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-Cells by Mechanisms that Mirror the Stimulus-Secretion Coupling in ß-Cells
Lilly Research Laboratories (H.L.O., K.Bo., J.G.), D-22419 Hamburg, Germany; Bartholin Instituttet (K.Bu.), Rigshospitalet, DK-2100 Copenhagen, Denmark; and Department of Cell Physiology and Metabolism (S.T., C.B.W.), University Medical Center, 1211 Geneva 4, Switzerland
Address all correspondence and requests for reprints to: Jesper Gromada, Lilly Research Laboratories, Essener Bogen 7, D-22419 Hamburg, Germany. E-mail: gromada{at}lilly.com.
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Abstract |
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-cells, glucose, arginine, and the sulfonylurea tolbutamide stimulated glucagon release. The effect of glucose was abolished by the KATP-channel opener diazoxide as well as by mannoheptulose and azide, inhibitors of glycolysis and mitochondrial metabolism. Glucose inhibited KATP-channel activity by 30% (P < 0.05; n = 5) and doubled the free cytoplasmic Ca2+ concentration. In cell-attached recordings, azide opened KATP channels. The N-type Ca2+-channel blocker 
-conotoxin and the Na+-channel blocker tetrodotoxin inhibited glucose-induced glucagon release whereas tetraethylammonium, a blocker of delayed rectifying K+ channels, increased secretion. Glucagon release increased monotonically with increasing K+ concentrations. -Conotoxin suppressed glucagon release to 15 mM K+, whereas a combination of -conotoxin and an L-type Ca2+-channel inhibitor was required to abrogate secretion in 50 mM K+. Recordings of cell capacitance revealed that glucose increased the exocytotic response evoked by membrane depolarization 3-fold. This correlated with a doubling of glucagon secretion by glucose in intact rat islets exposed to diazoxide and high K+. In whole-cell experiments, exocytosis was stimulated by reducing the cytoplasmic ADP concentration, whereas changes of the ATP concentration in the physiological range had little effect. We conclude that glucose stimulates glucagon release from isolated rat -cells by KATP-channel closure and stimulation of Ca2+ influx through N-type Ca2+ channels. Glucose also stimulated exocytosis by an amplifying mechanism, probably involving changes in adenine nucleotides. The stimulatory action of glucose in isolated -cells contrasts with the suppressive effect of the sugar in intact islets and highlights the primary importance of islet paracrine signaling in the regulation of glucagon release. |
Introduction |
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-cells. The ability of high glucose concentrations to suppress glucagon release has been attributed to both direct actions on the -cells (1, 2, 3) and paracrine effects exerted by factors released by neighboring ß-cells and 
-cells. Candidate paracrine inhibitors of glucagon secretion include insulin (4, 5, 6, 7, 8), Zn2+, which is cosecreted with insulin (6, 7), and 
-amino butyric acid (GABA) (9, 10, 11) as well as somatostatin (12, 13). Insulin is also known as a physiological suppressor of glucagon secretion in vivo (4).
Pancreatic -cells are electrically excitable and generate spontaneous Na+- and Ca2+-dependent action potentials (14, 15, 16, 17). Glucagon release is a Ca2+-dependent process (18, 19, 20, 21), and both capacitance recordings and hormone release measurements have revealed a close relationship between N-type Ca2+ channels and secretion at low glucose concentrations (11, 12, 16). Interestingly, pancreatic -cells are equipped with ATP-sensitive K+ channels (KATP channels) of the same type as those constituting the resting conductance in ß-cells (15, 22, 23, 24, 25). We have recently provided evidence that in mouse -cells KATP channels are involved in glucose regulation of glucagon secretion by controlling the membrane potential in a way reminiscent of that previously described for the ß-cell (25). However, because mouse -cells possess a different complement of voltage-gated ion channels involved in action potential generation from the ß-cell, moderate membrane depolarization in mouse -cells is associated with reduced rather than increased electrical activity and secretion (3, 25).
Different mechanisms may control glucagon secretion in different species because it has been reported that rat -cells share common features with ß-cells (6, 7). Both cell types transduce a metabolic signal (ATP) into ionic signals (membrane depolarization and Ca2+ influx) and stimulation of secretion in isolated cells. However, it is the simultaneous activation of ß-cells within the intact islet that inhibits Ca2+ influx and glucagon secretion in rat -cells (6, 7). Here we have applied measurements of whole-cell conductances and cell-attached recordings of KATP-channel activity as well as imaging of the intracellular free Ca2+ concentration, [Ca2+]i, and glucagon secretion measurements to study the stimulus-secretion coupling in isolated rat -cells. We show that glucose stimulates glucagon secretion in isolated -cells and that the stimulus-secretion coupling in the rat -cell mirrors that of the ß-cell.
