1Departments of Pharmacology, Tulane University Health Sciences Center and 4Department of Genetics, Louisiana State University Health Sciences Center, New Orleans, Louisiana 70112; 2Department of Pharmacology, Yale University School of Medicine, New Haven, Connecticut 06520; and 3Department of Pharmacology, Medical College of Virginia, Richmond, Virginia 23298
Submitted 12 August 2003 ; accepted in final form 25 September 2003
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ABSTRACT |
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internalization; intracellular free calcium; glucose desensitization
Alternating levels of [Ca2+]i influence glucose-evoked insulin release. It has been reported that after chronic exposure to high glucose concentrations, pancreatic islets are severely impaired in their ability to secrete insulin during an acute glucose challenge. This is associated with an elevation in basal [Ca2+]i and the inability of glucose to provoke a Ca2+ influx (5, 18). These results suggest that regulation of basal [Ca2+]i also plays an important role in glucose-evoked insulin release.
The regulation and significance of basal [Ca2+]i in -cells have not been investigated in detail. It is known that glucose causes an initial decrease in [Ca2+]i before elevating [Ca2+]i through mechanisms possibly involving ATP-dependent removal of Ca2+ from the cytoplasm (9). The purpose of this initial lowering of [Ca2+]i still remains unclear.
We have observed that a significant proportion of the high-voltage-activated (HVA) Ca2+ currents is under substantial Ca2+-dependent downregulation in both rat primary cultured islet cells and the insulin-secreting cells, INS-1, because these currents exhibit a pronounced "run-up" when Ca2+ chelators are introduced inside the cell. It is generally accepted that Ca2+ channels sensitive to dihydropyridines (DHPs), L-type, contribute to most of the Ca2+ current required for insulin release and are ubiquitous in all insulin-secreting cells (6). The function of L-type Ca2+ channels has been further attributed to either the CaV1.2 (1C) or CaV1.3 (
1D), the pore-forming subunits of L-type Ca2+ channels in different preparations (3, 10, 21, 27). We hypothesized that, in rat pancreatic
-cells, changes in basal [Ca2+]i determined the total number of surface Cav1.3. Therefore, we decided to study the effects of basal [Ca2+]i on the HVA Ca2+ currents and the specific HVA Ca2+ channel protein translocation in rat insulin-secreting cells.
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MATERIALS AND METHODS |
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Islet cell preparation. Pancreas of Sprague-Dawley rats (Charles River Laboratory, Wilmington, MA) were removed and cut into small pieces with a pair of surgical scissors in Hanks' solution (Life Technologies, Rockville, MD) containing collagenase (4 mg/ml; Boehringer Mannheim, Indianapolis, IN), DNase I (10 µg/ml; Sigma, St. Louis, MO), CaCl2 (1.28 mM), and bovine serum albumin (1 mg/ml; Life Technologies). 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 manually under a x20 stereomicroscope and treated with 0.25% trypsin-EDTA (Sigma) for 5 min at 37°C and 5% CO2 in a 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 (Life Technologies) containing 5 mM glucose, 10% FBS, and P/S at 37°C, 5% CO2, in a humidified atmosphere for 25 days before experiments.
Patch-clamp electrophysiology. Whole cell voltage-clamp recordings were carried out by the suction pipette method. Pipette resistance was in the range of 25 M in our internal solutions. An EPC-9 patch-clamp amplifier (HEKA, Göttingen, Germany) filtered at 2.9 kHz was used, and data were acquired by using Pulse/PulseFit software (HEKA). Voltage-dependent currents have been corrected for linear leak and residual capacitance by using an online P/n subtraction paradigm. For intracellular perfusion experiments, perfusate was introduced into the interior of the pipette through a thin quartz capillary (25 µm in diameter, a kind gift from Dr. D. L. Armstrong at National Institute of Environmental Health Science, Research Triangle Park, NC) inserted within 100200 µm of the tip.
