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Am J Physiol Endocrinol Metab 289: E900-E908, 2005. First published June 14, 2005; doi:10.1152/ajpendo.00101.2005
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Role of high-voltage-activated calcium channels in glucose-regulated {beta}-cell calcium homeostasis and insulin release

James T. Taylor,1 Luping Huang,1 Brian M. Keyser,1 Hean Zhuang,2 Craig W. Clarkson,1 and Ming Li1

1Department of Pharmacology, Tulane University Health Sciences Center, New Orleans, Louisiana; and 2Department of Pharmacology, University of South Alabama College of Medicine, Mobile, Alabama

Submitted 8 March 2005 ; accepted in final form 2 June 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
High-voltage-activated (HVA) calcium channels are known to be the primary source of calcium for glucose-stimulated insulin secretion. However, few studies have investigated how these channels can be regulated by chronically elevated levels of glucose. In the present study, we determined the level of expression of the four major HVA calcium channels (N-type, P/Q-type, LC-type, and LD-type) in rat pancreatic {beta}-cells. Using quantitative real-time PCR (QRT-PCR), we found the expression of all four HVA genes in rat insulinoma cells (INS-1) and in primary isolated rat islet cells. We then determined the role of each channel in insulin secretion by using channel-selective antagonists. Insulin secretion analysis revealed that N- and L-type channels are both involved in immediate glucose-induced insulin secretion. However, L-type was preferentially coupled to secretion at later time points. P/Q-type channels were not found to play a role in insulin secretion at any stage. It was also found that long-term exposure to elevated glucose increases basal calcium in these cells. Interestingly, chronically elevated glucose decreased the mRNA expression of the channels involved with insulin secretion and diminished the level of stimulated calcium influx in these cells. Using whole cell patch clamp, we found that N- and L-type channel currents increase gradually subsequent to lower intracellular calcium perfusion, suggesting that these channels may be regulated by glucose-induced changes in calcium.

glucose desensitization; insulinoma cells; diabetes

HIGH-VOLTAGE-ACTIVATED (HVA) CALCIUM CHANNELS are membrane-spanning proteins that are involved in the regulation of intracellular calcium in many different cell types (14). The HVA calcium channel family is composed of seven different genes (A-F, S) that encode the various {alpha}1-pore-forming subunits (7, 22). Calcium entry through these channels plays a key role in a variety of physiological responses, including membrane excitability and modulation of transmitter or hormone release. However, the differences in tissue distribution and efficacy with which HVA channels stimulate vesicular secretion vary among the channel classes (20). In pancreatic {beta}-cells, the expression of many HVA calcium channels' mRNA has been demonstrated (5, 13, 27, 30, 33, 39, 46, 58, 64, 68). These include the {alpha}1A (Cav2.1), {alpha}1B (Cav2.2), {alpha}1C (Cav1.2), {alpha}1D (Cav1.3), and {alpha}1E (Cav2.3). However, only a few studies have been published to show HVA calcium channel protein expression in pancreatic {beta}-cells. Expression of the {alpha}1C protein has been found in mouse islets cells, whereas the {alpha}1D protein has been shown to be expressed in normal (42) and obese mouse islet cells (69) and in rat islet {beta}-cells (28). Protein expression of {alpha}1A, {alpha}1B, and {alpha}1E has also been found in rat {beta}-cells (33, 60, 63).

Even though many types of HVA channels have been identified in pancreatic {beta}-cells, it is generally accepted that HVA calcium channels sensitive to dihydropyridines (DHPs, L-type) contribute to most of the {beta}-cell calcium current and subsequent insulin secretion (9, 26). However, it is unclear as to which L-type isoform (Cav1.2 or Cav1.3) is directly linked with insulin secretion. Cav1.3 has been shown to be preferentially coupled to insulin secretion in the rat {beta}-cell line INS-1 and rat islet cells (33, 35), whereas Schulla et al. (56) demonstrated that the Cav1.2 was coupled with insulin secretion in Cav1.2-null mice. It has been found that Cav1.3 is ~10 times less sensitive to block by DHPs than Cav1.2 (21), and doses required to sufficiently inhibit L-type activity in pancreatic {beta}-cells are above 10 µM, which suggests that Cav1.3 is the dominant isoform in the pancreas (21, 44, 47). Another study found that decreased L-type channel activity and expression were associated with type 2 diabetes (50). Non-DHP-sensitive calcium channels have also been shown to play a role in insulin secretion. Studies have shown that Cav2.1 (33) and Cav2.3 (31, 45, 63) play a role in insulin secretion in INS-1 cells. Additionally, Cav2.2 has been found to play a role in {beta}-cell calcium current (1, 37) and shown to be linked with insulin secretion in RINm5F cells (51). However, Davalli et al. (17) found that the Cav2.2 antagonist, {omega}-cGVIA, had no effects on glucose stimulated insulin secretion in human islet cells.