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Materials and Methods |
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-cells-cells were obtained by fluorescence-activated cell sorting (FACS) as described elsewhere (26). Based on the hormone contents (26) and immunohistochemistry, we estimate that the preparations contain more than 90% -cells and less than 3% ß-cells. To minimize any perturbations of -cell function and glucagon secretion by the low number of contaminating ß- and -cells, we have used zero glucose as basal condition to maximally inhibit the release of insulin and somatostatin. Cells were plated on plastic petri dishes (Nunc A/S, Roskilde, Denmark) and, for microfluorometry of the [Ca2+]i, on 22-mm glass coverslips and maintained for 2 d in RPMI 1640 medium (Invitrogen, Carlsbad, CA) supplemented with 10% (vol/vol) heat-inactivated fetal calf serum, 100 IU/ml penicillin, and 100 µg/ml streptomycin at 37 C in a humidified atmosphere. Measurement of glucagon content revealed no difference between freshly isolated (11.8 ± 0.9 ng glucagon per 1000 cells) and 2-d cultured -cells (10.7 ± 1.4 ng glucagon per 1000 cells; three different preparations).
Hormone secretion assays
FACS-isolated -cells were seeded on polyornithine-coated 24-well plates (10,000 cells per well) and cultured overnight in RPMI 1640 medium supplemented with 10% (vol/vol) heat-inactivated fetal calf serum, 100 IU/ml penicillin, and 100 µg/ml streptomycin at 37 C in a humidified atmosphere. Cells were washed twice with 0.5 ml Krebs Ringer bicarbonate HEPES (KRBH) buffer consisting of (in mM) 115 NaCl, 4.7 KCl, 2.6 CaCl2, 1.2 NaH2PO4, 1.2 MgCl2, 20 NaHCO3, 0.5% BSA (fraction V), and 10 HEPES (pH 7.4 with NaOH) and then preincubated in 0.5 ml of the same buffer for 30 min at 37 C. After a second wash, cells were incubated at 37 C for 30 min with KRBH buffer supplemented with different glucose concentrations and additional reagents, as indicated in the text. At the end of the test incubation, the medium was aspirated and assayed for glucagon using a commercial assay (Linco Research, St. Charles, MO). Glucagon release from intact islets was performed as described previously (27) using KRBH buffer. Tetrodotoxin (TTX) and thapsigargin were purchased from Alomone Labs (Jerusalem, Israel). SNX482 was from Peptides International (Louisville, KY). Mannoheptulose was obtained from Glycoteam GmbH (Hamburg, Germany). All other chemicals were from Sigma Chemical Co. (St. Louis, MO).
Electrophysiology
Patch-clamp electrodes were pulled from borosilicate capillaries, coated with Sylgard, and fire polished. The pipette resistance was between 2.5 and 4 M
when filled with the pipette solutions as specified below. Whole-cell KATP currents were recorded by applying 10-mV hyper- and depolarizing voltage pulses (duration, 200 msec; pulse interval, 2 sec) from a holding potential of –70 mV using the perforated-patch whole-cell configuration and an EPC-9 patch-clamp amplifier (Heka Elektronik, Lambrecht/Pfalz, Germany). The extracellular solution consisted of (in mM) 138 NaCl, 5.6 KCl, 2.6 CaCl2, 1.2 MgCl2, and 5 HEPES (pH 7.4 with NaOH) and supplemented with glucose as indicated. The pipette solution consisted of (in mM) 76 K2SO4, 10 KCl, 10 NaCl, 1 MgCl2, 5 HEPES (pH 7.35 with KOH), and 0.24 mg/ml of the pore-forming antifungal agent amphotericin B. Perforation required a few minutes, and the voltage clamp was considered satisfactory when the Gseries (series conductance) was stable and greater than 35 nS.
Recordings of KATP channels in the cell-attached recording mode were performed using the same extracellular solution as described above and with a pipette-filling solution containing (in mM) 140 KCl, 1 CaCl2, 1 MgCl2, and 10 HEPES (pH 7.3 with KOH). The pipette potential was held at 0 mV. The current signal was sampled at 4 kHz and filtered at 2 kHz, using the internal filter of the EPC-9 amplifier.
