP production by reducing the H⁺ gradient across the inner membrane, and the subsequent decrease in intracellular ATP concentration augments K_ATP channel activity. Consistently, in the present study DPH suppressed high glucose-induced Ca²⁺ elevation, but did not suppress K⁺-induced Ca²⁺ elevation in the presence of basal glucose.

Ca²⁺ and ATP directly affect the exocytotic system and enhance insulin release synergistically in experiments using single ß-cells (37, 38) and permeabilized islets (39). To quantify Ca²⁺ efficacy at clamped concentrations of ATP in the exocytotic process of insulin secretory granules directly, islets were electrically permeabilized to manipulate the intracellular Ca²⁺ and ATP concentration according to the extracellular medium, and insulin release was examined. DPH had no effect on Ca²⁺ efficacy at a clamped concentration of ATP in the exocytotic system. In addition, decreased intracellular ATP reduces Ca²⁺ efficacy in the exocytotic system (37, 38, 39). Accordingly, the lower ATP level due to reduced ATP production by cytoplasmic alkalization plays a role in the attenuation of insulin secretion from DPH-treated islets in response to high glucose. Biphasic glucose-induced insulin secretion can be described as the release of distinct pools of granules. The first phase of insulin secretion represents the release of readily releasable (primed/docked) granules located immediately below the plasma membrane. The second slower phase results from the time- and ATP-dependent mobilization of granules situated farther away from the plasma membrane (37). Because both phases are Ca²⁺ dependent, the precise site at which ATP and H⁺ modulate Ca²⁺ efficacy in these processes cannot be determined in the present study.

In the presence of 2.8 mM glucose, 20 µM DPH suppressed K⁺-induced monophasic insulin release without affecting Ca²⁺ and ATP levels, which is compatible with the finding that increased pH decreased insulin release from permeabilized islets at clamped concentrations of ATP and Ca²⁺. Consistent with our findings, important roles of secretory granule pH in exocytosis of insulin granules have been proposed recently. Blocking V-type H⁺-ATPase by bafilomycin increases secretory granule pH (40) and decreases exocytosis of insulin granules measured by cell capacitance (41) and by insulin release from permeabilized ß-cells (42). However, the fact that protein kinase A activation, which potentiates insulin release, increases secretory granule pH (40) indicates that granular pH is not the sole regulatory factor in the exocyotsis of insulin granules.

Our results demonstrate that DPH decreases Ca²⁺ efficacy in the exocytotic system by reducing both the cytoplasmic ATP level and the cytoplasmic H⁺ level, and clearly show that the inhibitory effect on insulin release of the agent does not necessarily require reduced [Ca²⁺]_i.

In pancreatic ß-cells, Na⁺/H⁺ exchange and Na⁺-dependent Cl^–/HCO₃^– exchange may play an important role in the regulation of cytoplasmic pH (17, 43). In the present study, DPH did not affect intracellular pH in Na⁺-deprived and HCO₃^–-deprived conditions, in which application of a permeable weak base, NH₄⁺, raised cytoplasmic pH. These results suggest that Na⁺ and HCO₃^– transport across the plasma membrane is involved in the increase in cytoplasmic pH by DPH. Additional study is necessary to elucidate precise mechanism of cytoplasmic alkalization by DPH.

	Acknowledgments

 
We thank Mr. S. Akagi and Mr. T. Yamaguchi for technical assistance.

	Footnotes

 
This work was supported in part by Grants-in-Aids for Scientific Research, Grants-in-Aids for Creative Scientific Research (15GS0301), and Grants for Leading Project for Biosimulation from the Ministry of Education, Culture, Sports, Science, and Technology of Japan; Health and Labor Sciences Research Grants for Research on Human Genome; Tissue Engineering Food Biotechnology and Health and Labor Sciences Research Grants for Comprehensive Research on Aging and Health from the Ministry of Health, Labor, and Welfare of Japan; and Establishment of International Center of Excellence (COE) for Integration of Transplantation Therapy and Regenerative Medicine (COE Program of the Ministry of Education, Culture, Sports, Science, and Technology of Japan).

