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Endocrinology Vol. 143, No. 1 213-221
Copyright © 2002 by The Endocrine Society

INSULIN-GLUCAGON-GI PEPTIDES-DIABETES MELLITUS

Prior Exposure to High Glucose Augments Depolarization-Induced Insulin Release by Mitigating the Decline of ATP Level in Rat Islets

Shimpei Fujimoto, Eri Mukai1, Yoshiyuki Hamamoto, Tomomi Takeda, Mihoko Takehiro, Yuichiro Yamada and Yutaka Seino

Department of Metabolism and Clinical Nutrition, Graduate School of Medicine, Kyoto University, Kyoto 606-8507, Japan

Address all correspondence and requests for reprints to: Shimpei Fujimoto, M.D., Ph.D., Department of Metabolism and Clinical Nutrition, Graduate School of Medicine, Kyoto University, 54 Shogoin Kawahara-cho, Sakyo-ku, Kyoto 606-8507, Japan. E-mail: fujimoto{at}metab.kuhp.kyoto-u.ac.jp


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
A brief exposure to elevated glucose augments the insulin secretory response of islets to subsequent stimulation. The site of this priming effect of glucose in the mechanism of the regulation of insulin secretion is not completely known, however. Insulin release triggered by a depolarizing concentration of K+ in the presence of basal glucose is markedly enhanced in primed rat islets. To clarify the role of priming on Ca2+ and ATP efficacy in the exocytotic apparatus, islets were electrically permeabilized to vary the intracellular Ca2+ and ATP concentrations according to the extracellular medium, and insulin release was evaluated. Ca2+ and ATP efficacy in Ca2+- and ATP-dependent insulin secretion was not affected by priming, and alteration of the intracellular Ca2+ concentration after depolarization cannot account for the phenomenon. There was no difference in ATP content before depolarization between nonprimed and primed islets. Moreover, the decline in ATP level after depolarization with basal glucose was observed in both primed and nonprimed islets. However, a reduced decline in ATP level in the early phase was observed in primed islets. In addition, oligomycin, a mitochondrial metabolism inhibitor, abolished the difference in ATP level between primed and nonprimed islets, suggesting that mitochondrial ATP production may be linked to the phenomenon.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GLUCOSE IS THE most important physiological secretagogue of insulin secretion from pancreatic ß-cells, and its mechanism of action is complex.

Glucose by itself triggers insulin secretion from ß-cells by a mechanism that is well documented: glucose entry into the ß-cells accelerates glycolysis and glucose oxidation that generates metabolic signals, including ATP, that close the ATP-sensitive K+ channels (KATP channel). Decrease in K+ conductance due to the closure depolarizes the membrane and opens the voltage-dependent Ca2+ channels. Ca2+ influx then raises the intracellular Ca2+ concentration ([Ca2+]i) to the level that triggers exocytosis of the insulin granule (1, 2). It has been reported that glucose enhances insulin secretion KATP channel-independently. This effect has been confirmed by treatment of ß-cells with diazoxide that prevents the KATP channels from closing and with a depolarizing concentration of extracellular K+ to restore Ca2+ influx (3, 4, 5). As glucose does not increase [Ca2+]i, but, nevertheless, augments insulin release under these conditions, glucose may exert its effects by increasing Ca2+ efficacy in stimulation-secretion coupling, which may be due at least partly to the direct effect of increased ATP derived from glucose metabolism on exocytosis (4, 6). These results indicate that [Ca2+]i, intracellular ATP concentration, and Ca2+ and ATP efficacy in exocytosis all are critical factors in insulin secretion.

It has been demonstrated that previous exposure within 60 min to a stimulatory concentration of glucose augments the insulin secretory response to a subsequent stimulation by glucose or other fuel and nonfuel secretagogues. This enhancing effect on insulin secretion from ß-cells is termed the priming effect, a time-dependent potentiation or "memory effect" of glucose (7, 8, 9, 10, 11, 12). The conditions required for priming are various, including glucose metabolism (9, 11) and possibly phosphoinositide metabolism (13, 14, 15, 16, 17); it is not dependent on cAMP (9, 10), and its dependence on Ca2+ is controversial (9, 11, 18, 19). However, there currently is no information regarding the site of action of the augmentation of depolarization-induced insulin secretion by the priming effect of glucose.

In the present study insulin release, [Ca2+]i, and intracellular ATP level after depolarization, in addition to the Ca2+ and ATP efficacy of the exocytotic system, were compared in primed and nonprimed islets.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Oligomycin, ADP, AMP, adenylate kinase, phosphocreatine, creatine kinase, NAD-dependent glucose-6-phosphate dehydrogenase, 12-O- tetradecanoyl-phorbol-13-acetate (TPA), mannoheptulose, and potassium aspartate were obtained from Sigma (St. Louis, MO). The oligomycin used is a mixture of several oligomycins (A, B, and C). Fura-PE3/AM was obtained from Calbiochem (La Jolla, CA). ATP was purchased from Kohjin (Tokyo, Japan). All other reagents are analytical grade and were obtained from Nacalai Tesque (Kyoto, Japan). Oligomycin was first prepared in dimethylsulfoxide (DMSO) and then was diluted to 1:1000 with buffer. The final concentration of DMSO did not exceed 0.1%, and the same concentration of DMSO was added to controls.