Exocytosis was monitored in single -cells as changes in cell capacitance using either the standard or the perforated-patch whole-cell configuration. An EPC-7 patch-clamp amplifier (List Elektronik, Darmstadt, Germany) was used, and exocytosis was elicited by 500-msec voltage-clamp depolarizations from –70 to 0 mV. Changes in cell capacitance were detected using in-house software written in Axobasic (Axon Instruments, Foster City, CA) as detailed elsewhere (28). The pipette solution for the perforated-patch experiments consisted of (in mM) 76 Cs2SO4, 10 KCl, 10 NaCl, 1 MgCl2, 5 HEPES (pH 7.35 with KOH), and 0.24 mg/ml amphotericin B. The pipette solution used for standard whole-cell recordings contained (in mM) 125 Cs-glutamate, 10 CsCl, 10 NaCl, 1 MgCl2, 5 HEPES, 0.05 EGTA, and 0.01 GTP as well as MgATP and ADP as indicated in the text (pH 7.15 using CsOH). The extracellular medium consisted of (in mM) 118 NaCl, 20 tetraethylammonium (TEA)-Cl, 5.6 KCl, 1.2 MgCl2, 2.6 CaCl2, and 5 HEPES (pH 7.40 with NaOH) and supplemented with test substances as indicated. All measurements were performed at 33 C, and the recording chamber was perfused at a rate of 1.5 ml/min.
Measurements of [Ca2+]i
The [Ca2+]i measurements were made using an Axiovert 135 inverted microscope with a Plan-Neofluar x100/NA 1.30 objective (Carl Zeiss, Göttingen, Germany) and an Ionoptix (Milton, MA) fluorescence imaging system. Excitation was effected at 340 and 380 nm and emitted light recorded at 510 nm with a video camera synchronized to the excitation light source and a computer interface. Cells were loaded for 20 min with 0.4 µM fura-2/AM (Molecular Probes, Eugene, OR) in extracellular solution with 2.5 mM glucose. The cells were perfused at a rate of 1.5 ml/min with extracellular solution at 33 C. Calibration of the fluorescence signal was performed by infusing single -cells using the standard whole-cell configuration of the patch-clamp technique with fura-2 and Ca2+-EGTA buffers (Molecular Probes) with known free Ca2+ concentrations ranging between 0 and 39.8 µM. These data were fitted to the equation described in Ref.29 and used to calculate [Ca2+]i.
cAMP measurements
Isolated rat -cells were plated (2000 cells per well) and cultured overnight as described above. Cells were preincubated for 1 h at 37 C in KRBH buffer supplemented with 0.5 mM of the phosphodiesterase inhibitor IBMX and incubated for another 30 min in the same buffer with 0 or 16.8 mM glucose. The reaction was stopped and the cells were lysed by the addition of HCl (50 mM final concentration) for 30 min and neutralized by an equimolar amount of NaOH. The cAMP content of the cell lysates was determined using the overnight acetylation protocol of the [125I]cAMP scintillation proximity assay (SPA) kit (Biotrak cAMP assay; Amersham, Little Chalfont, UK). In short, lysates and cAMP standards were acetylated using the kit acetylation reagent and incubated with [125I]cAMP plus cAMP antibodies overnight. The next day, the SPA beads were added, the antibody-bound fraction was separated by centrifugation, and the samples were decanted and counted in a -scintillation counter.
Statistical analysis
Results are presented as mean values ± SE for the indicated number of experiments. Statistical significances were evaluated using Student’s t test for pairs of data, Dunnett’s test for multiple comparisons with a control, and Tukey’s test when multiple comparisons between groups were required.
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Results |
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-cells-cells to study the effects of glucose on glucagon release. We found that glucose stimulated glucagon release in a dose-dependent manner (EC50 = 3.4 mM) (Fig. 1A
). Glucose did not elevate intracellular cAMP concentration (Table 1), whereas epinephrine (5 µM) via activation of ß-adrenergic receptors (16, 30) produced a more than 4-fold stimulation. These observations are consistent with an earlier report by Pipeleers et al. (21). Thus, the effects of glucose are unlikely to be mediated by an increase in intracellular cAMP with resulting stimulation of glucagon secretion.