Solutions for recording. For whole cell voltage-clamp recordings, the extracellular bath solution contained (in mM) 10 CaCl2, 110 tetraethylammonium chloride (TEA-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) for "zero calcium": 130 N-methyl-D-glucamine (NMDG), 10 EGTA (free acid), 15 HEPES, 2 MgCl2, 10 CsCl, 5 NaCl; for 132 nM calcium solution: 130 NMDG, 2.15 EGTA (free acid), 1 CaCl2, 15 HEPES, 2 MgCl2, 10 CsCl, 5 NaCl; and for 284 nM Ca2+ solution: 130 NMDG, 1.4 EGTA, 1 CaCl2, 15 HEPES, 2 MgCl2, 10 CsCl, and 5 NaCl. All solutions were pH adjusted to 7.4 with methanesulfonic acid.
[Ca2+]i measurement with fura 2 fluorescence. Rat islet cells or INS-1 cells were seeded onto glass coverslips (Fisher, Houston, TX). [Ca2+]i was estimated using the Ca2+-sensitive fluorophore, fura 2. The measurement solution contained (in mM) 130 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, and 15 HEPES, with pH adjusted to 7.4 with NaOH. Cells were washed twice with measurement solution before fura 2-AM loading. The depolarizing solution contained (in mM) 85 NaCl, 50 KCl, 2 CaCl2, 1 MgCl2, and 15 HEPES, pH adjusted to 7.4 with NaOH. The loading solution (1 ml) consisted of the measurement solution plus 3 µM fura 2-AM and 3 µl of a 10% pluronic acid solution. Cells were fura 2-AM loaded for 20 min at 37°C, 5%CO2, in a humidified atmosphere. After this loading period, cells were again washed and incubated for deesterification for an additional 20 min. After deesterification, [Ca2+]i was assessed with an inverted microscope in conjunction with a 175-watt Xenon arc lamp Metafluor Imaging System (Nikon Instrument, Lewisville, TX). Switching of excitation wavelengths (340 and 380) was controlled by a computer-driven filter wheel. Emitting fluorescent signal was acquired by an ADC 20-MHz camera (Photometric Coolsnap FX Monochrome, Nikon). The acquisition time per data was analyzed with Metafluor software (Nikon). The acquisition times per image and per image pair are 100 ms and 250 ms, respectively. Fluorescence was measured from single cells, and the values in [Ca2+]i were determined as the fluorescence ratio (R) of the Ca2+-bound fura 2 (340 nm) to Ca2+-unbound (380 nm) excitation wavelengths emitted at 510 nm. R was converted to free Ca2+ concentration by using a standard curve that was generated with a calibration kit (Molecular Probes, Eugene, OR). The empirical dissociation constant (Kd) obtained for Ca2+ binding to fura 2 in our system was 307 nM.
Immunofluorescence microscopy. Immunofluorescence samples were prepared as described previously (16). Briefly, INS-1 cells were cultured on coverslips overnight, fixed in 4% paraformaldehyde (Polysciences, Warrington, PA) in PBS (Dulbecco's phosphate-buffered saline; Mediatech, Herndon, VA) for 30 min, permeabilized, and blocked in 5% normal goat serum in PBS for 30 min with 0.1% Triton X-100. The cells were simultaneously incubated for 2 h with rabbit anti-Cav1.3 (Calbiochem-Novabiochem, San Diego, CA) and mouse anticlathrin (Calbiochem-Novabiochem) primary antibodies at a 1:240 dilution each. The coverslips were washed well and simultaneously incubated with goat anti-rabbit IgG (H+L) conjugated to Oregon green (also referred to here as FITC) and goat anti-mouse IgG (H+L) conjugated to rhodamine for 1 h. All steps were performed at room temperature and protected from light. The coverslips were mounted on slides and analyzed on a Leica DMRXA automated upright epifluorescent microscope (Leica Microsystems, Bannockburn, IL) equipped with a x63 (NA 1.4) oil-immersion objective, a Sensicam QE charge-coupled device digital camera (Cooke, Auburn Hills, MI), and filter sets optimized for Oregon Green (exciter HQ480/20, dichroic Q495LP, and emitter HQ510/20 nm), as well as rhodamine (exciter x545/30, dichroic Q570DLP, and emitter HQ620/60 nm). z-Axis plane capture, deconvolution, and analysis were performed with Slidebook deconvolution software (Intelligent Imaging Innovations, Denver, CO).