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

Islet isolation and culture
Islets of Langerhans were isolated from Wistar rats by collagenase digestion as described previously (20). Islets were cultured for 12 h in RPMI 1640 medium containing 10% FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 5.5 mM glucose.

Measurement of insulin release from intact islets
Insulin release from intact islets was monitored using static incubation and perifusion conditions described previously with Krebs-Ringer bicarbonate buffer (KRBB) supplemented with 0.2% BSA (20). For static incubation experiments, the islets were preincubated at 37 C for 30 min with 2.8 mM glucose. One group of islets was exposed to 16.7 mM glucose for 30 min (priming); the other group continued to be exposed to 2.8 mM glucose with subsequent incubation with the buffer containing 2.8 mM glucose for 15 min (interval), after which insulin release was stimulated with 30 mM K+ in the presence of 2.8 mM glucose with or without test materials for 30 min (final incubation). At the end of the final incubation period, islets were pelleted by centrifugation (10,000 x g, 180 sec), and aliquots of the buffer were sampled. For perifusion experiments, groups of islets were placed in each of the parallel chambers (400 µl each) of a perifusion apparatus and perifused with KRBB supplemented with 2.8 mM glucose and 0.2% BSA, with 10 mM HEPES adjusted to pH 7.4, at a rate of 0.7 ml/min at 37 C. The medium was continuously gassed with 95% O2 and 5% CO2. Islets usually were perifused for 30 min to establish a stable insulin secretory rate at the basal level of glucose, and then the priming process, interval process, and final incubation were performed. The time intervals at which perifusate samples were collected and detailed protocols are indicated in the figures. The amount of immunoreactive insulin was determined by RIA using rat insulin as standard (20). Experiments using the same protocol were repeated at least three times to ascertain reproducibility.

Measurement of insulin release from permeabilized islets (20)
Cultured islets were preincubated with KRBB containing 2.8 mM glucose and 0.2% BSA for 30 min. The islets were then divided into two groups; one was exposed to 16.7 mM glucose (priming), and the other continued to be incubated with 2.8 mM glucose for an additional 30 min. After both groups of islets were incubated again with 2.8 mM glucose for 15 min (interval), they were washed twice in cold potassium aspartate buffer (KA buffer) containing 140 mM potassium aspartate, 7 mM MgSO4, 2.5 mM EGTA, 30 mM HEPES, and 0.5% BSA (pH 7.0), with CaCl2 added to a Ca2+ concentration of 30 nM. The islets were then permeabilized by high voltage discharge (four exposures, each of 450 µsec duration, to an electrical field of 4.0 kV/cm) in KA 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 KA buffer with various concentrations of Ca2+ and ATP. The Ca2+ concentrations were determined as previously described (21) and were verified using Ca2+ electrode (Horiba, Kyoto, Japan). 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 immunoreactive insulin determination. The amount of immunoreactive insulin was determined as described above. Experiments using the same protocol were repeated three times to ascertain reproducibility.

