Uncoupling of Cav1.2 from Ca(2+)-induced Ca(2+) release and SK channel regulation in pancreatic β-cells.

ORIGINAL

RESEARCH

Uncoupling of Cav1.2 From Ca2ⴙ-Induced Ca2ⴙ Release and SK Channel Regulation in Pancreatic ␤-Cells Yuchen Wang,* Rachel E. Jarrard,* Evan P.S. Pratt, Marcy L. Guerra, Amy E. Salyer, Allison M. Lange, Ian M. Soderling, and Gregory H. Hockerman Purdue University Life Sciences Graduate Program (R.E.J., E.P.S.P., A.M.L.) and Department of Medicinal Chemistry and Molecular Pharmacology (Y.W., M.L.G., A.E.S., I.M.S., G.H.H.), Purdue University, West Lafayette, Indiana 47907-2091

We investigated the role of Cav1.2 in pancreatic ␤-cell function by expressing a Cav1.2 II-III loop/ green fluorescent protein fusion in INS-1 cells (Cav1.2/II-III cells) to disrupt channel-protein interactions. Neither block of KATP channels nor stimulation of membrane depolarization by tolbutamide was different in INS-1 cells compared with Cav1.2/II-III cells, but whole-cell Cav current density was significantly increased in Cav1.2/II-III cells. Tolbutamide (200 ␮M) stimulated insulin secretion and Ca2⫹ transients in INS-1 cells, and Cav1.2/II-III cells were completely blocked by nicardipine (2 ␮M), but thapsigargin (1 ␮M) blocked tolbutamide-stimulated secretion and Ca2⫹ transients only in INS-1 cells. Tolbutamide-stimulated endoplasmic reticulum [Ca2⫹] decrease was reduced in Cav1.2/II-III cells compared with INS-1 cells. However, Ca2⫹ transients in both INS-1 cells and Cav1.2/II-III cells were significantly potentiated by 8-pCPT-2⬘-O-Me-cAMP (5 ␮M), FPL-64176 (0.5 ␮M), or replacement of extracellular Ca2⫹ with Sr2⫹. Glucose (10 mM) ⫹ glucagon-like peptide-1 (10 nM) stimulated discrete spikes in [Ca2⫹]i in the presence of verapamil at a higher frequency in INS-1 cells than in Cav1.2/II-II cells. Glucose (18 mM) stimulated more frequent action potentials in Cav1.2/II-III cells and primary rat ␤-cells expressing the Cav1.2/II-II loop than in control cells. Further, apamin (1 ␮M) increased glucose-stimulated action potential frequency in INS-1 cells, but not Cav1.2/II-III cells, suggesting that SK channels were not activated under these conditions in Cav1.2/II-III loop-expressing cells. We propose the II-III loop of Cav1.2 as a key molecular determinant that couples the channel to Ca2⫹-induced Ca2⫹ release and activation of SK channels in pancreatic ␤-cells. (Molecular Endocrinology 28: 458 – 476, 2014)

ype 2 diabetes is typified by hyperglycemia resulting from insufficient insulin secretion from pancreatic ␤-cells, diminished sensitivity of target tissues to the effects of insulin, or both (1). Glucose-stimulated increases in intracellular Ca2⫹ concentration are the primary drivers of insulin secretion from ␤-cells (2), and derangements in Ca2⫹ signaling are observed in ␤-cells from diabetic rodents (3, 4). Uptake of glucose into pancreatic ␤-cells and subsequent metabolism of glucose increases the ATP: ADP ratio within ␤-cells (5). ATP binds to and blocks ␤-cell KATP channels (6), which are composed of the sul-

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fonylurea receptor 1 (SUR1) and the inwardly rectifying K⫹ channel Kir6.2 (7), resulting in membrane depolarization (8). If of sufficient magnitude, membrane depolarization activates voltage-gated Ca2⫹ (Cav) channels (9). Ca2⫹ influx via the L-type channels present in ␤-cells, Cav1.2 and 1.3 (10), is thought to be particularly important for driving insulin secretion in response to elevated glucose or sulfonylurea stimulation (11). Sulfonylurea drugs have long been used to stimulate insulin secretion from pancreatic ␤-cells in persons with type 2 diabetes (12). Similar to ATP, binding of sulfonylureas to SUR1/