Analysis involved the creation of a digital binary overlay by segmenting or filtering arbitrary areas over threshold from both green and red fluorescence channels within all captured planes (total of 68 per cell at 0.1 µm each) from cells demonstrating Oregon green indirectly labeled to anti-Cav1.3 primary antibody and constitutive anticlathrin indirectly labeled to rhodamine. Slidebook-driven mathematical voxel calculations were applied to 20 cells per group after a background subtraction and segmentation (masking) of fluorescent Cav1.3 and anticlathrin channels within all planes from each cell's z-axis stack. The anticlathrin channel served as a template for surface-specific labeling. Therefore, by using the anticlathrin mask and including the space within as an addition to the mask, volume calculations could be performed. Discrimination of anti-Cav1.3 membrane vs. intracellular expression was performed by establishing an anti-Cav1.3 mask based on the above threshold voxels colocalized with the anticlathrin mask. Subtracting the anticlathrin mask from the total cell volume mask rendered a representative mask for the intracellular signal. Mask segmentation through calculation of voxels above threshold and mask arithmetic operations were created by input of consistent intensity histogram values throughout samples and the Slidebook software. A "nearest neighbors" deconvolution was applied to a representative z-axis stack from each group for qualitative purposes only and after all quantitative data were obtained. Analysis also entailed a pixel correlation algorithm as a measure of colocalization (or lack thereof) between the FITC and rhodamine areas within each image (or plane) through thresholding and channel segmentation. This statistical measurement determines trends in the degree by which two pixels in an image correspond to one another. If the trends in number of pixels overlap or colocalize entirely, then they have a correlation measurement of 1.0 and are considered to be completely correlated. If the two sets of pixels correspond to each other in the exactly opposite manner, then they have a correlation measurement of 1.0 and are considered to be negatively correlated. Thresholding and channel segmentation were also utilized to measure the mean anti-Cav1.3-Oregon Green intensity in individual cells throughout their three-dimensional multiplane digital rendering. The thirty-fourth plane from a nearest neighbors deconvolved z-axis stack from each group was extracted for representative and qualitative purposes only (see Fig. 6A).
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Isolation of surface membrane protein and Western blot. The surface membrane protein was prepared as described previously (7). Briefly, INS-1 cells were washed twice with warm PBS and incubated in either PBS or PBS supplemented with 1 µM calcium ionophore A-23187 at 37°C, 5% CO2, in a humidified atmosphere for 30 min. The cells were washed twice with the warm PBS and incubated with the membrane-impermeable reagent N-hydroxysuccinimide-SS-biotin (EZ-Link Sulfo-NHS-SS-biotin; Pierce, Rockford, IL) at a concentration of 1.5 mg/ml in the PBS for 30 min at 4°C with gentle shaking. The cells were then washed twice with the PBS supplemented with 100 mM glycine and incubated with this solution for 30 min to remove any unbound NHS-SS-biotin. After being washed three times with the PBS, cells were scraped into 1 ml of ice-cold harvest buffer (1 mM EGTA, 1 mM EDTA, 200 µM phenylmethylsulfonyl fluoride, 1 µg/ml pepstatin A, 1 µg/ml leupeptin, 1 µg/ml aprotinin, 0.1 µg/ml benzamidine, and 8 µg/ml calpain inhibitor) into Eppendorf tubes prechilled on ice, sonicated (10 s), and centrifuged (14,000 rpm for 30 min at 4°C). The precipitates, which correspond to the membrane fraction, were redissolved by sonication (10 s) in solubilization buffer (harvest buffer plus 1% Triton X-100), followed by end-over-end mixing for 30 min in a 4°C cold room. After another centrifugation at 14,000 rpm for 30 min at 4°C, the supernatants were stored at 80°C for Western blot analysis. Protein concentrations were determined by Bio-Rad protein assay (Bio-Rad Laboratories, Hercules, CA). The antibody against Na+-K+-ATPase 1-subunit was purchased from Upstate Biotechnology (Lake Placid, NY).
Tissue preparation. One rat brain was removed and placed in 5 ml of ice-cold harvest buffer, homogenized in a PowerGen model 125 homogenizer (Fisher Scientific, Houston, TX), and centrifuged at 4°C at 14,000 rpm for 10 min. The pellet was resuspended in 2 ml of solubilization buffer, followed by end-over-end mixing for 30 min in a4°C cold room. After another centrifugation at 4°C at 14,000 rpm for 30 min, the supernatants were determined for protein concentration and stored at 80°C for Western blot analysis.