Measurement of adenine nucleotide and phosphocreatine contents
After groups of cultured intact islets were preincubated with 2.8 mM glucose for 30 min, they were incubated in KRBB supplemented with 0.2% BSA and 10 mM HEPES adjusted to pH 7.4 containing 2.8 or 16.7 mM glucose at 37 C for 30 min (priming). After both of them were exposed to 2.8 mM glucose for 15 min (interval), they were batch-incubated in 1 ml medium containing 30 mM K+ and 2.8 mM glucose. The incubation was stopped by the addition of 0.2 ml trichloroacetic acid to a final concentration of 5% at the intervals indicated in Table 1Go. The tubes were immediately mixed with vortex and then sonicated in ice-cold water for 3 min. They were then centrifuged (2000 x g, 180 sec), and a fraction (0.9 ml) of the supernatant was mixed with 1 ml water-saturated diethyl ether. The ether phase containing trichloroacetic acid was discarded. The step was repeated four times. After the extracts (0.4 ml) were diluted with 0.1 ml 40 mM HEPES solution (final pH 7.4), they were frozen at -80 C until assays. ATP, ADP, and AMP were assayed by a luminometric method (22, 23, 24). For measurement of the sum of ATP + ADP, ADP was converted into ATP by adding 210 µl solution containing 20 mM HEPES (pH 7.75), 3 mM MgCl2, 1.5 mM phosphoenolpyruvate, and 2.2 U/ml pyruvate kinase to 70 µl of the thawed extracts, with incubation at 37 C for 15 min. For measurement of the sum of ATP + ADP + AMP, AMP was converted into ADP before conversion to ATP by adding 210 µl solution containing HEPES (pH 7.75), MgCl2, phosphoenolpyruvate, pyruvate kinase, and 36 U/ml adenylate kinase to 70 µl of the thawed extracts, with incubation at 37 C for 15 min. The ATP concentration in the solutions was measured by adding 200 µl luciferin-luciferase solution (Turner Designs, Sunnyvale, CA) to a fraction of sample (100 µl) in a bioluminometer (luminometer model 20e, Turner Designs, Sunnyvale, CA). The ATP concentration was shown as a signal that was the integrated luminescence strength for 10 sec from 5 sec after the reaction started. For the measurement of ATP, the same procedure was performed, except that the incubation step was performed without pyruvate kinase and adenylate kinase. The ADP concentration and AMP concentration were calculated as the difference between the value of ATP + ADP and that of ATP and between the value of ATP + ADP + AMP and that of ATP + ADP from the same sample, respectively. For measurement of phosphocreatine (25), in a first step the ATP present in the sample was removed by adding 100 µl solution containing 20 mM HEPES (pH 7.75), 3 mM MgCl2, 1.4 mM glucose, 0.6 mM NAD, 0.5 U/ml hexokinase, and 0.7 U/ml NAD-dependent glucose-6-phosphate dehydrogenase to 100 µl of the thawed extracts, with incubation at 37 C for 30 min (26). This was followed by boiling for 1 min to destroy the enzyme. Measurement of the efficacy of this procedure showed that 100% of the ATP was removed. In a second step, phosphocreatine was converted into ATP by adding 100 µl solution containing 20 mM HEPES (pH 7.75), 3 mM MgCl2, 1.25 µM ADP, and 60 U/ml creatine kinase with incubation at 37 C for 30 min. Finally, the ATP formed was measured by the luminometric method described above. To draw a standard curve, blanks, AMP, ADP, ATP, and phosphocreatine standards were run through the entire procedure, including the extraction steps. Experiments using the same protocol were repeated three times to ascertain reproducibility.


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  		Table 1. Time course of changes of ATP content in 16.7 mM glucose-primed and nonprimed intact islets after depolarization by 30 mM K+

 
Measurement of intracellular Ca2+
Cultured islets were loaded with fura-PE3 during 2 h of preincubation in the presence of 2 µM fura-PE3. The islets were then divided into two groups; one was exposed to 16.7 mM glucose (priming), and the other group continued to be incubated with 2.8 mM glucose for another 30 min in KRBB supplemented with 0.2% BSA in humidified air containing 5% CO2 at 37 C. Afterward this group of islets was immediately placed in a heat-controlled chamber on the stage of an inverted microscope kept at 36 ± 1 C, superfused with KRBB supplemented with 2.8 mM glucose and 10 mM HEPES adjusted to pH 7.4 for 15 min (interval period), and subsequently exposed to the medium containing a high concentration of K+. These islets 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). The images were analyzed with a Meta Fluor image analyzing system (Universal Imaging Corp., West Chester, PA). The 340-nm (F340) and 380-nm (F380) fluorescence signals were detected every 20 sec, and the ratios (F340/F380) were calculated. In vitro calibration was performed using fura-PE3, and F340/F380 was converted into calibrated values of [Ca2+]i.

Statistical analysis
Results are expressed as the mean ± SE. Statistical significance was evaluated by unpaired t test. P < 0.05 was considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Time course of glucose-induced or a depolarizing concentration of K+-induced insulin release from high concentration glucose-primed intact islets
One group of islets was exposed to 16.7 mM glucose for 30 min (priming period) while the other continued to be exposed to 2.8 mM glucose. In 16.7 mM glucose-primed islets, the typical biphasic glucose-induced insulin release was observed during the priming period. Both groups of islets were then exposed to 2.8 mM glucose for 15 min (interval period). During that period in the 16.7 mM glucose-primed group, a gradual decrease in insulin secretion for the first 10 min became constant for the last 5 min. However, the insulin secretion during this 5-min period remained significantly greater than that in the control group (at 0 min: 16.7 mM glucose-primed, 0.23 ± 0.02; control, 0.14 ± 0.01 ng/10 islets·min; n = 7; P < 0.01). After the interval period, both groups of islets were simultaneously stimulated with 16.7 mM glucose to determine the biphasic glucose-induced insulin release. The first phase of insulin release was remarkably increased in the 16.7 mM glucose-primed group (insulin secretion within the initial 10 min: control, 4.29 ± 0.01; 16.7 mM glucose-primed, 11.11 ± 0.04 ng/10 islets·10 min; n = 7; P < 0.01). However, during the second phase of insulin release, the insulin release was similar in the two groups 22 min after introduction of 16.7 mM glucose (Fig. 1AGo). The effect of the 16.7 mM glucose-induced priming also was observed when both groups of islets were exposed to a depolarizing concentration (30 mM) of K+ in the presence of 2.8 mM glucose. The 30 mM K+-induced monophasic insulin release also was prominently augmented in 16.7 mM glucose-primed islets. Peak values of insulin secretion and total insulin secretion for the initial 10 min after stimulation in the 16.7 mM glucose-primed group were significantly greater than those in controls [peak value (at 2 min): 16.7 mM glucose-primed, 1.41 ± 0.14; control, 0.56 ± 0.05 ng/10 islets·min; n = 7; P < 0.01; insulin release for initial 10 min: 16.7 mM glucose-primed, 5.68 ± 0.43; control, 2.14 ± 0.09 ng/10 islets·10 min; n = 7; P < 0.01; Fig. 1BGo].