Regardless of which HVA calcium channels are preferentially coupled to insulin secretion, maintenance of regulated calcium influx remains vital for normal {beta}-cell activity and survival. The HVA channel class may play an important role in the maintenance of calcium homeostasis in insulin-secreting cells in addition to providing localized calcium influx for vesicular secretion. Several studies have shown that a variety of cell types, including pancreatic {beta}-cells, have abnormalities with calcium homeostatic mechanisms in many diabetic and hyperglycemic models (8, 25, 32). We (28, 54) previously reported that changes in intracellular calcium can modulate HVA calcium current and that calcium influx can be attenuated under increased intracellular basal calcium conditions and augmented under decreased intracellular basal calcium conditions. Additionally, Cav1.3 translocation was demonstrated under changing calcium conditions. Because several studies have found that elevated intracellular calcium in {beta}-cells is associated with diabetes (8, 25, 32), calcium-induced channel migration may play a role in glucose-induced desensitization.

Overstimulation has been implicated as a possible cause of increased intracellular calcium in {beta}-cells (24). This would most likely involve L-type calcium channels. The L-type-selective antagonist nifedipine has been shown to normalize intracellular calcium as well as restore glucose sensitivity in mouse {beta}-cells (41). More recently, low-voltage-activated (LVA) T-type calcium channels have also been shown to play a role in enhanced excitability of {beta}-cell insulin secretion (6). Additionally, glucose has been shown to increase expression of these LVA channels (34), and data suggest that these channels mediate sustained increases in intracellular calcium (12, 15, 23, 65). However, the exact role of these channels in {beta}-cell physiology remains to be determined.

To assess the role of overstimulation in increased basal calcium, we examined the effects of chronic elevated glucose on basal calcium in the presence of nifedipine (L-type antagonist) or mibefradil or NNC-55-396 (selective LVA antagonists). Mibefradil has been found to inhibit both LVA current (IC50 = 0.7 µM) and L-type current (IC50 = 2 µM) in myoblasts (36), whereas NNC-55-396 has been found to block LVA current, with an IC50 = 7 µM having no activity on HVA current at doses up to 100 µM (29). Additionally, we set out to examine the effects of elevated glucose on HVA gene expression and investigate the effects of selective HVA channel antagonists on calcium influx and calcium current. To further characterize the role of HVA subtypes in {beta}-cell function, we examined insulin secretion at distinct stages of glucose stimulation in the presence of selective channel antagonists.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture, islet isolation, and drug treatment. INS-1 cells were cultured in RPMI 1640 medium (Life Technologies) containing 10% fetal bovine serum (FBS), 100 U/ml penicillin, 100 mg/ml streptomycin (P/S), and 50 µM 2-mercaptoethanol (3) in a humidified atmosphere of 5% CO2 at 37°C. Culture medium was replaced every 2–5 days. After cells were grown to confluence, they were treated with trypsin and 1 mM EDTA for 5 min (diluted 1:3) and replated on 75-cm2 flasks or 35-cm2 culture dishes.

For islet isolation, Sprague-Dawley rats (Charles River Laboratory, Wilmington, MA) were anesthetized with pentobarbital sodium (1 mg/kg). The pancreas was exposed by a midline abdominal incision. The pancreatic duct was exposed and canulated with a thin tube connected to a 10-ml syringe that contained a digestion solution (Hanks' solution; Life Technologies, Rockville, MD) containing collagenase (4 mg/ml; Boehringer Mannheim, Indianapolis, IN), DNase I (10 mg/ml; Sigma, St. Louis, MO), CaCl2 (1.28 mM), and bovine serum albumin (1 mg/ml, Life Technologies). The pancreas was slowly inflated with the digestion solution and removed and cut into small pieces with a pair of surgical scissors and placed into a 50-ml conical tube containing fresh digestion solution. The pancreatic tissue was incubated at 37°C for 20 min and then washed five times with enzyme-free Hanks' solution. Islets were picked up by hand under a x20 stereomicroscope and treated with 0.25% trypsin-EDTA (Sigma) for 5 min at 37°C, 5% CO2, in humidified atmosphere. Isolated cells were obtained by triturating the islets with plastic pipette tips and vortexing for 3 s. Cells were then transferred into 35-mm culture dishes and cultured in RPMI 1640 medium containing 5 mM glucose, 10% FBS, and P/S at 37°C, 5% CO2, in humidified atmosphere for 2–5 days before experiments.

For all experiments using HVA antagonists the following doses were used: 10 µM nifedipine, 1 µM N-type antagonist {omega}-conotoxin GVIA ({omega}-cGVIA), and 1 µM P/Q antagonist {omega}-conotoxin MVIIC ({omega}-cMVIIC, Sigma). Doses for mibefradil and NNC-55-396 (Novo Nordisk, Bagsvaerd, Denmark) were 1 µM. Because glucose desensitization can occur in as little as 4–6 h (49), our chronic elevated glucose treatments were done over a 24-h period to maximize the effects of glucose on calcium homeostasis.