For analysis of Cav1.3 or Cav1.2 in total membrane expression, identical amounts (50 µg) of protein from the membrane fraction of each cell lysate were loaded to 7% SDS-PAGE. For the assessment of Cav1.3 surface expression, the remainders of each cell lysate were incubated with 100 µl (spun down from a 200-µl suspension) of NeutrAvidin-linked beads (Pierce) by end-over-end rotation for 2 h at 4°C. Beads were extensively centrifuged and washed with harvest buffer to remove unbound proteins. The surface membrane proteins were eluted by incubating the bead-bound proteins with dithiothreitol-containing SDS-PAGE loading buffer and loaded to 7% SDS-PAGE. After an hour-long transfer of gels to nitrocellulose membranes, the membranes were blocked with 5% skim milk and blotted with rabbit anti-Cav1.3 (Calbiochem-Novabiochem) or anti-Cav1.2 (Chemicon International, Temecula, CA) polyclonal antibody at 1.5 µg/ml concentration. The membranes were then incubated with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (KPL Europe, Guildford, UK) diluted in 1:2,000. Bands were visualized by enhanced chemiluminescence (ECL; Amersham Biosciences, Piscataway, NJ). Relative intensities of the bands were determined by a GS-700 imaging densitometer (Bio-Rad Laboratories).
Statistical analysis. All results were expressed as means ± SE. Statistical analysis was performed using two-tailed unpaired t-test for comparisons between two groups, and P values of 0.05 or less were considered significant.
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RESULTS |
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To elucidate the effect of changing basal [Ca2+]i on the regulation of HVA Ca2+ channels, we measured INS-1 cell voltage-dependent Ca2+ influx induced by replacing the measurement solution with an osmotically balanced depolarizing solution containing 50 mM KCl. The prestimulating basal [Ca2+]i was determined for each cell before the perfusion.
The 50 mM KCl perfusion induced an immediate increase in [Ca2+]i that decayed gradually (Fig. 2A). Again, the relationship between the basal [Ca2+]i and the Ca2+ influx stimulated by 50 mM KCl was determined by plotting Ca2+ influx (peak value of [Ca2+]i subtracted by the value of prestimulation basal [Ca2+]i) over the basal [Ca2+]i. As shown in Fig. 2B, the smaller Ca2+ influx was coupled to the higher basal [Ca2+]i. The slope of best-fit values is 3.155 ± 0.7771 and significantly deviated from zero (P < 0.001). Because influx was substantially reduced when the pretreated (and perfused) solution contained 10 µM nifedipine (Fig. 2A), the Ca2+ influx was mediated primarily by dihydropyridine-sensitive Ca2+ channels.
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We decided to characterize the effects of different [Ca2+]i on the Ca2+ currents with the combination of whole cell patch-clamp recording and intracellular perfusion techniques. Ca2+ currents were recorded at 0 mV when held at 70 mV. This protocol allowed us to record both T-type and HVA Ca2+ currents. In the cells we studied, however, the major currents were the HVA Ca2+ currents indicated by sustained activation of the currents during the 100-ms depolarization duration, as shown in Fig. 3A. After breaking in, the pipette solution entered the cell to set up a first controlled [Ca2+]i. This intracellular solution was then replaced by a second solution that contained different concentration of free Ca2+, which accordingly changed the [Ca2+]i in the recording cell. Intracellular perfusion of a solution containing high Ca2+ (284 nM) (Fig. 3A) caused a substantial reduction in the HVA Ca2+ current. This process was reversible, as shown in Fig. 3B. Perfusing a low [Ca2+]i solution to replace a high [Ca2+]i solution (284 nM 0 nM) caused an increase in the HVA Ca2+ current before current rundown, whereas perfusing a high [Ca2+]i solution to replace a low [Ca2+]i solution (0 nM
284 nM) reduced the HVA Ca2+ current over this time period. When an intermediate concentration of Ca2+ solution was perfused to replace a low Ca2+ solution (132 nM
0 nM), the HVA Ca2+ current remained relatively unchanged during this period. The free Ca2+ concentrations of intracellular perfusing solutions were approximately determined with Fabiato's equation (8) and then adjusted by direct measurement using fura 2 ratiometric fluorescence.