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). Diazoxide (100 µM), an opener of KATP channels, suppressed basal glucagon secretion by 40% (P < 0.05; n = 5) and abolished glucose-induced glucagon secretion (Fig. 1B). This is supported by our previous findings that rat -cells express KATP channels and that diazoxide inhibits spontaneous electrical activity (15). Sodium azide, an inhibitor of mitochondrial cytochrome c oxidase and consequently ATP formation suppressed basal release by 50% (P < 0.05; n = 5) and completely abolished glucose-induced glucagon secretion (Fig. 1B). Consistent with the expression of glucokinase in rat -cells (31), we found that mannoheptulose (10 mM) suppressed glucose-induced glucagon secretion (Fig. 1B).
Figure 1C shows that in an extracellular medium without glucose, closure of KATP channels by the sulfonylurea tolbutamide (100 µM) or depolarization of the -cell with 10 mM arginine produced robust stimulation of glucagon secretion. Comparable stimulation of glucagon release was induced by epinephrine (5 µM) (Fig. 1C). Glucose (16.8 mM) enhanced the stimulatory effects of tolbutamide, arginine, and epinephrine on glucagon release (Fig. 1C). It has previously been reported that glucose inhibits glucagon release from isolated rat -cells in the presence of amino acids and epinephrine (21). The reason for this discrepancy is currently unclear and is unlikely to result from differences in cell culture conditions. Furthermore, we have not observed changes in ion channel composition or current densities of KATP and Na+ channels as well as N- and L-type Ca2+ channels in the isolated -cells cultured for 2 d compared with freshly isolated cells (data not shown). These data suggest that cultured -cells show an equivalent channel phenotype to freshly isolated cells. Furthermore, Fig. 1D shows that the isolation procedure is not harmful to the cells and causes them to lose a signaling mechanism present in situ because the paracrine signals, insulin (100 nM), Zn2+ (30 µM), and GABA (0.5 mM) inhibit glucagon release from isolated -cells in both the absence and presence of 16.8 mM glucose. The above results suggest that isolated and cultured -cells maintain their phenotype and sensitivity to paracrine signaling molecules and that glucose via mitochondrial metabolism and KATP-channel closure is implicated in the initiation of glucagon secretion.
Effects of glucose on KATP-channel activity in -cells
We next measured KATP-channel activity in single rat -cells using the perforated-patch whole-cell configuration. In the absence of glucose, the membrane conductance normalized to cell capacitance amounted to 0.48 ± 0.09 nS/pF (n = 6). Addition of 20 mM glucose led to a 26 ± 7% (P < 0.05; n = 6) reduction in the membrane conductance, and the normalized conductance amounted to 0.36 ± 0.06 nS/pF (n = 6) (Fig. 2A). Tolbutamide (100 µM) reduced the input conductance by 84 ± 9% (P < 0.01; n = 5) (Fig. 2B). These and the above data suggest that, like in ß-cells, glucose metabolism and increased ATP production in rat -cells leads to KATP-channel closure and ultimately glucagon secretion. This concept is supported by the previous findings that glucose stimulates ATP production (6) and that tolbutamide stimulates electrical activity in single rat -cells (15).
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). Application of sodium azide (2 mM), which inhibits mitochondrial cytochrome c oxidase leading to a reduction in cellular ATP levels, resulted in activation of channels and cessation of action potential firing. The channel activated by sodium azide was identified as the KATP channel because they were rapidly and reversibly blocked by 200 µM tolbutamide. Furthermore, channel blockage was associated with the reappearance of action potentials. These experiments demonstrate, for the first time on the single-channel level in intact cells, that the KATP channel in -cells is activated by lowering the ATP-to-ADP ratio. These results thus reinforce the conclusions reached above from whole-cell experiments.
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-cells shows a recording of [Ca2+]i from an individual -cell initially exposed to glucose-free medium. Spontaneous oscillations in [Ca2+]i was observed in 27 of 45 cells. The average [Ca2+]i measured in the absence of glucose was 132 ± 14 nM (n = 45). Increasing the glucose concentration to 20 mM resulted in an initial decrease in [Ca2+]i (27 ± 6 nM in amplitude) followed after 3.7 ± 0.9 min by a pronounced and monophasic [Ca2+]i rise (peak amplitude, 254 ± 23 nM; P < 0.05; n = 14) (Fig. 4A). Subsequent application of epinephrine (5 µM) produced a transient elevation in [Ca2+]i, confirming the identity of the -cells (Fig. 4A). The ability of glucose to stimulate an increase in [Ca2+]i was inhibited in the presence of 10 mM mannoheptulose, whereas subsequent addition of 100 µM tolbutamide produced a prompt and reversible increase in [Ca2+]i to a peak amplitude of 344 ± 19 nM (P < 0.01; n = 9) (Fig. 4B).
