1Unit of Endocrinology and Metabolism and 2Service of Pathology, University of Louvain Faculty of Medicine, B-1200 Brussels, Belgium
Submitted 22 September 2003 ; accepted in final form 16 April 2004
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ABSTRACT |
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pancreatic -cell; cytosolic calcium concentration; mitochondrial activity; glucose toxicity
Previous studies have shown that prolonged in vivo or in vitro exposure to high glucose concentrations causes parallel decreases in maximal GSIS and -cell insulin content and abolishes GSIS within the physiological range of glucose concentrations (510 mmol/l) (32, 37, 52). However, the cause of this loss of GSIS remains controversial. In one group of studies (5, 7, 8, 13, 23, 39, 51),
-cells exposed to high glucose concentrations became insensitive to subsequent acute glucose stimulation, i.e., for changes in ATP production, intracellular Ca2+ concentration ([Ca2+]i), and insulin secretion, defects that were tentatively attributed to an increased expression of the mitochondrial uncoupling protein (UCP)2 (34, 47). In sharp contrast, in another group of studies (1, 38, 4042, 48, 60),
-cells exposed to high glucose concentrations became hypersensitive to subsequent acute glucose stimulation, i.e., for the acceleration of mitochondrial metabolism, proinsulin biosynthesis, and insulin secretion. In association with
-cell degranulation, this increased sensitivity to glucose resulted in an already stimulated secretion at low fasting glucose concentrations (
5 mmol/l) and a lack of further stimulation of insulin secretion by physiological postprandial glucose concentrations (510 mmol/l).
These two apparently irreconcilable explanations for the absence of GSIS after prolonged exposure to high glucose concentrations have distinct and even opposite implications for the development of new therapeutic strategies to improve GSIS in type 2 diabetes. Therefore, to clarify this issue, we reinvestigated the effects of prolonged exposure to high glucose concentrations on key -cell stimulus-secretion coupling events in a single experimental model, the in vitro cultured rat islet, with a particular focus on changes in islet energetic metabolism and both triggering and amplifying pathways of GSIS (27, 59).
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MATERIALS AND METHODS |
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Islet isolation and culture. Islet isolation was performed using sterile-filtered Krebs solution containing 10 mmol/l glucose, 5 mmol/l HEPES, 100 IU/ml penicillin, 100 µg/ml streptomycin, and 0.3 µg/ml fungizone. Briefly, male Wistar rats (180200 g) were anesthetized by intraperitoneal injection of 25 mg/kg pentobarbital (Nembutal, Abbott Laboratories) and laparotomized. After the animals were killed by incision of the heart, the pancreas was inflated by injecting 5 ml of medium containing 1.5 mg/ml of collagenase (Serva, Heidelberg, Germany) through the common pancreatic duct. After collagenase digestion, the islets were separated from the digest by density gradient centrifugation using Histopaque 1077 (Sigma). The islets were washed thrice and hand-picked under a stereomicroscope to ensure high purity of the preparation. They were then separated into two batches and cultured at 37°C in RPMI 1640 medium containing 10 (Control) or 30 (High-glucose) mmol/l glucose and 5 g/l bovine serum albumin instead of 10% fetal calf serum, to prevent proliferation of fibroblasts and attachment and flattening of islets on the bottom of the dish. Unless otherwise specified, the islets were cultured for 78 days, and the medium was renewed after 1 day, and then every other day. Islets with a diameter >150200 µm, which usually develop central necrosis during culture, were discarded. High-glucose and Control islets of similar size were then chosen for functional comparison. Although mycoplasmal contamination was not specifically tested, culture contamination was routinely excluded by phase contrast microscopy (x2040 objective).
All animal experimentations performed for this study were conducted in accord with accepted standards of humane animal care and were approved by the Institutional Committee on Animal Experimentation from the Faculty of Medicine of the University of Louvain.
Optical and electron-microscopic analysis of islets. After 1 wk of culture in G10 or G30, the islets were washed in RPMI without BSA, fixed for 4 h in Bouin's fluid or in 2.5% glutaraldehyde solution for light and electron microscopy, respectively. For light-microscopic analysis, 4-µm-thick sections of paraffin-embedded tissue were mounted on glass slides and processed routinely for hematoxylin and eosin staining. Structures containing fragments of basophilic chromatin material were considered characteristic of "apoptotic bodies." For electron-microscopic analysis, ultrathin sections (80 nm) of epoxy-embedded islets were counterstained with aqueous uranyl acetate and lead citrate. They were then examined on a Zeiss 109 transmission electron microscope (Carl Zeiss, New York City, NY).