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The effects of a "high" [Ca2+]i (284 nM) on the current-voltage (I-V) relationship and the isochronic inactivation of the HVA Ca2+ current are shown in Fig. 3, C and D, respectively. The high [Ca2+]i caused shifts in HVA Ca2+ current activation and inactivation to negative voltages [by simulating the data with the Boltzmann equation, the values of half-inactivation voltage (V1/2) and slope (k) are changed from 45.6 ± 0.8 and 4.2 to 54.0 ± 0.3 and 4.0 for the 0 and 284 nM [Ca2+]i conditions, respectively; n = 3]. The high [Ca2+]i also caused a potentiation of the T-type Ca2+ current, as indicated by increased currents at 20 mV and lower voltages. The increase in T-type Ca2+ current under high [Ca2+]i conditions was further investigated by measuring the slow deactivating tail currents at 80 mV after stepping to 30 mV for 10 ms (Fig. 3E). The voltage-dependent activation of T-type Ca2+ channel currents shifted 11 mV to negative potentials with 284 nM Ca2+ perfusion as shown in Fig. 3F (V1/2 and k are changed from 17.1 ± 1.1 and 7.6 to 28.1 ± 0.7 and 6.2 for the 0 and 284 nM [Ca2+]i conditions, respectively; n = 3).
Our standard pipette solution contained a high concentration of Ca2+ chelators, and we frequently observed the HVA Ca2+ current increases over time (run-up) upon establishment of a whole cell patch (Fig. 4, A and B). We speculated that the run-up phenomenon was due to Ca2+ chelation inside the cells and not a phosphorylation-mediated event because the run-up took place whether or not 3 mM Mg-ATP or KN-62, a calmodulin kinase inhibitor, was included in the pipette solution (data not shown). When the intracellular solution contained a free Ca2+ concentration of 1 µM, the HVA Ca2+ current exhibited no initial run-up but, indeed, a rapid rundown (data not shown). We used run-up to further study the mechanism of basal [Ca2+]i-dependent regulation of HVA Ca2+ channel activity.
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Changes in the Ca2+ channel activity may result from changes in the opening probability of the channels or changes in total number of channels residing in the plasma membrane. The latter mechanism most likely occurs via a cytoskeleton-mediated channel migration between the plasma membrane and intracellular locations. To test the possibility of cytoskeleton-mediated channel transport mechanism, we preincubated INS-1 cells with cytoskeleton modulators, 30 nM cytochalasin B for 60 min, or 50 µM colchicine for 24 h and then examined the effects of these drugs on HVA Ca2+ current run-up. HVA Ca2+ current run-up was significantly prevented when cells were preincubated with cytochalasin B or colchicine (Fig. 4, C and D). At the 180-s point, the normalized amplitudes of HVA Ca2+ current recorded from the control cells, cytochalasin B-treated cells, and colchicine-treated cells were 1.916 ± 0.142 (n = 7), 0.7654 ± 0.0489 (n = 6), and 0.986 ± 0.155 (n = 6), respectively. These values are statistically different between the control and the cytochalasin B-treated groups (P < 0.0001) or the colchicine-treated groups (P < 0.0001) determined by an unpaired t-test. The viability of the cells after colchicine incubation was determined by using the trypan blue staining method, which showed that 98 ± 2% of the cells were viable (n = 4). These results suggest that depletion of [Ca2+]i and concomitant HVA Ca2+ current run-up involves a cytoskeleton-related process.
Because nifedipine attenuated 50 mM KCl-induced Ca2+ influx, we tested its effect on the HVA Ca2+ current run-up. We first conducted a set of experiments to characterize the dose-dependent effect of nifedipine on the total HVA Ca2+ current recorded in INS-1 cells. Figure 4E shows that 60% of HVA current was blocked by 10 µM nifedipine. When the bath solution contained 10 µM nifedipine, the run-up of HVA Ca2+ current was substantially reduced (Fig. 4F), indicating that dihydropyridine-sensitive Ca2+ channels are involved in [Ca2+]i-dependent HVA Ca2+ current regulation.