Patch clamp electrophysiology. Recordings were carried out using gigaseal whole cell patch methods. Pipette resistance was in the range of 2–5 M{Omega} in our intracellular pipette solutions. An EPC-9 patch-clamp amplifier (HEKA, Gottingen, Germany) filtered at 2.9 kHz was used, and data were acquired using Pulse/PulseFit software (HEKA). Voltage-dependent currents were corrected for linear leak and residual capacitance by use of an on-line P/n subtraction. Following whole cell access, the cells were held at –70 mV with test pulses ranging from –60 to +40 mV with 10-mV increments. For the normalized current recordings, peak currents were measured at +10 mV. Extracellular bath solutions contained (in mM) 10 CaCl2, 110 tetraethylammonium-Cl, 10 CsCl, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), 40 sucrose, and 0.5 3,4-diaminopyridine at pH 7.3. The intracellular pipette solutions contained (in mM): 130 N-methyl-D-glucamine, 10 EGTA, 15 HEPES, 2 MgCl2, 10 CsCl, 5 NaCl, and 1.2 mM CaCl2 at pH 7.4.

Calcium imaging. INS-1 cells were plated onto 25-mm glass coverslips (Fisher, Pittsburgh, PA). Cytosolic Ca2+ concentration ([Ca2+]i) was estimated using the Ca2+-sensitive fluorophore fura 2. The measurement solution contained (in mM) 125 NaCl, 5.9 KCl, 1.28 CaCl2, 1.2 MgCl2, 17 HEPES, and 3.3 glucose, with pH adjusted to 7.4 with NaOH. Cells were incubated with fura 2-AM for 30 min (37°C in 5% CO2 in humidified atmosphere) in a loading solution that contained the measurement solution plus 3 µM fura 2-AM and 3 µl of a 10% pluronic acid solution. After fura 2 loading, cells were washed twice and incubated in a fura-free measurement solution for deesterification for an additional 20 min. After deesterification, calcium entry was assessed using a depolarizing solution that contained (in mM) 85 NaCl, 50 KCl, 2 CaCl2, 1 MgCl2, 15 HEPES, and 3.3 glucose adjusted to 7.4 pH with NaOH. Experiments were carried out using an inverted microscope in conjunction with a 175-W Xenon arc lamp Metafluor Imaging System (Nikon Instrument, Lewisville, TX). Switching of excitation wavelengths (340 and 380 nM) was controlled by a computer-driven filter wheel. The emitting fluorescent signal (510 nM) was acquired by an ADC 20-MHz camera (Photometric Coolsnap FX Monochrome, Nikon). Acquisition time per image was 250 ms, and the empirical Kd obtained for Ca2+ binding to fura 2 in our system was 269 nM. Data were acquired using MetaFluor 5.0R3 on an OEI (Konica) P4 1400 MHz processor with 1-GB RAM using Windows 2000 Professional SP3.

RNA isolation/real-time quantitative RT-PCR. Total RNAs were prepared from INS-1 or pancreatic islet cells by a modified method of Chomczynski and Sacchi (16) using TRIzol reagent (Life Technologies, Gaithersburg, MD) according to the manufacturer's instructions. Real-time quantitative (Q)RT-PCR with SYBR Green detection was performed using GenAmp PCR system 9600 and GeneAmp Sequence Detection System version 1.3. Primers were designed using ABI Primer Express 2.0. Total RNA was treated with a DNAse (Promega, Madison, WI) prior to cDNA production using the High Capacity cDNA Kit (Applied Biosystems/ABI Prism). Reaction conditions for cDNA synthesis were 80°C for 3 min and 37°C for 2 h.

Fifty nanograms of total RNA were used for each PCR reaction using the SYBR Green PCR Master Mix (Applied Biosystems) with 26-µl reaction volumes. The reaction conditions for PCR were (stage 1, Rep 1x) 50°C for 2 min, (stage 2, Rep 1x) 95°C for 10 min, and (stage 3, Rep 40x) 95°C at 15 s and then 60°C at 1 min. Sequences for real-time PCR primers are shown in Table 1.


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  		Table 1. HVA primer sets for quantitative real-time PCR

 
PCR products from unsorted islets cells cannot be attributed to {beta}-cells alone. Even though islets contain ~90% {beta}-cells, islets also contain {alpha}-, {delta}-, and PP-cells. These cells constitute the remaining 10% of cells in pancreatic endocrine tissue. INS-1 cells, however, are a glucose-responsive, homogeneous population of rat insulinoma {beta}-cells. The common link between INS-1 cells and other insulinoma cells used in similar studies is the fact that they are all glucose-responsive, insulin-secreting cells.

The mean CT values of the replicate wells run for each sample were calculated (n = independent experiments each run in triplicate). The difference ({Delta}CT) between the mean CT values of the samples in the target wells and those of the endogenous controls ({beta}-actin) was calculated. Then the difference between the {Delta}CT values of the samples for each target and the mean CT value of the calibrator for that target ({Delta}{Delta}CT) was calculated. The relative quantitation value was calculated using the equation 2= fold difference.