Insulin secretion and insulin content measurements.
After culture, batches of 2030 islets were perifused (1 ml/min) with a medium containing Gx. Insulin was measured in 2-min effluent collections by radioimmunoassay with rat insulin as a standard. After perifusion, the islets were disrupted by sonication in TNE buffer (10 mmol/l Tris, 0.2 M NaCl, and 10 mmol/l EDTA) for measurement of their insulin (RIA) and DNA contents (fluorescent assay using bisbenzimide) (35).
[Ca2+]i measurements.
[Ca2+]i was measured by microspectrofluorimetry with Quanticell 700m (Visitech, Sunderland, UK). Islets were loaded for 23 h at 37°C with 2 µmol/l fura PE 3-acetoxymethylester (fura PE 3-AM; Teflabs, Austin, TX) in RPMI medium containing the same glucose concentration as that used during culture. They were then transferred into a thermostatized (37°C) chamber mounted on the stage of an inverted microscope used in the epifluorescence mode with a x20 objective and were perifused at a flow rate of 1 ml/min. The islets were alternately excited at 340 and 380 nm by a monochromator. A dichroic mirror centered at 430 nm reflected the light of excitation to the perifusion chamber and transmitted the emitted fluorescence to a charge-coupled device video camera (Photonic Science, Tunbridge Wells, UK) through a 510-nm filter. The data were automatically corrected for background fluorescence. [Ca2+]i was calculated by comparing the ratio of fluorescence at each pixel to a calibration curve based on the equation of Grynkiewicz et al. (24). The mean [Ca2+]i in the islet was then calculated by averaging the [Ca2+]i of all pixels of the islet.
45Ca2+ efflux measurements. After 1 wk of culture in RPMI medium containing 10 or 30 mmol/l glucose, the islets were loaded overnight with 45CaCl2 (80 µCi/ml; Amersham Pharmacia Biotech, Buckinghamshire, UK) in the same medium. After several washes to remove the extracellular tracer, batches of 30 islets were perifused with a medium containing various glucose concentrations. The radioactivity lost by the islets was measured in 2-min effluent collections by liquid scintillation using Pico-fluor 15 (Packard Bioscience, Meriden, CT). At the end of the experiment, the islets were collected to measure their total 45Ca2+ radioactivity. The fractional rate of 45Ca2+ efflux was calculated by dividing the radioactivity lost in each sample by the total sum of radioactivity present in the islets just before collection of that sample (15).
Islet 45Ca2+ uptake measurements. After 1 wk of culture, Control and High-glucose islets were divided into three batches of 30 each. For each type of islets, one batch was used for duplicate DNA content measurement, and the other two groups were loaded for 18 h with 45CaCl2 in the same medium, with eventual addition of 1 µmol/l thapsigargin (Tg) for the last hour of culture. The islets were then rinsed thrice in modified Krebs solution without bicarbonate and containing 5 mmol/l Tris, pH 7.4, supplemented with 2 mmol/l LaCl3, to block Ca2+ fluxes (25). Each group of islets was then divided into batches of 8 for triplicate determination of their 45Ca2+ content. After normalization for variations in the total radioactivity concentration of the loading medium and for differences in islet DNA content, the results were expressed as a percentage of the 45Ca2+ content of Control islets within the same experiment.
Reduced pyridine nucleotide fluorescence measurements. After culture, the islets were transferred to the same setup as for [Ca2+]i measurements. The reduced forms of NAD and NADP, collectively referred to as NAD(P)H, were excited at 360 nm, and the fluorescence emitted through a dichroic mirror centered at 400 nm was filtered at 470 nm (22). After background subtraction, the results were normalized to the maximal fluorescence level measured 25 min after addition of 5 mmol/l azide in the presence of 20 mmol/l glucose.