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  		Figure 1. Glucose-induced priming effect on glucose-induced biphasic insulin secretion (A) and on a depolarizing concentration (30 mM) of K+-induced monophasic insulin secretion in the presence of 2.8 mM glucose (B). Two groups of cultured islets were preincubated with 2.8 mM glucose for 30 min (from -75 to -45 min). One group of islets then was exposed to 16.7 mM glucose, and the other group of islets continued to be incubated with 2.8 mM glucose for another 30 min (from -45 to -15 min; priming period). After both groups of islets were incubated with 2.8 mM glucose again for the following 15 min (from -15 to 0 min; interval period), they were stimulated simultaneously with 16.7 mM glucose (A) or 30 mM K+ in the presence of 2.8 mM glucose (B), and the biphasic insulin release (A) or the monophasic insulin release (B) in each group was measured for 30 min (from 0–30 min; final incubation period). Values in A and in B represent the mean ± SE of seven determinations in the same experiment. G, Glucose. A, All values measured, except those at -48, -45, -44, 22, 25, 28, and 30 min, were significantly greater in 16.7 mM glucose-primed islets than the corresponding values in control islets (P < 0.01). B, All indicated values from -5 to 30 min measured, except that at 30 min, were significantly greater with 16.7 mM glucose-primed islets than the corresponding values in control islets (P < 0.01).

 
[Ca2+]i elevation induced by depolarization in the presence of a basal level of glucose in glucose-primed intact islets
Two minutes before depolarization during the 15-min interval, [Ca2+]i in the presence of 5 mM K+ and 2.8 mM glucose in 16.7 mM glucose-primed islets was significantly higher than that in nonprimed islets [average for 2 min before depolarization: primed, 104.5 ± 4.0 (n = 20); nonprimed, 76.2 ± 7.2 nM (n = 21); P < 0.01]. After depolarization induced by 30 mM K+ in the presence of 2.8 mM glucose, [Ca2+]i was slightly higher in primed islets than in nonprimed islets [average for 10 min after depolarization: primed, 171.5 ± 4.1 (n = 20); nonprimed, 155.5 ± 7.2 nM (n = 21); P < 0.05]. However, there was no difference between the increment induced by depolarization in primed islets and that in nonprimed islets (primed, 67.0 ± 5.0 (n = 20); nonprimed, 79.3 ± 8.4 nM; n = 21, not significant; Fig. 2Go).



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  		Figure 2. [Ca2+]i elevation induced by 30 mM K+-induced depolarization in the presence of 2.8 mM glucose in primed and nonprimed islets. Two groups of cultured islets were preincubated with 2.8 mM glucose for 30 min. One group of islets then was exposed to 16.7 mM glucose, and the other group of islets continued to be incubated with 2.8 mM glucose for another 30 min (priming period). After both groups of islets were incubated with 2.8 mM glucose again for the following 15 min (interval period), they were stimulated with 30 mM K+ in the presence of 2.8 mM glucose at time zero. A, Time course of [Ca2+]i. Values represent the mean ± SE of 20 (for primed) or 21 (for nonprimed) determinations from the several experiments. B, Average values calculated from the data from A. Basal, average value from -2 to 0 min in the presence of 5 mM K+ with 2.8 mM glucose. Stimulated, average value from 0–10 min in the presence of 30 mM K+ with 2.8 mM glucose. {triangleup}, Stimulated value minus basal value. *, P < 0.01 vs. basal, nonprimed. {dagger}, P < 0.05 vs. stimulated, nonprimed.

 
Insulin release from electrically permeabilized islets primed with a high concentration of glucose
In the presence of 5 mM ATP, raising the Ca2+ concentration from 30 nM to 3 µM elicited a dose-dependent increase in insulin release from electrically permeabilized islets. However, in the absence of ATP, raising the Ca2+ concentration had no effect on the increase in insulin secretion (Fig. 3AGo). In the presence of 1000 nM Ca2+, raising the ATP concentration from 0 to 5 mM produced a dose-dependent increase in insulin release from permeabilized islets. However, in the presence of 30 nM Ca2+, raising the ATP concentration had no effect on the increase in insulin secretion (Fig. 3BGo).