Insulin secretion. INS-1 cells were plated onto 24-well plates at a density of 10,000 cells/well and incubated in normal medium for 2 days. Before treatments, cells were washed with Hanks' balanced salt solution (HBSS, 0.1% BSA) with 0 mM glucose for 10 min. Groups were then treated with HBSS with 0 or 16 mM glucose for 40 min. For each time period, 10 µl of solution were aspirated and placed on ice before being subjected to the insulin secretion assay. Insulin was assessed using a High Range Insulin ELISA (ALPCO). After treatment, samples from each well were pipetted into an ELISA 96-well plate in triplicate. The anti-insulin conjugate solution was then added, and the plate was rotated at 900 rpm for 2 h at room temperature. After 2 h, 3,3',5,5'-tetramethylbenzidine (TMB) peroxidase substrate solution was added and incubated for 15 min at room temperature in the dark. Stop buffer was added, and the plate was rotated again for 5 s to allow mixing, and then the absorbance was measured at 450 nM with a MultiSkan ELISA plate reader.

Data analysis. All results are expressed as means ± SE. Statistical analysis was performed using a two-tailed unpaired t-test for comparisons between two groups, and P values of ≤0.05 were considered significant. Data were analyzed using GraphPad Prism version 4.0. Statistical significance for QRT-PCR experiments was performed on {Delta}CT values before calibrator genes were set to 1 (subsequent {Delta}{Delta}CT and FD ± SE values were calculated for each step).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
{beta}-Cells and HVA expression. To determine which HVA calcium channels are expressed in pancreatic {beta}-cells, we used a QRT-PCR method to show HVA mRNA expression in isolated rat islets and INS-1 cells. These data are represented primarily to show HVA channel mRNA expression. The SYBR Green {Delta}{Delta}CT method was used to calculate the relative fold difference of the HVA genes Cav1.2, Cav1.3, Cav2.1, and Cav2.2. Figure 1A shows the relative fold difference for the four HVA genes in INS-1 cells. After selecting the gene with highest {Delta}CT as a calibrator, the {Delta}{Delta}CT can be calculated for each gene. As shown, Cav2.2 has the lowest expression and its mean has been set to 1 ({Delta}CT = 7.61 SE ± 0.07, n = 5). Relative expression for Cav2.1 was 1.10x ± 0.08 ({Delta}CT = 7.59 SE ± 0.20, n = 5) and is not significantly higher than the calibrator, Cav2.2. Cav1.2 has a relative expression of 6.12x ± 1.13 ({Delta}CT = 4.99 SE ± 0.28, n = 5) and is significantly higher than Cav2.1, Cav2.2, and Cav1.3 (P < 0.05). Cav1.3 has a relative expression of 3.05x ± 0.28 ({Delta}CT = 6.00 SE ± 0.12, n = 5) and is significantly greater than the calibrator (P < 0.05).



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  		Fig. 1. Fold differences for high-voltage-activated (HVA) mRNA from quantitative (Q)RT-PCR. A: fold differences in HVA mRNA expression from INS-1 cells. B: data from isolated rat islets. *P < 0.05; n = 5–6.

 
Isolated pancreatic islet cells were also subjected to QRT-PCR analysis to test for HVA gene expression. Fold differences are shown in Fig. 1B for isolated islets cells. Cav2.2 had the lowest expression and was chosen as the calibrator for the fold difference calculation ({Delta}CT = 1.59 SE ± 0.41, n = 3). Cav2.1 ({Delta}CT = 0.55 SE ± 0.17, n = 3) and Cav1.2 ({Delta}CT = 0.41 SE ± 0.15, n = 3) were both significantly different from the calibrator and Cav1.3 ({Delta}CT = 1.30 SE ± 0.22, n = 3; P < 0.05). Fold difference for HVA genes: Cav2.1 (FD = 2.06x, SE ± 0.06), Cav1.2 (FD = 2.27x, SE ± 0.04), Cav1.3 (FD = 1.22x, SE ± 0.02).

PCR products from QRT-PCR experiments were run on a 3% agarose gel for product verification. Figure 2 shows HVA products from both INS-1 and isolated islets. Figure 2A, lanes 1–4 (Cav2.1, Cav1.2, Cav1.3, and Cav2.2, respectively), represents INS-1 HVA products, and lane 5 represents the internal standard, {beta}-actin. Figure 2B, lanes 1–4 (Cav2.1, Cav1.2, Cav1.3, and Cav2.2, respectively), represents HVA gene products from islets, and lane 5 represents the internal standard {beta}-actin. Lane 6 represents a 25-bp DNA ladder in both A and B. All primers for HVA genes and internal standards are ~75 bp in length and were designed with ABI Primer Express 2.0.



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  		Fig. 2. PCR products from QRT-PCR run on a 3% agarose gel for product verification. A: HVA mRNA from INS-1 cells (lanes 1–4: Cav2.1, Cav1.2, Cav1.3, and Cav2.2, respectively). B: HVA mRNA from islet cells (lanes 1–4: Cav2.1, Cav1.2, Cav1.3, and Cav2.2, respectively). Lane 5 in A and B represents the internal standard {beta}-actin; lane 6 in A and B represents 25-bp DNA ladder.