Mitochondrial membrane potential measurements. The islets were loaded for 20 min with 10 µg/ml rhodamine 123 (Molecular Probes, Eugene, OR) in the presence of the same glucose concentration as that during culture, and changes in the fluorescence of rhodamine 123 were measured by spectrofluorimetry (excitation/emission 490/530 nm, 505 nm long pass dichroic mirror) (55). After background subtraction, the rhodamine 123 fluorescence in each islet was normalized to the maximal fluorescence level measured 25 min after addition of 5 mmol/l azide in the presence of 20 mmol/l glucose. A decrease in rhodamine fluorescence reflects mitochondrial membrane hyperpolarization.
Measurements of islet adenine nucleotides. After culture, batches of 58 islets were preincubated for 30 min in 500 µl of medium containing 0.5 mmol/l glucose. They were then incubated for 30 min in 1 ml of medium containing 0.520 mmol/l glucose. At the end of the experiment, the islets were disrupted with trichloracetic acid (5% vol/vol), and their ATP and ADP contents (pmol/islet) were measured by a luminometric method (17). The fraction of the sum (ATP + ADP) that was in the form of ATP in the islets was calculated [ATP/(ATP + ADP)] and used as an indicator of islet cell energetic level (48).
Measurements of islet mRNA expression of mitochondrial UCP2.
The UCP2/cyclophilin mRNA ratio was determined by reverse transcription and duplex radioactive semiquantitative PCR in Control and High-glucose islets, as previously described (31). Primers for cyclophilin (31) and UCP2 (sense 5'-ggcctctacgactctgtaaagc-3'; antisense 5'-TGGCATTTCGGGCAACATTG-3') were used at a concentration of 200 nmol/l. The thermal cycle profile was a 10-min denaturing step at 94°C followed by cycles of amplification (1 min of denaturation at 94°C, 1 min of annealing at 60°C, 1 min of extension at 72°C) and a final extension step of 10 min at 72°C. PCR were stopped in the middle of the exponential phase of amplification of both genes (21 cycles) (31). The amplimers were separated on a 6% PAGE, and their [-32P]dCTP content was quantitated by a Cyclone Storage Phosphor System (Packard, Meriden, CT). The UCP2-to-cyclophilin mRNA ratio was calculated for each sample and normalized to the ratio measured in islets cultured in G10.
Data analysis.
High-glucose and Control islets from the same preparation were always tested simultaneously within a single experiment, with an equal number of islets (2) of each type in each experiment. The results are means ± SE for the indicated number of islets from at least three different preparations. Statistical significance of differences between means was assessed by one-way ANOVA, followed by a Newman-Keuls test or an unpaired t-test, unless specified otherwise. Differences were considered significant at P < 0.05.
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RESULTS AND DISCUSSION |
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Morphological and functional characteristics of Control islets.
Optical and electron microscopy of Control islets showed well-preserved islet architecture, no morphological signs of -cell apoptosis, and normal ultrastructure of
- and non-
-cells with uncondensed and unfragmented nuclei, well-defined endoplasmic reticulum (ER), Golgi complex, and mature secretory granules (Fig. 1, A, C, and E). In these islets, a stepwise increase in glucose concentration from G0.5 to G20 stimulated key GSIS coupling events as expected (27, 46, 65): a glucose concentration-dependent increase in islet NAD(P)H autofluorescence, mitochondrial membrane hyperpolarization, and rise in the ATP/(ATP+ADP) ratio; a transient decrease in [Ca2+]i, reflecting increased Ca2+ pumping by the ER (61) without changes in insulin secretion at subthreshold glucose concentrations (G3G6), and a parallel increase in [Ca2+]i and insulin secretion above that threshold (Fig. 2). Although [Ca2+]i did not reach steady state within the short duration of each glucose step, we can estimate that the glucose sensitivity of [Ca2+]i and insulin secretion changes in Control islets was similar to that previously reported in freshly isolated rat islets (2), with half-maximal and maximal effective glucose concentrations close to G10 and G20, respectively. These glucose-induced [Ca2+]i changes were similar after 121 days of culture in G10 (data not shown).
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Insulin and DNA content of Control and High-glucose islets.
On average, the insulin content was 50% lower in High-glucose than in Control islets (23.5 ± 2.2 and 48.4 ± 4.7 ng/islet, respectively, n = 23, P < 0.0001), whereas the DNA content was similar in the two types of islets (87.8 ± 9.5 and 87.5 ± 9.5 ng/islet, n = 23).