The authors have nothing to disclose.

First Published Online March 9, 2006

Abbreviations: AM, Acetoxymethyl ester; [Ca²⁺]_i, intracellular Ca²⁺ concentration; DAPP, diadenosine pentaphosphate; DPH, diphenylhydantoin; FCCP, carbonyl cyanide p-trifluoromethoxyphenylhydrazone; JC-1, 5,5', 6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide; KG buffer, potassium gluconate buffer; KRBB, Krebs-Ringer bicarbonate buffer; m, mitochondrial membrane potential; NADPH, reduced nicotinamide adenine dinucleotide phosphate; Na⁺,K⁺-ATPase, Na⁺,K⁺-adenosine triphosphatase; VDCC, voltage-dependent Ca²⁺ channel.

Received October 4, 2005.

Accepted for publication March 1, 2006.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 

1. Malherbe C, Burrill KC, Levin SR, Karam JH, Forsham PH 1972 Effect of diphenylhydantoin on insulin secretion in man. N Engl J Med 286:339–342[Medline]
2. Goldberg EM, Sanbar SS 1969 Hyperglycemic, nonketotic coma following administration of dilantin (diphenylhydantoin). Diabetes 18:101–106[Medline]
3. Hofeldt FD, Dippe SE, Levin SR, Karam JH, Blum MR, Forsham PH 1974 Effects of diphenylhydantoin upon glucose-induced insulin secretion in three patients with insulinoma. Diabetes 23:192–198[Medline]
4. Pelkonen R, Taskinen MR 1973 Effect of diphenylhydantoin on plasma-insulin in insulinoma. Lancet 1:604–605[Medline]
5. Levin SR, Booker Jr J, Smith DF, Grodsky GM 1970 Inhibition of insulin secretion by diphenylhydantoin in the isolated perfused pancreas. J Clin Endocrinol Metab 30:400–401[Medline]
6. Kizer JS, Vargas-Gordon M, Brendel K, Bressler R 1970 The in vitro inhibition of insulin secretion by diphenylhydantoin. J Clin Invest 49:1942–1948[Medline]
7. Henquin JC 2000 Triggering and amplifying pathways of regulation of insulin secretion by glucose. Diabetes 49:1751–1760[Abstract]
8. Matthews EK, Sakamoto Y 1975 Pancreatic islet cells: electrogenic and electrodiffusional control of membrane potential. J Physiol 246:439–457[Abstract/Free Full Text]
9. Levin SR, Kasson BG, Driessen JF 1978 Adenosine triphosphatases of rat pancreatic islets: comparison with those of rat kidney. J Clin Invest 62 :692–701
10. Herchuelz A, Lebrun P, Sener A, Malaisse WJ 1981 Ionic mechanism of diphenylhydantoin action on glucose-induced insulin release. Eur J Pharmacol 73:189–197[Medline]
11. Pace CS, Livingston E 1979 Ionic basis of phenytoin sodium inhibition of insulin secretion in pancreatic islets. Diabetes 28:1077–1082[Medline]
12. Siegel EG, Janjic D, Wollheim CB 1982 Phenytoin inhibition of insulin release. Studies on the involvement of Ca²⁺ fluxes in rat pancreatic islets. Diabetes 31:265–269[Abstract]
13. Fujimoto S, Ishida H, Kato S, Okamoto Y, Tsuji K, Mizuno N, Ueda S, Mukai E, Seino Y 1998 The novel insulinotropic mechanism of pimobendan: direct enhancement of the exocytotic process of insulin secretory granules by increased Ca²⁺ sensitivity in ß-cells. Endocrinology 139:1133–1140[Abstract/Free Full Text]