 
Using QRT-PCR, we examined the effects of elevated glucose on mRNA expression of Cav1.2, Cav1.3, Cav2.1, and Cav2.2 after a 24-h incubation period. Figure 3 shows fold differences for all four genes compared with low-glucose controls. Fold differences for each HVA gene were calculated on the basis of a comparison with the {Delta}CT for each low-glucose control. Statistical significance was calculated on the basis of the {Delta}CT values. Cav2.1 had an FD = 1.13 ± 0.035 ({Delta}CT = 5.70 ± 0.04, n = 6) and was not significantly different from the low-glucose control ({Delta}CT = 5.51 ± 0.07, n = 6). However, elevated glucose resulted in a significant decrease in Cav2.2 expression, with an FD = 0.54 ± 0.05 ({Delta}CT = 7.78 ± 0.05, n = 6; P < 0.01) vs. controls ({Delta}CT = 6.90 ± 0.05, n = 6). The Cav1.2 high-glucose group also had significantly lower expression than that of the low-glucose control group ({Delta}CT = 3.70 ± 0.15) with an FD = 0.32 ± 0.10 ({Delta}CT = 5.33 ± 0.10, n = 6; P < 0.05). Additionally, Cav1.3 mRNA was significantly lower with an FD = 0.50 ± 0.02 ({Delta}CT = 10.02 ± 0.023, n = 6; P < 0.05) vs. controls ({Delta}CT = 9.0 ± 0.049).



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  		Fig. 3. Fold difference results from QRT-PCR for all 4 HVA genes after 24-h treatment with 27 mM glucose (GLU; INS-1 cells). {alpha}1A, Cav2.1; {alpha}1C, Cav1.2; {alpha}1D, Cav1.3; {alpha}1B, Cav2.2. *P < 0.01, **P < 0.05, ***P < 0.001; n = 5–6.

 
High glucose and basal intracellular calcium. We wanted to test whether chronic high glucose had an effect on intracellular calcium. Figure 4A shows various concentrations of glucose treatment over a 24-h period. Basal calcium, as measured by fura 2 fluorescence, for the control (3.7 mM glucose) group was 80 nM (SE ± 4.0, n = 18). The group treated with 11.1 mM glucose had an intracellular calcium of 95 nM (SE ± 5.5, n = 18; P < 0.05) and was significantly different from the control. The other two treatment groups represented high-glucose treatment with 27 and 33.3 mM glucose. The 27 mM glucose treatment group had an intracellular calcium of 130 nM (SE ± 4.4, n = 54), whereas the 33.3 mM treatment group had an intracellular calcium of 235 nM (SE ± 11.8, n = 32), both of which were significantly different from the control (P < 0.001). To test whether the increased calcium was due to an osmotic effect, we incubated INS-1 cells with 33 mM mannose. The group treated with mannose had an intracellular calcium of 78.0 nM (SE ± 3.3, n = 18) that was not significantly different from that of the control glucose group. We then wanted to test whether the glucose effect on intracellular calcium was mediated, at least in part, by overstimulation of voltage-gated calcium channels.



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  		Fig. 4. Effects of 24-h glucose treatment on intracellular calcium in INS-1 cells. A: 4 glucose concentrations (3.7, 11, 27, and 33 mM) vs. basal calcium. *P < 0.05, **P < 0.0001. Mannose (33 mM) was used as an osmotic control. B: 24-h high-glucose treatment vs. basal calcium in the presence of nifedipine, mibefradil, or NNC-55-396 (NNC). *P < 0.001; n = 18.

 
To determine whether increased calcium was due to voltage-gated calcium channels, we incubated cells for 24 h with 33 mM glucose with nifedipine (10 µM), mibefradil (1 µM), or NNC-55-396 (4 µM). Mibefradil and NNC-55-396 are low-voltage-activated (LVA) calcium channel blockers. Mibefradil has been found to also block HVA calcium current (67), whereas NNC-55-0396 has been shown to be more selective for LVA calcium channels (29). As stated above, LVA calcium channels have been found to be increased during chronic high glucose and also play a role in increased basal calcium (12, 15, 23, 65, 70). Figure 4B shows the high-glucose control with 235 nM intracellular calcium (SE ± 11.8, n = 32). Cells that were treated with mibefradil had significantly lower calcium (140 nM, SE ± 21.4, n = 12) than controls (P < 0.001). Additionally, cells that were treated with NNC-55-396 had significantly lower calcium (135 nM, SE ± 6.8, n = 12) than controls (P < 0.0001). The HVA calcium channel blocker nifedipine also had significantly lower calcium (105 nM, SE ± 2.2, n = 12) than the high-glucose/no-drug controls (P < 0.0001). These data suggest that HVA, and possibly LVA, calcium channels are involved with increased basal calcium. However, the link between LVA calcium channels and changes in basal calcium needs to be investigated further.