High-glucose islets have a high energetic state at low glucose despite increased UCP2 mRNA levels. As in Control islets, a stepwise increase in glucose concentration from G0.5 to G20 induced concentration-dependent increases in NAD(P)H autofluorescence, mitochondrial membrane hyperpolarization, and ATP/(ATP+ADP) ratio in High-glucose islets (Fig. 2, A-C). However, these effects of glucose were of much smaller amplitude than those in Control islets. To permit direct comparison of the semiquantitative changes in NAD(P)H and rhodamine 123 fluorescence between the two types of islets, the results were normalized to the maximal fluorescence level observed 25 min after addition of azide to G20. At the concentration of 5 mmol/l, this inhibitor of complex IV of the electron transport chain depolarized the mitochondrial membrane (maximal level of rhodamine 123 fluorescence), transiently increased NAD(P)H autofluorescence, and reduced the ATP/(ATP+ADP) ratio to the level measured in Control islets incubated in G0.5 (Fig. 2C). The NAD(P)H autofluorescence, mitochondrial membrane hyperpolarization, and ATP/(ATP+ADP) ratio measured in the presence of G0.5 were all significantly higher in High-glucose than in Control islets (P < 0.05 or less), whereas the maximal and minimal energetic levels, measured in the presence of G20 and G20+azide, respectively, were similar in the two types of islets. These results are not compatible with the hypothesis that prolonged exposure to high glucose concentrations abolishes the glucose-induced rise in ATP production by increasing the expression of the mitochondrial UCP2 (47). We therefore compared the expression of UCP2 in High-glucose and Control islets by RT-PCR, with cyclophilin as an internal control gene. After 12 wk of culture, the UCP2/cyclophilin mRNA ratio was significantly higher in High-glucose than in Control islets (174 ± 12 and 100 ± 9%, respectively, P < 0.001).
These results therefore indicate that the lower glucose responsiveness of mitochondrial metabolism in High-glucose islets results from an increase in mitochondrial metabolism and ATP production in G0.5, rather than from its reduction in G20, despite an 1.7-fold increase in islet UCP2 mRNA expression. The functional significance of the latter increase is not clear (36). Although we agree that a large overexpression of UCP2 can be detrimental to GSIS in any type of islets (12), our results suggest that, as in other cell types, a moderate (<2-fold) increase in UCP2 expression in rat
-cells does not impair ATP production (53). It is even possible that a limitation in the production of reactive oxygen species at complexes I and III by mild mitochondrial uncoupling explains the surprising observation that mild UCP2 overexpression rather improves
-cell dysfunction from Zucker Diabetic Fatty rats (63).
Increased glucose sensitivity of insulin secretion in the presence of a stable elevation of [Ca2+]i in High-glucose islets.
Compared with Control islets, [Ca2+]i was 50 nmol/l higher in High-glucose islets perifused with G0.5 and did not further increase upon glucose stimulation, except for a slow regular increase that varied between experiments and likely resulted from the progressive leakage of fura PE 3 from the islets (compare Figs. 2D and 4C) (3, 22). These [Ca2+]i alterations developed progressively during culture in G30 (normal response after 1 day, partial alteration after 3 days, and complete lack of response after 721 days). They were fully reversible upon further culture in G10 (no correction after 1 day, partial and complete normalization after 3 and 7 days, respectively; data not shown, n = 46 islets from 23 independent cultures). Despite the sustained and glucose-insensitive elevation of [Ca2+]i, High-glucose islets displayed a clear GSIS that was rapidly inhibited by addition of 5 mmol/l azide. Compared with Control islets, insulin secretion from High-glucose islets was similar in G0.5, higher in G6 (P < 0.05), not different in G10, and lower in G20 (P < 0.001), with nearly maximal stimulation in G10 (Fig. 2E). After correction for the decrease in islet insulin content, insulin secretion was much larger in High-glucose than in Control islets during perifusion with G6-G10, but it was not different in G20 (Fig. 2F).