14. Fujimoto S, Tsuura Y, Ishida H, Tsuji K, Mukai E, Kajikawa M, Hamamoto Y, Takeda T, Yamada Y, Seino Y 2000 Augmentation of basal insulin release from rat islets by preexposure to a high concentration of glucose. Am J Physiol 279:E927–E940
15. Takehiro M, Fujimoto S, Shimodahira M, Shimono D, Mukai E, Nabe K, Radu RG, Kominato R, Aramaki Y, Seino Y, Yamada Y 2005 Chronic exposure to ß-hydroxybutyrate inhibits glucose-induced insulin release from pancreatic islets by decreasing NADH contents. Am J Physiol 288:E365–E371
16. Kajikawa M, Fujimoto S, Tsuura Y, Mukai E, Takeda T, Hamamoto Y, Takehiro M, Fujita J, Yamada Y, Seino Y 2002 Ouabain suppresses glucose-induced mitochondrial ATP production and insulin release by generating reactive oxygen species in pancreatic islets. Diabetes 51:2522–2529[Abstract/Free Full Text]
17. Shepherd RM, Henquin JC 1995 The role of metabolism, cytoplasmic Ca²⁺, and pH-regulating exchangers in glucose-induced rise of cytoplasmic pH in normal mouse pancreatic islets. J Biol Chem 270:7915–7921[Abstract/Free Full Text]
18. Juntti-Berggren L, Civelek VN, Berggren PO, Schultz V, Corkey BE, Tornheim K 1994 Glucose-stimulated increase in cytoplasmic pH precedes increase in free Ca²⁺ in pancreatic ß-cells. A possible role for pyruvate. J Biol Chem 269:14391–14395[Abstract/Free Full Text]
19. Thomas JA, Buchsbaum RN, Zimniak A, Racker E 1979 Intracellular pH measurements in Ehrlich ascites tumor cells utilizing spectroscopic probes generated in situ. Biochemistry 18:2210–2218[CrossRef][Medline]
20. Ashcroft FM, Ashcroft SJ, Harrison DE 1988 Properties of single potassium channels modulated by glucose in rat pancreatic ß-cells. J Physiol 400:501–527[Abstract/Free Full Text]
21. Radu RG, Fujimoto S, Mukai E, Takehiro M, Shimono D, Nabe K, Shimodahira M, Kominato R, Aramaki Y, Nishi Y, Funakoshi S, Yamada Y, Seino Y 2005 Tacrolimus suppresses glucose-induced insulin release from pancreatic islets by reducing glucokinase activity. Am J Physiol 288:E372–E380
22. Shimono D, Fujimoto S, Mukai E, Takehiro M, Nabe K, Radu RG, Shimodahira M, Kominato R, Aramaki Y, Nishi Y, Funakoshi S, Yamada Y, Seino Y 2005 ATP enhances exocytosis of insulin secretory granules in pancreatic islets under Ca²⁺-depleted condition. Diabetes Res Clin Pract 69:216–223[Medline]
23. Mitchell P 1979 Keilin’s respiratory chain concept and its chemiosmotic consequences. Science 206:1148–1159[Free Full Text]
24. Meglasson MD, Matschinsky FM 1986 Pancreatic islet glucose metabolism and regulation of insulin secretion. Diabetes Metab Rev 2:163–214[Medline]
25. Patterson GH, Knobel SM, Arkhammar P, Thastrup O, Piston DW 2000 Separation of the glucose-stimulated cytoplasmic and mitochondrial NADPH responses in pancreatic islet ß cells. Proc Natl Acad Sci USA 97:5203–5207[Abstract/Free Full Text]
26. Hutson SM 1987 pH regulation of mitochondrial branch chain -keto acid transport and oxidation in rat heart mitochondria. J Biol Chem 262:9629–9635[Abstract/Free Full Text]
27. Sener A, Malaisse WJ 1991 Hexose metabolism in pancreatic islets. Regulation of D-[6-¹⁴C]glucose oxidation by non-nutrient secretagogues. Mol Cell Endocrinol 76:1–6[Medline]