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-cells-cells (3, 14, 15, 16, 25). We therefore correlated the electrophysiological recordings to changes in glucagon secretion from isolated rat -cells. The importance of voltage-gated Na+ channels for -cell electrical activity was underscored by the strong inhibitory action of the Na+-channel blocker TTX (0.1 µg/ml) of both basal and glucose-stimulated (16.8 mM) secretion (Table 2). The significance of Ca2+ influx through plasma membrane voltage-gated Ca2+ channels for glucagon secretion was illustrated by the 37 ± 6% inhibition by 5 mM Co2+ of basal secretion and complete suppression of glucose-induced release. These inhibitory effects were mimicked by the N-type Ca2+-channel blocker -conotoxin (1 µM) whereas addition of the L-type Ca2+-channel inhibitor nifedipine (50 µM) or the R-type Ca2+-channel blocker SNX482 (100 nM) lacked inhibitory action (Table 2). This corroborates the notion that basal glucagon secretion depends principally on Ca2+ influx through N-type Ca2+ channels (11, 12, 16) whereas L-type Ca2+ channels play an important role for regulation of secretion after activation of protein kinase A (12, 16) or in response to strong membrane depolarization (see below). We also tested the effects of 4-aminopyridine (4-AP), a blocker of a rapidly activating and inactivating TEA-resistant K+ current. Contrary to mouse -cells (25), application of 5 mM 4-AP did not affect glucagon secretion (Table 2). This contrasts to the stimulation of both basal and glucose-evoked secretion in the presence of 20 mM TEA, a blocker of delayed rectifying K+ channels (Table 2).
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-cell electrical activity (33). Table 3 compares the ability of glucose to stimulate glucagon secretion in batches of rat -cells in the absence and presence of thapsigargin, an inhibitor of sarcoplasmic-endoplasmic reticulum calcium ATPase (34). It can be seen that increasing the glucose concentration from 0 to 16.8 mM produced 70% stimulation of glucagon secretion and that this effect was not affected by 5 µM thapsigargin (Table 3).
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). This contrasts to our earlier observation in mouse -cells where increasing the extracellular K+ concentration up to 15 mM resulted in inhibition of glucagon release and only K+ concentrations beyond 25 mM produced stimulation of secretion (25).
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shows that the effect of raising the external K+ concentration to 15 mM on glucagon release was suppressed by inclusion of 1 µM -conotoxin in the extracellular solution. On the contrary, nifedipine (50 µM) did not affect secretion in the presence of 15 mM K+. Rather different results were obtained after application of 50 mM K+. Under these experimental conditions, -conotoxin reduced glucagon release by 40%, whereas the combination of -conotoxin and nifedipine suppressed secretion to the level observed in the presence of 5.6 mM K+ (Fig. 5B). -Conotoxin and nifedipine did not significantly reduce glucagon release under basal conditions (5.6 mM K+).
Effects of glucose on glucagon release from intact islets exposed to diazoxide and elevated K+
In Fig. 6A (closed circles) we investigated the effects of glucose on glucagon release from intact islets depolarized with 30 mM K+ and in the presence of diazoxide (250 µM). Under these experimental conditions, where the membrane potential is clamped and action potential firing is suppressed, glucose enhanced glucagon secretion in a dose-dependent manner (EC50 = 3.1 mM). This effect must have occurred independent of KATP-channel closure because this pathway is bypassed under these conditions (35, 36). This contrasts to a clear and dose-dependent inhibition of glucagon secretion from parallel experiments in which the islets were exposed to increasing glucose concentrations in normal KRBH buffer containing 4.7 mM K+ and no added diazoxide (Fig. 6A, open circles). Figure 6B shows the corresponding insulin secretion from the same islets as described above. Under control conditions, a sigmoidal relationship between the glucose concentration in the extracellular medium and insulin release was observed. In extracellular solution with elevated K+ concentration and diazoxide, glucose again produced a dose-dependent increase in insulin secretion. Consistent with a previous study (35), the dose-response curve displayed two components. A first increase was observed between 0 and 3 mM glucose, whereas the second increase occurred at more than 6 mM. These data support our conclusion that islet paracrine action is important in the regulation of glucagon release. Figure 6C shows that mannoheptulose (10 mM) did not significantly reduce glucagon secretion in depolarized islets in the absence of glucose. The lack of effect of mannoheptulose reflects the fact that glucagon secretion in zero glucose is mainly controlled by the depolarization-induced Ca2+ influx through the voltage-gated Ca2+ channels, a process that is distal to inhibition of glucokinase by mannoheptulose. However, mannoheptulose treatment completely suppressed the ability of glucose (16.8 mM) to enhance glucagon secretion (Fig. 6C). This is consistent with the data that mannoheptulose suppressed glucose-induced but not basal glucagon secretion in isolated -cells (see Fig. 1B).