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  		Figure 3. Dose-dependent effects of Ca2+ and ATP on insulin release from electrically permeabilized islets. After preincubation with 2.8 mM glucose for 75 min, islets were electrically permeabilized and incubated with medium containing Ca2+ and ATP at the concentrations indicated in the figure. A, Ca2+ dose-response curve in the presence of 5 mM ATP and in the absence of ATP. Values represent the mean ± SE of five determinations in the same experiment. B, ATP-dose-response curve in the presence of 1000 and 30 nM Ca2+. Values represent the mean ± SE of eight determinations in the same experiment.

 
In the absence of ATP, insulin release was greater in primed islets than in nonprimed islets, independent of the Ca2+ concentration; however, no increment in insulin release induced by the elevation of Ca2+ was observed in either primed or nonprimed islets [at 30 nM Ca2+: nonprimed, 0.24 ± 0.04; primed, 0.42 ± 0.03 (n = 5; P < 0.01); at 100 nM: nonprimed, 0.23 ± 0.02; primed, 0.40 ± 0.03 (n = 5; P < 0.01); at 1000 nM: nonprimed, 0.25 ± 0.03; primed, 0.42 ± 0.03 ng/islet·30 min (n = 5; P < 0.01)]. In the presence of 5 mM ATP, insulin release at 30, 100, and 1000 nM Ca2+ was greater in 16.7 mM glucose-primed islets than in nonprimed islets, and the Ca2+-dependent increase in insulin secretion was observed in both primed and nonprimed islets (Fig. 4AGo). In the presence of 5 mM ATP, when the increment in insulin release in the presence of 30 nM Ca2+ due to the priming effect was subtracted from the value of insulin release from primed islets, there was no difference in the values of insulin secretion at 100 and 1000 nM compared with the values from nonprimed islets (Fig. 4AGo). In the presence of 1000 nM Ca2+, insulin release at 0, 1, 3, and 5 mM ATP was greater in primed islets than in nonprimed islets, and the ATP-dose-dependent increase in insulin secretion was observed in both primed and nonprimed islets (Fig. 4BGo). In the presence of 1000 nM Ca2+, however, when the increment in insulin release in the absence of ATP due to the priming effect was subtracted from the value of insulin release from primed islets, no significant augmentation was observed in the values of insulin secretion at 1, 3, and 5 mM ATP compared with the values from nonprimed islets (Fig. 4BGo).



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  		Figure 4. Effect of 16.7 mM glucose-induced priming on insulin release from electrically permeabilized islets in the presence of 5 mM ATP and various concentrations of Ca2+ (A) and in the presence of 1000 nM Ca2+ and various concentrations of ATP (B). Two groups of cultured islets were preincubated with 2.8 mM glucose for 30 min. One group of islets then was exposed to 16.7 mM glucose, and the other group of islets was incubated with 2.8 mM glucose for another 30 min (priming period). After both groups of islets were incubated with 2.8 mM glucose again for 15 min (interval period), they were electrically permeabilized and incubated with medium containing ATP and Ca2+ at the concentrations indicated in the figure. The values obtained by subtracting the increment in insulin release in the presence of 30 nM Ca2+ due to the priming effect from insulin release from primed islets and the values obtained by subtracting the increment in insulin release in the absence of ATP due to the priming effect from insulin release from primed islets are also indicated in A and B, respectively. All values represent the mean ± SE of 20 (A) and 9 (B) determinations in the same experiment. *, P < 0.01 vs. corresponding nonprimed control.

 
Time course of ATP level changes after depolarization in the presence of basal level of glucose in glucose-primed islets
Insulin content and DNA content at the end of the interval period (time zero) was the same in nonprimed and primed islets (data not shown). ATP content at time zero was the same in nonprimed and primed islets. Twenty minutes after 30 mM K+-induced depolarization, the ATP content was the same in primed and nonprimed islets, but it was significantly decreased in both primed and nonprimed islets compared with that in islets that continued to be incubated in the presence of 5 mM K+. On the other hand, ATP content 2 and 4 min after depolarization was significantly greater in primed islets than that in nonprimed islets (Table 1AGo).

Adenine nucleotide and phosphocreatine content in glucose-primed islets
ATP, ADP, AMP, and phosphocreatine contents at the end of the interval period (before depolarization) were similar in primed and nonprimed islets (Table 2Go).