{beta}-Cells and calcium current. Because the majority, but not all, of {beta}-cell calcium influx elicited by depolarization is sensitive to block by nifedipine (28), we wanted to determine the role of other HVA channel blockers on depolarization-induced calcium influx and calcium current. We used fura 2 fluorescence and whole cell patch clamp to test the effects of the following blockers on calcium current/influx: nifedipine, {omega}-cGVIA, and {omega}-cMVIIC. Figure 5A shows current-density means from whole cell patch clamp using different calcium channel blockers. Current-density (C/D) is calculated by dividing the current (picoAmps) of each cell by its membrane capacitance (picoFarads). Currents represent peak currents obtained with a test pulse of +10 mV and a holding potential of –70 mV. Figure 5A shows the mean C/Ds for all three calcium channel blockers. The control group has a C/D of 16.0 pA/pF (SE ± 2.2, n = 10). Peak current amplitudes for the Cav2.2 blocker {omega}-CGVIA were significantly smaller compared with control with a C/D of 10.1 pA/pF (SE ± 0.98, n = 10) and a P value of <0.05. The L-type antagonist nifedipine was also significantly smaller than controls, with a C/D of 9.23 pA/pF (SE ± 0.93, n = 10) and a P value of <0.01. However, the Cav2.1 blocker {omega}-cMVIIC was not significantly different from controls, with a 14.50 pA/pF (SE ± 2.34, n = 10) and a P value of >0.05.



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  		Fig. 5. A: current-density plots from patch clamp experiments using various calcium channel blockers. B: depolarization-induced (50 mM KCl) calcium influx data using fura 2 fluorescence in the presence of HVA antagonists. MVIIC, {omega}-conotoxin MVIIC; GVIA, {omega}-conotoxin GVIA; NIF, nifedipine. Data are represented as means ± SE. *P < 0.01, **P < 0.05, ***P < 0.0001; n = 9.

 
We also examined the effects of these calcium channel inhibitors on depolarization-induced calcium influx as measured by fura 2 calcium imaging (Fig. 5B). Depolarization was induced by perfusion of an osmotically balanced 50 mM KCl solution. Calcium influx for the control group was 721.8 nM (SE ± 36.1, n = 10). Calcium influx in the presence of the Cav2.1 inhibitor was significantly lower (580.6 nM, SE ± 16.0, n = 10) than the control group, with a P value of <0.01. Calcium influx in the presence of the Cav2.2 inhibitor was also significantly lower (569.4, SE ± 45.6, n = 11) than controls, with a P value of <0.05. As expected, calcium influx in the presence of the L-type channel blocker was significantly lower (205.8, SE ± 38.4, n = 8) than that of the control group, with a P value of <0.0001.

HVA channel subtypes and calcium current run-up. We (28) previously reported that HVA calcium current gradually increases "run-up" when measured using whole cell patch clamp and that this increase is due, at least in part, to L-type calcium channels. In the present study, we wanted to examine whether other HVA channels are involved in calcium current run-up.

Previous studies have implicated Cav1.2, Cav1.3, Cav2.1, and Cav2.2 calcium channels in run-up, whereas Cav2.3 has been shown to play a role in run-down (13, 19, 38, 62). We wanted to test which HVA channels are involved in run-up in {beta}-cells by using whole cell patch clamp in the presence of HVA calcium channel antagonists {omega}-cGVIA (Cav2.2), {omega}-cMVIIC (Cav2.1), and nifedipine (Cav1.2 and Cav1.3). Figure 6 shows normalized calcium current after the addition of HVA channel blockers. Figure 6A shows normalized currents with a test pulse of +10 mV elicited every 30 s. Inverted triangles represent six individual recordings after {omega}-cGVIA (1 µM) application and shows that run-up current is partially inhibited. We also used {omega}-cGVIA (1 µM) plus nifedipine (10 µM) (diamonds), which completely inhibited run-up in these cells (n = 6). Figure 6A also shows that the Cav2.1 channel blocker {omega}-cMVIIC does not inhibit run-up current (upright triangles, n = 7). Figure 6B represents peak run-up currents measured at 180 ms from data in Fig. 6A after {omega}-cGVIA plus nifedipine (0.659, SE ± 0.12) subtraction. Normalized peak currents show that {omega}-cMVIIC (1.240, SE ± 0.98, n = 7; P > 0.05) was not significantly different compared with controls (solid squares; 1.257, SE ± 0.084, n = 7). The partial block of run-up current by {omega}-cGVIA (0.591, SE ± 0.12, n = 6; P < 0.05) was significantly inhibited compared with controls (1.257, SE ± 0.084, n = 7), and the subtracted data revealed that nifedipine treatment (0.750 ± 0.084, n = 6; P < 0.002) significantly inhibited peak run-up current compared with controls (1.257, SE ± 0.084, n = 7). Previously, nifedipine (10 µM) was also shown to partially inhibit run-up under similar conditions (17).



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  		Fig. 6. A: normalized calcium current using whole cell patch clamp in the presence of MVIIC ({blacktriangleup}), GVIA ({blacktriangledown}), and GVIA + nifedipine ({blacklozenge}) measured every 30 s. B: peak current measured at 180 s after GVIA + nifedipine subtraction. Data represented as means ± SE. *P < 0.05; n = 7–11.