It has been suggested that Cav1.3 is the major component of HVA Ca2+ channels in rat pancreatic -cells and that it plays a crucial role in the regulation of insulin secretion in normal and altered metabolic states (10, 21). In addition, Cav1.3-mediated Ca2+ influx is preferentially coupled with glucose-stimulated insulin secretion in INS-1 cells (15). We decided to use a Cav1.3-specific antibody to assay the translocation of Cav1.3 channels between the plasma membrane and cytoplasm. In Fig. 5A, using the anti-Cav1.3 antibody, a band of
240 kDa was detected by Western blot. This band was absent when the antibody had undergone pretreatment with Cav1.3-specific inhibitory peptide. Thus this validates the antibody's specificity for the Cav1.3 protein. We also performed a similar analysis to determine Cav1.2 expression in INS-1 cells. As shown in Fig. 5B, Cav1.2 was not detected in INS-1 cells, whereas a Cav1.2 (
210 kDa)-positive band was shown in a rat brain preparation. The band of Cav1.2 was absent in rat brain preparation when the anti-Cav1.2 antibody was preincubated with Cav1.2-specific inhibitory peptide.
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Both plasma membrane and intracellular expression of the Cav1.3 protein in INS-1 cells were analyzed by using indirect immunofluorescence and epifluorescence microscopy of anti-Cav1.3, which is visualized as green (Fig. 6A). The red channel represents the coimmunofluorescent labeling of clathrin. Under the control condition, in which cells were incubated in PBS for 30 min, the majority of Cav1.3 proteins were located within or adjacent to the plasma membrane. Because the intracellular perfusion and run-up experiments indicated a fast increase in the HVA current amplitude as the intracellular Ca2+ was depleted, it is likely that the channels were in close proximity to the plasma membrane. In contrast, cells incubated with PBS plus 1 µM calcium ionophore A-23187 for 30 min showed an increased cytoplasmic distribution of Cav1.3. No green fluorescent signal was identified within cells in which the Cav1.3 antibody was omitted, indicating that secondary antibodies are specific to the Cav1.3 antibodies.
Red fluorescence represents the location of clathrin, which outlines the plasma membrane of the cell. By subtracting the Cav1.3 (green)- and clathrin (red)-colocalized pixels from the total visible pixels within each cell's volume, we have estimated the relative distribution of Cav1.3 under the condition of the control and high [Ca2+]i. Intensity measurements depict a clear cytoplasmic distribution increase as represented by the mean anti-Cav1.3 intensities recorded within projection planes composing the z-axis stack from cells under the high [Ca2+]i condition (n = 20) (Fig. 6B). The plasma membrane fraction is determined by the FITC intensity within the volume containing clathrin immunofluorescence. The mean volumes and the SE of plasma membrane fraction are 35.4 ± 3.2, 41.24 ± 2.9, and 38.89 ± 2.7 µm3 for control, high [Ca2+]i, and anti-Cav1.3 negative [labeled with ()] control, respectively. The mean volumes of cytoplasmic fraction were derived from the difference between total cell and anticlathrin membrane-labeled volumes. Cytoplasmic volumes averaged 308.57 ± 22.28, 329.43 ± 19.45, and 319.12 ± 18.56 µm3 for control, high [Ca2+]i, and anti-Cav1.3 negative control, respectively. The degree of colocalization of Cav1.3 and clathrin between groups was qualitatively assessed by indirect immunofluorescent labeling of cells with antibodies against each protein (Fig. 6A) but also objectively demonstrated through their green to red fluorescence intensity trends as a pixel correlation graph (Fig. 6C). These data demonstrate a significant difference in digital pixel colocalization between the control and high [Ca2+]i conditions, thus suggesting Cav1.3 redistribution within the high [Ca2+]i as the Cav1.3 signal deviates toward a negative value (inverse correlation).
These results suggest an increased intracellular distribution, in addition to the plasma membrane expression, of Cav1.3 proteins under the high [Ca2+]i conditions. Short period treatment with calcium ionophore A-23187 did not cause -cell death. We treated INS-1 cells with 1 µM A-23187 for 30 min in PBS and observed no significant decrease in viability as determined by the trypan blue staining (98.6 ± 1.1%, n = 4).