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Increased glucose sensitivity of the amplifying pathway of GSIS in High-glucose islets. Besides its ability to increase Ca2+ influx (triggering pathway of GSIS), glucose also amplifies the efficacy of Ca2+ on insulin granule exocytosis by a mechanism that is independent from the closure of ATP-sensitive K+ (KATP) channels (amplifying pathway) (20, 21, 26, 27, 54). In normal rodent islets, both triggering and amplifying pathways correlate with changes in the islet ATP/ADP ratio (16, 20) and contribute simultaneously to GSIS under physiological conditions (27). We therefore tested whether the amplifying pathway of GSIS is more sensitive to glucose in High-glucose than in Control islets.
During perifusion with a medium containing 0, 6, or 20 mmol/l glucose (G0, G6, or G20) and 250 µmol/l of the KATP channel opener diazoxide (DZ), [Ca2+]i was higher in High-glucose than in Control islets, but insulin secretion remained below the RIA detection limit in both types of islets (Fig. 3). Upon membrane depolarization with 30 mmol/l K+ (K30), [Ca2+]i rapidly increased to a plateau that was similar in High-glucose and Control islets and was unaffected by the glucose concentration (Fig. 3, A and B). In Control islets, the stimulation of insulin secretion by K30 was markedly glucose dependent, with a monophasic increase in the absence of glucose, a slightly larger and more sustained increase in G6, and a much larger increase in G20 (Fig. 3C). In High-glucose islets, however, K30-induced insulin secretion was already sustained in G0, resembling the response observed in G6 with Control islets, and the maximal amplifying effect of glucose was observed in G6 instead of G20 (Fig. 3D). Thus, in the presence of G6-K30 DZ, insulin secretion was about fourfold greater in High-glucose than in Control islets, even though [Ca2+]i was not different between the two types of islets. In the presence of G20-K30 DZ, the absolute rate of secretion was similar in the two types of islets, despite the 50% lower insulin content of High-glucose islets (329 ± 4 and 645 ± 8 pg insulin/ng DNA in High-glucose and Control islets, respectively, P < 0.0001). These results therefore demonstrate that High-glucose islets are also more sensitive to glucose under conditions that measure the amplifying pathway of GSIS only, and that their maximal rate of secretion, measured under these conditions, is not reduced in parallel with their insulin content.
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In Control islets perifused with G0.5, 45Ca2+ efflux was low and stable (Fig. 4B), and [Ca2+]i was 120 nmol/l and slowly increased with time (Fig. 4C). Under these conditions, 45Ca2+ efflux and [Ca2+]i were significantly higher in High-glucose than in Control islets (P < 0.001; Fig. 4, B and C). In both types of islets, they were unaffected by addition of DZ to the medium but decreased in parallel upon removal of extracellular Ca2+ (Fig. 4, D-F), so that 45Ca2+ efflux and [Ca2+]i remained higher in High-glucose vs. Control islets even in the absence of extracellular Ca2+. These results indicate that the elevations of 45Ca2+ efflux and [Ca2+]i in High-glucose islets perifused with G0.5 do not result from a stimulation of Ca2+ influx and therefore reflect an elevation of resting [Ca2+]i. This may explain why the rate of insulin secretion was not higher in High-glucose than in Control islets during perifusion with G0.5 (Fig. 4A).
Increased glucose sensitivity, but reduced glucose responsiveness, of the triggering pathway of GSIS in High-glucose islets.
When Control islets were stimulated with glucose, 45Ca2+ efflux slightly decreased in G6 and markedly increased in G20 (Fig. 4B). Parallel changes in [Ca2+]i were observed under these conditions (Fig. 4C). Accordingly, insulin secretion was not affected by G6 but was stimulated 30-fold by G20 (Fig. 4A). As expected (27, 28), the rises in 45Ca2+ efflux, [Ca2+]i, and insulin secretion produced by G20 were completely inhibited by DZ in these Control islets (Fig. 4, A-C). When the stimulation by G6 and G20 was applied in the presence of DZ or absence of extracellular Ca2+, 45Ca2+ efflux was inhibited in a concentration-dependent manner (Fig. 4, D and E), likely as a result of the stimulation of Ca2+ pumping by the ER (61). These results therefore confirm that the rise in 45Ca2+ efflux and [Ca2+]i induced by G20 in Control islets results from the stimulation of Ca2+ influx after closure of KATP channels and plasma membrane depolarization (27, 28).