 
HVA calcium channels and insulin secretion. It is generally accepted that the HVA L-type calcium channels are primarily associated with stimulus secretion coupling. However, other channel subtypes have been implicated in glucose-induced calcium influx (e.g., Cav2.3) (63). We examined glucose-induced insulin secretion in the presence of different HVA channel antagonists at various time points to determine which channels are involved in stimulus secretion coupling. Figure 7 shows insulin secretion from INS-1 cells measured at different time points. Figure 7A shows unstimulated basal insulin secretion measured at 1 min (6.3 ng/µl, SE ± 0.8, n = 3), 3 min (7.0 ng/µl, SE ± 2.0, n = 3), 10 min (11.1 ng/µl, SE ± 1.4, n = 3), and 40 min (12.1 ng/µl, SE ± 1.3, n = 3). Figure 7B represents insulin secretion measured after 1 min of 16 mM glucose exposure in the presence of HVA channel blockers. Insulin secretion in the presence of the Cav2.1 blocker {omega}-cMVIIC (23.0 ng/µl, SE ± 2.2, n = 6) was not significantly different from the 16 mM controls (22.5 ng/µl, SE ± 2.0, n = 6). Insulin secretion in the presence of the Cav2.2 blocker {omega}-cGVIA (10.5 ng/µl, SE ± 0.34, n = 6) was significantly lower than controls, with a P value of <0.01. Additionally, insulin secretion in the presence of the L-type blocker nifedipine was significantly lower (14.7 ng/µl, SE ± 1.8, n = 6) from the control group, with a P value of <0.05. Figure 7C represents cumulative (1 min + 10 min) insulin secretion measured after 10-min exposure to 16 mM glucose in the presence of antagonists. Again, the Cav2.1 blocker {omega}-cMVIIC did not significantly (39.5 ng/µl, SE ± 2.0, n = 6) inhibit insulin secretion compared with controls (39.0 ng/µl, SE ± 1.6, n = 6). The presence of the Cav2.2 blocker did not significantly lower (31.0 ng/µl, SE ± 4.7, n = 6) insulin secretion compared with controls. Insulin secretion in the presence of the Cav1.2 and Cav1.3 (L-type) blocker was significantly lower (28.4 ng/µl, SE ± 2.2, n = 6) than the control group, with a P value of <0.01. Measurement of insulin at a third time period (40 min) revealed similar results to the 10-min measurement. Figure 7D shows no significant inhibition of insulin secretion in the presence of {omega}-cMVIIC (62.9 ng/µl, SE ± 0.37, n = 6) or {omega}-cGVIA (55.0 ng/µl, SE ± 1.05, n = 6) compared with controls (65.5 ng/µl, SE ± 5.59, n = 6). As expected, the presence of nifedipine significantly inhibited insulin secretion (37.8 ng/µl, SE ± 1.48, n = 6) compared with controls, with a P value of <0.05.



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  		Fig. 7. A: basal insulin secretion from INS-1 cells. B: glucose-stimulated (16 mM glucose) insulin secretion in the presence of MVIIC, GVIA, and nifedipine after 1 min (0, no glucose). C: cumulative insulin secretion from 10 min. D: insulin secretion from 40 min. Data represented as means ± SE. *P < 0.05; n = 6.

 
These data suggest that Cav2.2 (N-type) in addition to Cav1.2 or Cav1.3 (L-type) plays a role in the immediate response to glucose. However, N-type blockade did not provide any additional inhibition of insulin secretion after early stimulation as did L-type inhibition.


   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Expression of HVA calcium channels. Almost all of the HVA genes have been found to be expressed in rat and human pancreatic {beta}-cells (27, 30, 33, 43, 58, 63, 64). However, the role of each channel subtype in these cells remains unclear. All of these channels are capable of being activated by the depolarization elicited by glucose metabolism, and most studies conclude that L-type calcium channels are primarily involved in insulin secretion from {beta}-cells. It is possible that these channels are selectively activated due to close proximity with KATP channels or insulin-secretory vesicles. This close association would allow for localized depolarization with subsequent localized calcium increases. Several studies have shown that calcium channels are colocalized with vesicles in neural tissue (18, 57, 59), which may also be the case in neuroendocrine tissue. In our study, we wanted to first determine the level of expression of four HVA calcium channel genes. The genes that we probed for were the Cav1.2, Cav1.3, Cav2.1, and Cav2.2 HVA calcium channels. We found that all four HVA calcium channels' mRNA were present in both INS-1 and isolated rat islet cells. In islet cells, quantitative analysis of mRNA revealed no significant differences for all four genes. This suggests that, at the message level, these genes are equally expressed. In INS-1 cells, the mRNA level for the four genes was not found to be equally expressed. Our data suggest that Cav1.2 mRNA is expressed at a much higher level under our culture conditions.

Pancreatic {beta}-cells share some common features with neurons, including expression of channels and proteins involved with synaptic vesicle secretion (55). Cav2.1 and Cav2.2 have been found to be associated with glutamate and GABA activity in neuronal tissue (10, 52). Many GABA and glutamate receptor subtypes have also been found to be expressed in pancreatic {beta}-cells (11), and glutamate receptors are known to facilitate neuronal GABA release (53). It is possible that Cav2.1 or Cav2.2 is involved with glucose- or glutamate-stimulated GABA release from pancreatic {beta}-cells. Once released, GABA then plays a crucial role in regulating {alpha}- and {beta}-cell activity (66).