The regulation of surface expression of Cav1.3 was further investigated by using a biotinylation-NeutrAvidin-elution method. INS-1 cells were incubated in either PBS with or without A-23187 (1 µM) for 30 min. After the treatments, surface proteins were biotinylated with sulfo-NHS-SS-biotin and then incubated with NeutrAvidin-linked beads. Bound proteins were eluted from the beads and subjected to Western blot analysis. We found that the expression of surface Cav1.3 (represented by the 240-kDa band) was decreased in A-23187 (high Ca2+)-treated cells compared with controls (Fig. 7A). The expression of Cav1.3 proteins in the total membrane preparation from whole cells (sampled before bead elution, indicated as total in Fig. 7) remained the same between the high-Ca2+-treated cells and the controls, whereas the channels in the surface membrane were significantly reduced in high-Ca2+ treated cells. The accumulated data are shown in Fig. 7, B and C.
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We further applied a Western blot analysis to verify that the surface protein fractions were truly from the plasma membrane. As shown in Fig. 7D, enriched Na+-K+-ATPase 1-subunit was present primarily in the biotinylation-eluted surface protein preparation. In contrast, this enzyme was not detected in the protein preparation obtained from the supernatant component of the cell lysate and was less enriched from the total membrane preparation that consisted of both surface and nonsurface membrane proteins. This ensured that our biotinylation assay reliably detected proteins from the plasma membrane.
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DISCUSSION |
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HVA Ca2+ channels play a central role in glucose-stimulated insulin secretion in pancreatic -cells (6). Studies of non-insulin-dependent diabetes mellitus (NIDDM) with Zucker diabetic fatty rats (ZDF) showed that the defect of insulin secretion in this model was associated with the loss of pancreatic
-cell L-type Ca2+ channel activity (20). Both Cav1.2 and Cav1.3 were observed in the mouse
-cell plasma membrane and are suspected to be associated with granule exocytosis (3, 27). It has been reported that a normalization of [Ca2+]i restored glucose responsiveness in a mouse cell line that is unresponsive to glucose (17). The [Ca2+]i was normalized by treating these cells with 10 µM of nifedipine for 24 h, which increased the mRNA level, as well as the protein expression of the
1-subunit of Ca2+ channels. The trafficking of Ca2+ channels, however, was not detected in these cells. In contrast, our results show the Cav1.3 subtype undergoing [Ca2+]i-dependent trafficking in rat insulin-secreting cells. Because human islet expresses primarily Cav1.3 (26), this [Ca2+]i-dependent Ca2+ channel trafficking may be important for the secretory capacity of human islet
-cells.
Abnormal handling of [Ca2+]i by cells is one of the primary defects initiating impairments in insulin action, as well as initiating diabetic complications (13). Altered [Ca2+]i homeostasis may also be the primary determinant of insufficient insulin secretion in diabetes. It is reported that desensitization of glucose-induced insulin secretion in human pancreatic islets is induced in parallel with major glucose-specific [Ca2+]i abnormalities (5). Our data suggest that the elevation of basal [Ca2+]i in -cells may induce the internalization of Cav1.3 from plasma membrane, thus reducing the capacity of Ca2+ influx in response to membrane depolarization. Indeed, it has been postulated that a decreased Ca2+-ATPase activity may contribute to the desensitization of
-cells to glucose in NIDDM (14, 19). Therefore a consequent rise in basal [Ca2+]i may diminish
-cell secretory function.
The mechanism of [Ca2+]i-dependent Cav1.3 Ca2+ channel trafficking is unclear. Run-up of Ca2+ current in rat ventricular cells has been previously reported (22). In that case, run-up was independent of phosphorylation and [Ca2+]i, and the authors concluded that mechanostimulation is probably the mechanism for increased Ca2+ channel activity. This is physiologically conceivable in heart cells; however, we propose a basal [Ca2+]i-dependent mechanism for insulin-secreting cells. It has also been reported that GTP modulated the run-up of whole cell Ca2+ channels in dissociated rat hippocampal neurons (23). More interestingly, the small G protein kir/Gem is reported to regulate the Cav1.2 expression at the cell surface via direct binding to the -subunits of Ca2+ channels (4). However, in our run-up experiments, the pipette solution contained no GTP; thus the role of the G protein-mediated channel migration in our experimental system remains unclear.
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ACKNOWLEDGMENTS |
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GRANTS
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-05151 and by the Louisiana Gene Therapy Consortium.
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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.
* L. Huang and A. Bhattacharjee contributed equally to this work.
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