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  		Table 2. Adenine nucleotide and phosphocreatine contents in primed and nonprimed islets in the presence of 2.8 mM glucose and 5 mM K+

 
Effects of oligomycin on ATP level and on insulin release after depolarization in glucose-primed islets
To investigate a role of mitochondrial metabolism in the reduced decline in ATP and the augmented insulin release in primed islets, 2 µg/ml oligomycin were used. Two minutes after K+-induced depolarization in the presence of 2.8 mM glucose, the ATP level was significantly greater in 16.7 mM glucose-primed islets. Addition of 2 µg/ml oligomycin during depolarization significantly reduced the ATP level in both primed and nonprimed islets; however, no significant difference in ATP level between nonprimed and primed islets was observed (Table 3Go). Basal insulin release and a depolarizing concentration of K+-induced insulin release from primed islets were significantly increased compared with those from nonprimed islets [basal, nonprimed, 0.25 ± 0.05; basal, primed, 0.43 ± 0.04 (n = 7; P < 0.01); stimulated, nonprimed, 0.76 ± 0.07; stimulated, primed, 1.24 ± 0.09 ng/islet·30 min (n = 7; P < 0.01)]. Addition of 2 µg/ml oligomycin during depolarization completely suppressed stimulated insulin release to the basal level in both nonprimed and primed islets (Fig. 5Go).


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  		Table 3. Effects of 2 µg/ml oligomycin on ATP content in primed and nonprimed islets after 2-min exposure to a depolarizing concentration of K+ in the presence of 2.8 mM glucose

 


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  		Figure 5. Effects of 2 µg/ml oligomycin on depolarization-induced insulin release from primed and nonprimed intact islets. Two groups of cultured intact islets were preincubated with 2.8 mM glucose for 30 min. One group of islets then was exposed to 16.7 mM glucose (priming), and the other group of islets continued to be incubated with 2.8 mM glucose for 30 min. After both groups of islets were incubated with 2.8 mM glucose (with 5 mM K+) again for the following 15 min (interval), they were incubated with 30 or 5 mM K+ in the presence of 2.8 mM glucose for 30 min (final incubation). Oligomycin was present only during the final incubation. Values represent the mean ± SE of seven determinations in the same experiment. *, P < 0.01 vs. corresponding nondepolarized without oligomycin. {dagger}, P < 0.01 vs. corresponding depolarized without oligomycin. {ddagger}, P < 0.01 vs. nondepolarized, nonprimed without oligomycin.

 
Depolarization-induced insulin release and time course of ATP level changes in mannoheptulose-preexposed islets
One group of islets was exposed to 16.7 mM glucose with 20 mM mannoheptulose for 30 min, and the other was exposed to 2.8 mM glucose with 20 mM mannoheptulose during the priming period. Basal insulin release in the presence of 2.8 mM glucose and 5 mM K+ and depolarization-induced insulin release in the presence of 2.8 mM glucose and 30 mM K+ were not significantly augmented from those in 16.7 mM glucose- and 20 mM mannoheptulose-preexposed intact islets compared with those from 2.8 mM glucose and 20 mM mannoheptulose-preexposed islets [basal release: 2.8 mM glucose-preexposed, 0.23 ± 0.04; 16.7 mM glucose-preexposed, 0.26 ± 0.05 (n = 6; not significant); stimulated release: 2.8 mM glucose-preexposed, 0.60 ± 0.06; 16.7 mM glucose-preexposed, 0.63 ± 0.08 ng/islet·30 min (n = 6; not significant)]. ATP content at 0, 2, and 8 min after depolarization was the same in 16.7 mM glucose- and 20 mM mannoheptulose-preexposed islets and 2.8 mM glucose- and 20 mM mannoheptulose-preexposed islets (Table 1BGo).

Depolarization-induced insulin release and time course of ATP level changes in TPA-primed islets
One group of islets was exposed to 2.8 mM glucose with 10 nM TPA for 30 min while the other continued to be exposed to 2.8 mM glucose during the priming period. Depolarization-induced insulin release was significantly augmented from TPA-primed intact islets compared with that from nonprimed islets (Table 4AGo). In the presence of 5 mM ATP, insulin release at 30, 100, and 1000 nM Ca2+ was greater in 10 nM TPA-primed permeabilized islets than in nonprimed islets, and the Ca2+-dependent increase in insulin secretion was observed in both primed and nonprimed permeabilized islets (Fig. 6AGo). In the presence of 30 nM Ca2+ and the absence of ATP, the increment in insulin release due to the priming effect was not observed (Fig. 6AGo). In the presence of 5 mM ATP, when the increment in insulin release in the presence of 30 nM Ca2+ due to the priming effect was subtracted from the insulin release from primed permeabilized islets, there was a significant increase in insulin secretion at 100 and 1000 nM compared with that from nonprimed islets (Fig. 6AGo). In the presence of 1000 nM Ca2+, insulin release at 1, 3, and 5 mM ATP was greater in primed permeabilized islets than in nonprimed islets, and the ATP dose-dependent increase in insulin secretion was observed in both primed and nonprimed permeabilized islets (Fig. 6BGo). In the presence of 1000 nM Ca2+ and the absence of ATP, however, no increment in insulin release due to the priming effect was observed (Fig. 6BGo). ATP content at 0, 2, and 8 min after depolarization was the same in nonprimed and TPA-primed islets (Table 4BGo).