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-cell surface area that occurs when the glucagon-containing granules fuse with the plasma membrane. Figure 7A illustrates whole-cell Ca2+ currents and the associated changes in cell capacitance elicited by 500-msec depolarizations from –70 to 0 mV in an intact -cell using the perforated-patch configuration. In the absence of glucose, the integrated Ca2+ current amounted to 6.1 pC and a capacitance increase of 22 fF was evoked. The latter value corresponds to the discharge of 11 granules using a conversion factor of 2 fF per granule (14). Six minutes after inclusion of 20 mM glucose in the bathing solution, the same membrane depolarization produced an integrated Ca2+ current of 7.6 pC (25% stimulation) and a capacitance increase of 69 fF (214% stimulation). On average, glucose produced a 178 ± 18% (P < 0.05; n = 7) stimulation of exocytosis. The effect of glucose on exocytosis was associated with a 26 ± 8% (P < 0.05; n = 7) enhancement of the integrated Ca2+ current (Fig. 7B). This stimulation of Ca2+ influx is likely to account for only a minor fraction of the total stimulatory action of glucose on exocytosis (16). Addition of 3 mM sodium azide to the extracellular solution suppressed the stimulatory action of glucose on the whole-cell Ca2+ current and even reduced the capacitance increase to 50% of that observed in the absence of glucose (Fig. 7A). The stimulatory action of glucose on exocytosis was secondary to metabolism and was abolished by inclusion of mannoheptulose (10 mM) in the extracellular medium (Fig. 7C). Under these experimental conditions, subsequent application of pyruvate (5 mM) to the extracellular solution produced a robust stimulation of exocytosis (Fig. 7, C and D).
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). In the absence of glucose, the capacitance increase elicited by the train amounted to 54 ± 15 fF (n = 6; Fig. 8A). After addition of glucose, the amplitude of the capacitance increase was stimulated 3.3-fold and averaged 178 ± 13 fF (P < 0.01; n = 6; Fig. 8B). Secretory granules in rat -cells, like other hormone-releasing cells, can be functionally subdivided into a reserve pool and a limited readily releasable pool (RRP), which undergoes rapid exocytosis upon stimulation (12, 16). The exhaustion of the exocytotic response during the train is likely to reflect depletion of the RRP rather than inactivation of the Ca2+ current with resulting suppression of Ca2+-induced exocytosis. This notion is supported by the observation that the integrated Ca2+ current measured at the end of the train, when secretion had ceased, was reduced by only 24 ± 9% (n = 6) with respect to the first depolarization.
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-cell was dialyzed with a solution containing 3 mM ATP. After establishment of the whole-cell configuration, the cell was allowed a 2-min equilibration period. A train consisting of 10 500-msec depolarizations from –70 to 0 mV was then applied to evoke exocytosis. In a series of five experiments, the total increase in cell capacitance amounted to 171 ± 24 fF (Fig. 9A, left trace). A second train applied to the same cell after an interval of 3 min evoked a capacitance increase of 164 ± 21 fF (Fig. 9A, right trace). When the same experiment was repeated after inclusion of both 5 mM ADP and 3 mM ATP in the pipette-filling solution, the first train was similar to that observed in the presence of standard ATP and averaged 176 ± 23 fF (n = 5; Fig. 9B, left trace). However, exocytosis during the second train (Fig. 9B, right trace) was strongly suppressed, and the total increase averaged 23 ± 11 fF (P < 0.01; n = 5). The ability of ADP to inhibit exocytosis was concentration dependent, and a half-maximal inhibitory action was observed at 0.39 mM (Fig. 9C). This suggests that ADP interferes with the refilling of the RRP but not exocytosis of granules that have already progressed into this pool. ATP (in the absence of ADP) stimulated exocytosis with an EC50 value of 0.67 mM (Fig. 9D). However, it is notable that exocytosis is little affected by variations of the intracellular ATP concentration between 1 and 5 mM. Figure 9E shows that AMP (5 mM) did not affect exocytosis, whereas inclusion of 5 mM GDP in the pipette-filling solution was associated with a 22% inhibition of exocytosis (P < 0.05; n = 5).