Glucose and intracellular calcium. It is commonly known that application of high concentrations of glucose will induce calcium entry in pancreatic {beta}-cells. Glucose is taken in by the {beta}-cells via the GLUT2 transporter and metabolized. Metabolism of glucose leads to changes in the ATP/ADP ratio, resulting in closure of KATP channels and subsequent depolarization. Depolarization leads to activation of HVA and LVA calcium channels and calcium influx. As illustrated in Fig. 8, elevated calcium levels resulting from calcium channel activation can then lead to a decrease of HVA calcium channel expression (41) and internalization of these channels (28). Additionally, elevated glucose increases LVA calcium channel gene expression, resulting in a sustained increase in basal calcium. LVA calcium channel may regulate calcium influx by itself (e.g., window current) or in a synergistic manner with the HVA calcium channels. This model can be used to explain our results that both HVA and LVA channel blockers can partially prevent the elevation of basal calcium induced by chronic high-glucose treatment.



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  		Fig. 8. Illustration of a hypothetical model of the regulation of intracellular basal calcium concentration via a mechanism involving glucose-modulated HVA and LVA Ca2+ channel expression. KATP, ATP-regulated potassium channels. HVA, high-voltage-activated Ca2+ channels; LVA, low-voltage-activated Ca2+ channels; {Delta}{psi}m, change in plasma membrane potential; extra, extracellular side of plasma membrane; intra, intracellular side of plasma membrane; Ch(T), LVA; Ch(L), HVA.

 
{beta}-Cells and HVA calcium current/influx. Because the majority, but not all, of depolarization-induced calcium influx is attributed to L-type calcium channels, we wanted to determine the role of other HVA calcium channels in calcium influx. Whole cell patch clamp studies showed that a small portion of calcium current is attributed to Cav2.2 and Cav2.1 on the basis of channel selective toxins. Any remaining HVA calcium current was attributed to Cav2.3, which is resistant to HVA antagonists. However, it has been recently shown that SNX-482 inhibits Cav2.3 current (4), but the role of these channels in {beta}-cells has yet to be determined.

Our results revealed that Cav2.2 as well as L-type channels are involved in calcium current run-up, whereas Cav2.1 were not found to play a role in increased calcium current. We believe that the run-ups of Cav2.2 and L-type current are primarily due to channel migration to the plasma membrane as a result of lowered intracellular calcium, primarily because this effect is seen only when calcium chelators are included with the intracellular solution. This conclusion is supported by our previous demonstration of L-type channel migration under changing calcium concentrations by use of immunofluorescence microscopy (28). Previous studies have found that Cav2.3 do not play a role in run-up but may be involved in run-down (decreased calcium current following repeated stimulation). The mechanism behind run-down is unknown, but it is clear that all HVA calcium channels are capable of exhibiting run-down in whole cell patch clamp experiments.

HVA channels and insulin secretion. The role of L-type channels was found to be consistent with previously reported studies showing that DHPs inhibit not only the majority of calcium current and calcium influx but also insulin secretion (9, 28, 40). Interestingly, it was found that Cav2.2 are primarily involved with immediate insulin secretion that occurs in less than 1 min, as shown in Fig. 7B. However, Cav2.2 inhibition did not provide any additional block of insulin secretion after 1 min (Fig. 7, C and D). One important factor in type 2 diabetes is the loss of the immediate first phase of insulin secretion. This effect was found to be reversible when blood glucose levels were returned to normal (2, 48). Cav2.2 may play an important role in the loss of the early glucose response seen in type 2 diabetics. However, at later stages, it appears that L-type channels are primarily linked with insulin secretion. Even though Cav2.1 were also found to be expressed in INS-1 and islet cells, it appears that they may play little or no role in insulin secretion under normal conditions. These channels, however, are known to be involved in calcium influx for GABA release in neurons (61) and may be involved in localized calcium influx associated with GABA vesicles in insulin secreting cells.

Having a multiple array of voltage-gated calcium channels in pancreatic {beta}-cells may allow these cells to fine-tune insulin-secretory efficiency. In a disease like diabetes, there may be a defect in the cells' ability to maintain this delicate balance. If selective HVA channel translocation to and from the plasma membrane is a process that can be regulated by glucose or basal calcium, then this may be an important factor when one considers {beta}-cell activity in the diabetic state.


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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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This work was supported by American Diabetes Association Research Award 7-03-RA-51 to M. Li.


   FOOTNOTES
 

Address for reprint requests and other correspondence: Ming Li, Pharmacology SL83, 1430 Tulane Ave., New Orleans, LA 70112 (e-mail: mli{at}tulane.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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Channel regulation of glucose sensing in the pancreatic {beta}-cell
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M. D. Nitert, C. L F Nagorny, A. Wendt, L. Eliasson, and H. Mulder
CaV1.2 rather than CaV1.3 is coupled to glucose-stimulated insulin secretion in INS-1 832/13 cells
J. Mol. Endocrinol., July 1, 2008; 41(1): 1 - 11.
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V. Navarro-Tableros, T. Fiordelisio, A. Hernandez-Cruz, and M. Hiriart
Physiological development of insulin secretion, calcium channels, and GLUT2 expression of pancreatic rat beta-cells
Am J Physiol Endocrinol Metab, April 1, 2007; 292(4): E1018 - E1029.
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