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  		Table 4. TPA (10 nM)-induced priming effect on insulin release from intact islets (A) and time course of changes of ATP content in 10 nM TPA-primed islets after depolarization by 30 mM K+ (B)

 


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  		Figure 6. Effects of 10 nM TPA-induced priming on insulin release from electrically permeabilized islets in the presence of 5 mM ATP and various concentrations of Ca2+ (A) and in the presence of 1000 nM Ca2+ and various concentrations of ATP (B). Two groups of cultured islets were preincubated with 2.8 mM glucose for 30 min. One group of islets then was exposed to 10 nM TPA in the presence of 2.8 mM glucose, and the other group of islets continued to be incubated with 2.8 mM glucose without TPA for 30 min (priming period). After both groups of islets were incubated with 2.8 mM glucose again for the following 15 min (interval period), they were electrically permeabilized and incubated in medium containing ATP and Ca2+ at the concentrations indicated. The values obtained by subtracting the increment in insulin release in the presence of 30 nM Ca2+ and 5 mM ATP due to the priming effect from insulin release from primed islets and insulin release in the presence of 30 nM Ca2+ and the absence of ATP also are indicated in A. All values represent the mean ± SE of eight determinations in the same experiment. A and B are different experiments. *, P < 0.01 vs. corresponding nonprimed control. {dagger}, P < 0.01 vs. no ATP, primed. {ddagger}, P < 0.01 vs. corresponding nonprimed control.

 

   Discussion
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 Abstract
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 Materials and Methods
 Results
 Discussion
References
 
We have shown that the difference in ATP level in the early phase after depolarization between primed and nonprimed islets may contribute to the difference in depolarization-induced insulin secretion.

In the perifusion experiments the first phase of glucose-induced insulin release from primed islets was remarkably augmented; during the second phase the insulin secretion from nonprimed and primed islets was similar. Aizawa et al. (5, 27) proposed that glucose-induced biphasic insulin secretion is the sum of glucose-induced triggering of insulin release produced solely by cytosolic Ca2+ elevation (first phase) and self-augmentation of insulin release by the glucose-induced priming effect (second phase). Therefore, as a secretagogue to quantify the priming effect during final incubation, a depolarizing concentration of K+ in the presence of the basal level of glucose was chosen to avoid self-augmentation during the final incubation. The monophasic insulin release triggered by a depolarizing concentration of K+ in the presence of basal levels of glucose also was markedly enhanced in primed islets. Accordingly, [Ca2+]i and intracellular ATP in this condition in addition to Ca2+ and ATP efficacy in the exocytotic system were evaluated in both nonprimed and primed islets to identify the mechanism involved in stimulation-secretion coupling.

To quantify the priming effect on Ca2+ and ATP efficacy in the exocytotic process of insulin secretory granules directly, high concentration glucose-primed islets were electrically permeabilized to manipulate the intracellular Ca2+ and ATP concentrations according to the extracellular medium, and insulin release was examined. Both the Ca2+ dose-response curve in the presence of 5 mM ATP and the ATP dose-response curve in the presence of 1 µM Ca2+ were shifted to the left by priming. However, the shift due to priming also was observed in the Ca2+ dose-response curve in the absence of ATP in which no increment in insulin release induced by Ca2+ elevation was observed. In a recent study (24) we found that prior exposure to high glucose enhanced basal insulin release from rat pancreatic islets and suggested that this enhancement was due to ATP-independent and Ca2+-independent insulin release, because it was not affected by reduction of the intracellular ATP or Ca2+ concentration. The GTP-sensitive site may play a role, as no augmentation was observed in GTP-depleted islets, and GDP analog significantly suppressed the augmentation in ATP- and Ca2+-depleted permeabilized islets. Accordingly, the leftward shift of the Ca2+ and ATP dose-response curves due to priming should be derived from the augmented ATP-independent and Ca2+-independent insulin release. When Ca2+-independent and ATP-independent insulin release enhanced by priming was subtracted from these dose-response curves in primed islets, there was no difference between nonprimed and primed islets. These results indicate that Ca2+ and ATP efficacy in Ca2+- and ATP-dependent insulin secretion is not affected by priming. It should be emphasized that no Ca2+-triggered insulin secretion was observed even from primed islets without ATP, contrary to the report that the priming process increased Ca2+-triggered exocytosis in the absence of ATP in permeabilized adrenal chromaffin cells (28) and PC12 cells (29). Our results are consistent with the report of capacitance measurements showing that the pool of insulin secretory granules that can be released by Ca2+ triggering in the absence of ATP is considerably smaller in pancreatic ß-cells compared with other neuroendocrine cells (30, 31). Those results also suggest that the Ca2+ dependency and the ATP dependency of exocytosis in the pancreatic ß-cell are tightly linked.