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Discussion |
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-cells by a mechanism that mirrors that of the ß-cell. This effect is secondary to glucose metabolism, closure of KATP channels, membrane depolarization, and stimulation of Ca2+ influx. Importantly, we have also uncovered an amplifying action of glucose on Ca2+-induced exocytosis. In the intact rat islet, glucose suppresses glucagon release. These findings therefore redefine our understanding of the rat -cell stimulus-secretion coupling and emphasize the role of islet paracrine signaling as the primary regulatory mechanism governing glucagon secretion in the intact rat islet.
Rat -cells express both glucokinase and the glucose transporter GLUT1, a lower capacity isoform than GLUT2, expressed in ß-cells of this species (31). They have a high ATP concentration (6.5 mM) and a high ATP-to-ADP ratio already at 1 mM glucose (39), and although steady-state glucose utilization is the same (40), glucose oxidation in rat -cells is only 30% of that in ß-cells (41). However, our data clearly show that sufficient ATP was generated within 6 min after glucose stimulation to reduce KATP-channel activity, elevate [Ca2+]i, and stimulate Ca2+-dependent exocytosis. We have previously reported that glucose inhibits spontaneous electrical activity in rat -cells (15). Our measurements of [Ca2+]i now show that this observation can be explained by the fact that, as in ß-cells, glucose-induced membrane depolarization is preceded by a transient hyperpolarization (reflected as a decrease in [Ca2+]i), which is likely to result from the initial ATP consumption by glucokinase (42, 43) and Ca2+ uptake into the endoplasmic reticulum (44). However, this hyperpolarization could also result from deactivation of a Ca2+ release-activated Ca2+ current and occurs over a much longer time (minutes rather than seconds in -cells) because of the relatively slow rate of glucose metabolism.
We have previously demonstrated that the maximal input conductance after complete wash-in of a pipette solution with low ATP and ADP content (using the standard whole-cell configuration) was 10 nS/pF (15). The input conductance measured in intact rat -cells in the absence of glucose (0.5 nS/pF) suggest that only 5% of the total KATP conductance is activated even when the -cells are exposed to glucose-free solution. The low KATP-channel conductance, because of the high ATP-to-ADP ratio (39), keeps the membrane potential sufficiently depolarized to allow spontaneous electrical activity even in the absence of glucose (15, 16, 27). This concept was supported by the finding that in the absence of glucose, in cell-attached recordings, KATP-channel activity was very low and accompanied by electrical activity, whereas application of sodium azide caused the prompt activation of channels sensitive to tolbutamide.
The concept of a similar mechanism of cell activation in - and ß-cells is supported by the observations that 1) high external K+ stimulates glucagon secretion (present study and Ref.45); 2) diazoxide suppressed glucose-induced glucagon secretion by KATP-channel activation and membrane hyperpolarization (15); and 3) tolbutamide stimulates electrical activity (15) and glucagon secretion (present study and Ref.7). Tolbutamide also stimulates glucagon release from perfused rat pancreas (46, 47) and circulating glucagon levels in patients with advanced type 1 diabetes (48).
We demonstrate here that glucagon secretion in response to glucose stimulation and moderate K+ depolarization is severely compromised by -conotoxin, an inhibitor of N-type Ca2+ channels. This is consistent with the observation that glucagon secretion triggered by hypoglycemia in intact islets depends principally on Ca2+ influx through N-type Ca2+ channels (12, 16, 49). We also observed that TTX inhibited glucose-evoked glucagon secretion as effectively as -conotoxin, confirming our previous data that a prominent voltage-gated and TTX-sensitive Na+ current is activated during the action potential and contributes to the fact that rat -cells, in contrast to ß-cells, produced overshooting action potentials (i.e. exceed 0 mV) (15, 16, 27). Furthermore, our data suggest that, like in the ß-cell, delayed rectifying K+ channels are required to restore the negative membrane potential after each action potential.