In High-glucose islets, stimulation with G6G20 induced parallel increases in 45Ca2+ efflux and insulin secretion (P < 0.001) that were both completely inhibited by DZ (Fig. 4, A and B). Surprisingly, these effects of glucose and DZ occurred in the absence of detectable [Ca2+]i changes (Fig. 4C). Compared with Control islets, the stimulation of 45Ca2+ efflux and insulin secretion in High-glucose islets occurred at a lower glucose concentration (G6 vs. G20) and was of smaller magnitude in G20. When the stimulation by G6 and G20 was applied in the presence of DZ or absence of extracellular Ca2+, 45Ca2+ efflux was inhibited in a concentration-dependent manner, as in Control islets (Fig. 4, D and E), but the maximal decrease of 45Ca2+ efflux was observed at a lower glucose concentration than in Control islets (G6 vs. G20).
These results demonstrate that, also in High-glucose islets, GSIS requires the closure of -cell KATP channels and the stimulation of Ca2+ influx. They therefore suggest that, under normal ionic conditions, the strong stimulation of insulin secretion by G6 in High-glucose islets reflects almost maximal glucose stimulation of both triggering and amplifying pathways.
Dissociation between [Ca2+]i and Ca2+ influx/insulin secretion in High-glucose islets. In Control islets, glucose stimulation and DZ inhibition of Ca2+ influx were accompanied by the expected increase and decrease in [Ca2+]i (27). This was not the case in High-glucose islets, in which glucose stimulation and DZ inhibition of Ca2+ influx and insulin secretion occurred in the absence of detectable [Ca2+]i changes (see previous paragraph).
What can be the cause of such a dissociation between [Ca2+]i and Ca2+ influx/insulin secretion in High-glucose islets? An experimental artifact of fura PE 3 microspectrofluorimetry seems unlikely, because parallel changes in [Ca2+]i were detected in High-glucose and Control islets upon stimulation with acetylcholine (ACh) in the absence of extracellular Ca2+ (Fig. 5A). Alternatively, the stimulation of Ca2+ influx by glucose could be smaller in High-glucose than in Control islets and escape detection by measuring [Ca2+]i because of the already elevated mean [Ca2+] in the whole cytoplasm. It could, however, be sufficient to increase the [Ca2+] beneath the plasma membrane in the vicinity of voltage-dependent Ca2+ channels (VDCC) and trigger exocytosis of insulin granules in their close proximity (6). This hypothesis is compatible with recent observations suggesting that insulin granule exocytosis may be better controlled by changes in Ca2+ influx through VDCC than by a rise in [Ca2+]i (57).
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After stimulation with G10, removal of extracellular Ca2+ abrogated the glucose-induced [Ca2+]i rise in Control islets and reduced [Ca2+]i by 20 nmol/l in High-glucose islets (Fig. 5A), so that [Ca2+]i remained elevated in High-glucose vs. Control islets. Subsequent addition of ACh to the medium induced a similar transient rise in [Ca2+]i in High-glucose and Control islets (58 ± 3 vs. 54 ± 1 nmol/l, respectively; Fig. 5A), followed by a prolonged lowering of [Ca2+]i that was slightly larger in High-glucose than in Control islets (18 ± 2 vs. 9 ± 1 nmol/l, P < 0.05).
Pretreatment of Control islets with the SERCA [sarco(endo)plasmic reticulum Ca2+-ATPase] pump inhibitor Tg did not affect their resting [Ca2+]i compared with nonpretreated islets from the same culture, but it had three effects: the initial decrease in [Ca2+]i induced by stimulation with G10 was abolished, the fall in [Ca2+]i after omission of extracellular Ca2+ was faster and larger, and ACh was ineffective (both the initial transient peak and the subsequent lowering of [Ca2+]i were abolished) (Fig. 5B). In High-glucose islets pretreated with Tg and perifused with G0.5, [Ca2+]i was lower than in nonpretreated islets from the same culture, so that the difference in [Ca2+]i between High-glucose and Control islets was reduced by about one-third from 50 ± 1 to 34 ± 1 nmol/l (P < 0.001). Despite this reduction, the inability of G10 to increase [Ca2+]i persisted. Both phases of the effect of ACh were also abolished in the absence of extracellular Ca2+ (Fig. 5B).