Because the observed differences in [Ca2+]i elevation after depolarization and in Ca2+ and ATP efficacy in the exocytotic system cannot account for the enhanced depolarization-induced insulin release due to preexposure to a high concentration of glucose, the ATP level after 30 mM K+-induced depolarization in the presence of basal glucose was examined. The ATP content before depolarization was the same in nonprimed and primed islets, and the decline in ATP level after depolarization in the presence of a basal level of glucose, which reflects a larger consumption than production of ATP produced by increased [Ca2+]i (32), was observed in both primed and nonprimed islets. However, a reduced decline in ATP in the early phase was observed in primed islets. Even in the presence of a low concentration of ATP that cannot close the KATP channel and cannot trigger Ca2+ influx by itself, insulin release occurs when Ca2+ influx is produced by membrane depolarization (6, 33). Moreover, the ATP dose-response curve in the presence of a clamped sufficient concentration of Ca2+ clearly shows that the increment in ATP level increases Ca2+ efficacy in exocytosis. In addition, in mannoheptulose-preexposed islets, in which no augmentation of depolarization-induced insulin release due to a glucose-induced priming effect was observed, the decline in ATP level in the early phase was the same in high glucose-preexposed islets and control islets. The difference in ATP level in the early phase, therefore, may affect depolarization-induced monophasic insulin release by a direct effect on the exocytotic apparatus.

We then investigated the mitigated decline of ATP level in primed islets. Because pancreatic ß-cells contain creatine kinase (34, 35) and adenylate kinase (36), intracellular phosphocreatine and ADP might function as an ATP reservoir in islets by the following reaction: phosphocreatine + ADP -> creatine + ATP by creatine kinase, and 2ADP -> AMP + ATP by adenylate kinase. If such an ATP reservoir pool were increased by preexposure to high glucose and its level maintained before depolarization, it would prevent the intracellular ATP level from decreasing even when ATP consumption is increased by depolarization. The fact that ADP, AMP, and phosphocreatine contents before depolarization in islets were the same in nonprimed and primed islets suggests that the reduced decline in ATP level in primed islets is not derived from an increase in ADP or the phosphocreatine pool. A rise in [Ca2+]i stimulates mitochondrial metabolism by activating enzymes, including pyruvate dehydrogenase, NAD+-isocitrate dehydrogenase, and oxoglutarate dehydrogenase in the Krebs cycle (37) and mitochondrial glycerol phosphate dehydrogenase in the glycerol phosphate shuttle (38, 39, 40). To determine whether mitochondrial ATP production plays a role in the reduced decline in ATP level and augmented insulin release in primed islets, 2 µg/ml oligomycin, a mitochondrial H+-adenosine triphosphatase inhibitor, at which concentration the elevation of [Ca2+]i after depolarization is not affected (41), was used during high concentration K+-induced depolarization. The ATP level 2 min after depolarization in the presence of basal glucose was suppressed by the reagent in both primed and nonprimed islets, which indicates that mitochondrial ATP production may participate in regulation of the ATP level after depolarization even in nonprimed islets. The facts that the difference in ATP level between primed and nonprimed islets in the early phase was abolished in the presence of oligomycin and that high K+-induced insulin release was completely suppressed to the basal level in both primed and nonprimed islets in the presence of the reagent suggest that mitochondrial ATP production may play a role in both the reduced decline in ATP level and the enhanced insulin release in primed islets.

As the involvement of PLC and PKC activation has been reported in the glucose-induced priming effect (13, 14, 15, 16, 17), the intracellular ATP level, in addition to Ca2+ and ATP efficacy in the exocytotic system in islets pretreated by TPA, a PKC activator, was examined. Depolarization-induced insulin release from islets primed by TPA at a basal level of glucose was significantly augmented compared with that from nonprimed islets, as previously reported (16). Interestingly, although ATP decline after depolarization was the same in TPA-primed and nonprimed islets, increased Ca2+ and ATP efficacy in insulin release was observed in TPA-primed islets. These results indicate at the least that a different mechanism is involved in the glucose-induced priming effect and the priming effect induced by the PKC activator and that multiple priming mechanisms are functional within the islet. Whether ATP and phorbol ester are common activators of the same downstream signaling molecule ultimately required for priming remains unknown.


   Acknowledgments
 
The authors thank Mr. S. Nawata for his technical assistance.


   Footnotes
 
This work was supported in part by Grants-in-Aid for Scientific Research from the Ministry of Education, Science, Sports, and Culture of Japan; a Grant-in-Aid for Creative Basic Research (NP10NP0201) from the Ministry of Education, Science, Sports, and Culture of Japan; and a grant from the Research for the Future Program of the Japan Society for the Promotion of Science (JSPS-RFTF97I00201).

1 Research fellow of the Japan Society for Promotion of Science. Back

Abbreviations: [Ca2+]i, Intracellular Ca2+ concentration; DMSO, dimethylsulfoxide; KA buffer, potassium aspartate buffer; KATP channel, ATP-sensitive K+ channel; KRBB, Krebs-Ringer bicarbonate buffer; TPA, 12-O-tetradecanoyl-phorbol-13-acetate.

Received May 16, 2001.

Accepted for publication September 12, 2001.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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