Based on [Ca2+]i measurements in isolated mouse -cells, it has recently been suggested that a store-operated membrane conductance plays a pivotal role in the regulation of glucagon secretion in mouse -cells (33). At low glucose, intracellular Ca2+ stores are empty, leading to activation of the depolarizing conductance with resulting initiation of -cell electrical activity and stimulation of glucagon secretion. After an increase in glucose concentration, metabolism is accelerated and the intracellular Ca2+ stores are filled, leading to a reduction of conductance, membrane repolarization, and suppression of glucagon secretion (33). However, this mechanism is not likely to govern glucagon secretion in rat -cells because thapsigargin did not affect the ability of glucose to stimulate secretion.
In the ß-cell, glucose stimulates insulin secretion not only by the well characterized triggering pathway involving glucose metabolism, closure of KATP channels, membrane depolarization, and Ca2+ influx but also by generating amplifying signals. The amplifying pathway has been intensively studied in ß-cells (35, 36). Under this condition and in intact rat islets, we found that glucose stimulated glucagon secretion. This increase in secretion was dose dependent, required glucose metabolism, and is likely to occur independently of paracrine signaling in the islet. Using capacitance measurements, we show that the stimulatory effect of glucose results from acceleration of granule mobilization resulting in a 3.3-fold increase in the number of readily releasable granules. The process by which granules attain release competence is poorly characterized in -cells but requires ATP hydrolysis (12). We now show that the effect of ATP on exocytosis is dose dependent (EC50 = 0.67 mM) and that the secretory capacity is suppressed by ADP (IC50 = 0.39 mM). It is therefore likely that the amplifying action of glucose on glucagon secretion results from an increase in the ATP-to-ADP ratio secondary to glucose metabolism. This is supported by the observations that mannoheptulose inhibited glucose-induced glucagon secretion from intact islets exposed to high K+ and diazoxide as well as glucagon secretion and Ca2+-dependent exocytosis in isolated -cells after glucose stimulation.
This study demonstrates that removing the rat -cell from the repressive environment in the intact islet has enabled us to characterize the signaling pathways leading to glucagon secretion. These findings also uncover islet paracrine signaling as the primary regulatory mechanism governing glucagon secretion. There is now strong evidence that the ß-cell secretory products Zn2+ (6, 7), insulin (4, 5, 6, 7, 8), and GABA (9, 10, 11) inhibit glucagon release. Furthermore, paracrine inhibition of glucagon secretion by somatostatin released from neighboring -cells also contributes to the glucose inhibitory action (12, 13, 50). The stimulatory action of glucose in intact depolarized (K+ and diazoxide) islets on glucagon release suggests that the paracrine inhibitory actions of the ß- and -cell secretory products primarily occurs via modulation of plasma membrane ion channel activity and inhibition of electrical activity. This is consistent with the observation that insulin secretion is stimulated under the same experimental conditions and that we and others show that insulin and Zn2+ inhibit glucagon secretion by KATP-channel activation (7) and that GABA reduces glucagon release by activation of Cl– currents in the -cell plasma membrane (9, 11).
In conclusion, our study demonstrates that the stimulus-secretion coupling in the rat -cell mirrors that of the ß-cell and that glucose stimulates glucagon secretion in isolated -cells. This clearly contrasts to the suppressive effect of glucose in intact islets and highlights the importance of islet paracrine signaling as the primary regulatory mechanism governing glucagon secretion. Finally, this study suggests that reduced paracrine signaling caused by loss of ß-cell function leads to hyperactivity of neighboring -cells and may account for the hyperglucagonemia associated with type 2 diabetes.
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Present address for H.L.O.: Sophion Bioscience A/S, Baltorpvej 154, DK-2750 Ballerup, Denmark.
First Published Online August 4, 2005
Abbreviations: 4-AP, 4-Aminopyridine; FACS, fluorescence-activated cell sorting; GABA, -amino butyric acid; KRBH, Krebs Ringer bicarbonate HEPES; RRP, readily releasable pool; TEA, tetraethylammonium; TTX, tetrodotoxin.
Received June 29, 2005.
Accepted for publication July 29, 2005.
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Cm; lower) evoked by 500-msec depolarization from –70 to 0 mV (Vm; top) in the absence of glucose (left), 6 min after the addition of 20 mM glucose (middle), and 2 min after application of 3 mM sodium azide (azide) in the continuous presence of 20 mM glucose (right). B, Histogram showing the capacitance increases (