These results suggest that the elevation of [Ca2+]i in High-glucose islets could partly result from the overfilling of intracellular Ca2+ stores during culture in G30 (cytosolic Ca2+-binding proteins, ER, and/or mitochondria). In support of this hypothesis, we observed that, after 18 h of loading with 45Ca2+, conditions under which we assume that isotopic equilibrium was reached, 1-h treatment with Tg decreased the islet 45Ca2+ content to a much larger extent in High-glucose than in Control islets (Fig. 5D). However, the partial reduction of the elevation of [Ca2+]i produced by Tg pretreatment was not sufficient to unmask a rise in [Ca2+]i upon glucose stimulation of Ca2+ influx. It is clear, therefore, that factors other than ER Ca2+ overfilling contribute to the elevation of resting [Ca2+]i and the absence of glucose-stimulated [Ca2+]i rise in High-glucose islets.
General discussion.
The present study demonstrates that High-glucose islets are more sensitive to glucose for the stimulation of both triggering (Ca2+ influx) and amplifying (augmentation of Ca2+-induced exocytosis) pathways of GSIS, with a lower threshold of glucose concentration for the stimulation of the triggering pathway than in Control islets, and nearly maximal stimulation of both pathways in G6G10 rather than G20. A similar increase in glucose sensitivity was observed for the glucose-induced reduction of Ca2+ efflux in the presence of DZ or absence of extracellular Ca2+. This increase in glucose sensitivity of High-glucose vs. Control islets likely results from their more highly energetic state in the presence of low glucose concentrations. Because energetic metabolism of High-glucose islets is almost maximally stimulated in the presence of G6, the changes in NAD(P)H production, mitochondrial hyperpolarization, ATP production, Ca2+ influx, and insulin secretion that are expected to occur when the glucose concentration is raised from G6 to G10G20 are strongly reduced, when not absent. This might have been interpreted as a lack of glucose responsiveness if the lower glucose concentration of G0.5 had not been tested. Our measurement of an increase in glucose sensitivity of High-glucose islets, which could partly result from the abnormal accumulation of glycogen in islet -cells (41, 44, 67), is in good agreement with data obtained with rat and human islets cultured under similar conditions (4042, 45, 60), with islets from 48-h glucose-infused rats (1), and with islets from Psammomys obesus made diabetic by 5 days on a high-energy diet (48). Other studies suggesting that
-cells exposed to high glucose concentrations become insensitive to glucose may be conflicting at first sight only. Thus the basal glucose concentration used as a reference was higher than here, and presumably already had induced nearly maximal stimulation of
-cells (7, 8, 13, 39, 47, 51).
Besides their increased glucose sensitivity, High-glucose islets displayed an 50% decrease in maximal GSIS under physiological conditions (G20). This reduction, which could result from both
-cell degranulation and reduced glucose stimulation of Ca2+ influx, was no longer observed when [Ca2+]i was further increased by plasma membrane depolarization with 30 mmol/l K+. Our results therefore suggest that correcting the triggering action of glucose (stimulation of Ca2+ influx through VDCC) could be more effective in improving GSIS in type 2 diabetes than further augmenting an already maximally stimulated amplifying action of the sugar. They also suggest that slightly reducing the basal rate of
-cell ATP production may restore GSIS within the physiological range of glucose concentrations while simultaneously decreasing fasting hyperinsulinemia and
-cell degranulation (23, 29).
In conclusion, after 1 wk of culture in G30, the loss of glucose stimulation of insulin secretion within the physiological range of glucose concentrations (from G5 in the fasting state to G10 in the postprandial period) results from the combination of two effects: first, an increased sensitivity to glucose of mitochondrial metabolism and both triggering and amplifying pathways of insulin secretion, which occur despite an approximately twofold increase in UCP2 mRNA expression; second, an 50% reduction in the maximal glucose stimulation of the triggering pathway, hence of the maximal glucose-stimulated insulin secretion under physiological conditions. These alterations are associated with a paradoxical dissociation between glucose-induced changes in Ca2+ influx and the absence of detectable [Ca2+]i rise of unclear origin.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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