28. Lindström P, Sehlin J 1986 Effect of intracellular alkalinization on pancreatic islet calcium uptake and insulin secretion. Biochem J 239:199–204[Medline]
29. Gunawardana SC, Sharp GW 2002 Intracellular pH plays a critical role in glucose-induced time-dependent potentiation of insulin release in rat islets. Diabetes 51:105–113[Abstract/Free Full Text]
30. Best L, Yates AP, Gordon C, Tomlinson S 1988 Modulation by cytosolic pH of calcium and rubidium fluxes in rat pancreatic islets. Biochem Pharmacol 37:4611–4615[CrossRef][Medline]
31. Juntti-Berggren L, Arkhammar P, Nilsson T, Rorsman P, Berggren PO 1991 Glucose-induced increase in cytoplasmic pH in pancreatic ß-cells is mediated by Na⁺/H⁺ exchange, an effect not dependent on protein kinase C. J Biol Chem 266:23537–23541[Abstract/Free Full Text]
32. Misler S, Barnett DW, Gillis KD, Pressel DM 1992 Electrophysiology of stimulus-secretion coupling in human ß-cells. Diabetes 41:1221–1228[Abstract]
33. Juntti-Berggren L, Rorsman P, Siffert W, Berggren PO 1992 Intracellular pH and the stimulus-secretion coupling in insulin-producing RINm5F cells. Biochem J 287:59–66
34. Pace CS, Goldsmith KT 1986 Effect of substitution of a permeable weak acid for the permissive role of glucose in amino acid-induced electrical activity in B-cells. Endocrinology 119:2433–2438[Abstract]
35. Misler S, Gillis K, Tabcharani J 1989 Modulation of gating of a metabolically regulated, ATP-dependent K⁺ channel by intracellular pH in B cells of the pancreatic islet. J Membr Biol 109:135–143[CrossRef][Medline]
36. Proks P, Takano M, Ashcroft FM 1994 Effects of intracellular pH on ATP-sensitive K⁺ channels in mouse pancreatic ß-cells. J Physiol 475:33–44[Abstract/Free Full Text]
37. Rorsman P 1997 The pancreatic ß-cell as a fuel sensor: an electrophysiologist’s viewpoint. Diabetologia 40:487–495[CrossRef][Medline]
38. Takahashi N, Kadowaki T, Yazaki Y, Ellis-Davies GC, Miyashita Y, Kasai H 1999 Post-priming actions of ATP on Ca²⁺-dependent exocytosis in pancreatic ß cells. Proc Natl Acad Sci USA 96:760–765[Abstract/Free Full Text]
39. Fujimoto S, Mukai E, Hamamoto Y, Takeda T, Takehiro M, Yamada Y, Seino Y 2002 Prior exposure to high glucose augments depolarization-induced insulin release by mitigating the decline of ATP level in rat islets. Endocrinology 143:213–221[Abstract/Free Full Text]
40. Tompkins LS, Nullmeyer KD, Murphy SM, Weber CS, Lynch RM 2002 Regulation of secretory granule pH in insulin-secreting cells. Am J Physiol 283:C429–C437
41. Barg S, Huang P, Eliasson L, Nelson DJ, Obermuller S, Rorsman P, Thevenod F, Renstrom E 2001 Priming of insulin granules for exocytosis by granular Cl^– uptake and acidification. J Cell Sci 114:2145–2154[Abstract/Free Full Text]
42. Maechler P, Wollheim CB 1999 Mitochondrial glutamate acts as a messenger in glucose-induced insulin exocytosis. Nature 402:685–689[CrossRef][Medline]
43. Lynch A, Best L 1990 Cytosolic pH and pancreatic ß-cell function. Biochem Pharmacol 40:411–416[CrossRef][Medline]

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