ISSN Print 0888-8809 ISSN Online 1944-9917 Printed in U.S.A. Copyright © 2014 by the Endocrine Society Received April 5, 2013. Accepted January 28, 2014. First Published Online February 7, 2014

* Y.W. and R.E.J. contributed equally to this work. Abbreviations: AHP, after hyperpolarization; AUC, area under the curve; CFP, cyan fluorescent protein; CICR, Ca2⫹-induced Ca2⫹ release; eIF3e, eukaryotic translation initiation factor 3 subunit e; EPAC2, exchange factor exchange protein directly activated by cAMP 2; ER, endoplasmic reticulum; FRET, fluorescence resonance energy transfer; GFP, green fluorescent protein; GLP-1, glucagon-like peptide 1; GFP, green fluorescent protein; KRBH, modified Krebs-Ringer buffer; PBST, PBS with 0.1% Tween 20; RIM, Rab 3 interacting molecule; SUR1, sulfonylurea receptor 1.

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Mol Endocrinol, April 2014, 28(4):458 – 476

doi: 10.1210/me.2013-1094

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doi: 10.1210/me.2013-1094

Kir6.2 decreases K⫹ efflux through these channels and triggers membrane depolarization (13). One role of Ca2⫹ influx via L-type Ca2⫹ channels into pancreatic ␤-cells is to stimulate further release of Ca2⫹ from the endoplasmic reticulum (ER), via Ca2⫹-induced Ca2⫹ release (CICR). This process is activated in ␤-cells depolarized with elevated concentrations of extracellular KCl (14) but apparently does not contribute to [Ca2⫹]i oscillations observed in ␤-cells stimulated with high glucose concentrations in the absence of other cosecretagogues (15, 16). The SUR1/Kir6.2 channel is known to interact directly or indirectly with several proteins that reside at the plasma membrane and play a role in insulin exocytosis. The scaffolding proteins Rab 3 interacting molecule (RIM)2 (17) and Piccolo (18) also interact with the intracellular II-III loop domain of Cav1.2 (19). We have previously reported that endogenously expressed RIM2 is present in lipid rafts and is immunoprecipitated by a Cav1.2/II-III-enhanced green fluorescent protein loop fusion protein, but not by a Cav1.3/II-III loop-enhanced green fluorescent protein fusion protein, in INS-1 cells (20). Further, we reported that overexpression of the Cav1.2/II-III loop causes accumulation of Cav1.2, but not Cav1.3, outside of lipid rafts in the rat insulinoma cell line, INS-1. Conversely, overexpression of the Cav1.3/IIIII loop selectively excluded Cav1.3, but not Cav1.2, from lipid rafts (20). The guanine nucleotide exchange factor exchange protein directly activated by cAMP 2 (EPAC2) interacts directly with both SUR1 and Piccolo (21) whereas Piccolo forms a dimer with RIM2, in a Ca2⫹dependent manner (21). Thus, a network of protein-protein interactions connecting the main components of the pathway that leads to insulin secretion stimulated by sulfonylureas implicates Cav1.2 in this process. This signaling complex of scaffolding proteins (RIM2, Piccolo), cAMP effector (EPAC2), and ion channel subunits (SUR1, Cav1.2) is of special significance because membrane depolarization-dependent calcium influx via Cav1.2 channels has been implicated in triggering Ca2⫹-induced Ca2⫹ release from the ER in pancreatic ␤-cells (22), a process that is amplified by cAMP, at least in part, through EPAC2 (22, 23). However, it is not clear whether interaction of Cav1.2 with other proteins via the II-III loop has functional significance in sulfonylurea-stimulated insulin secretion or [Ca2⫹]i transients in pancreatic ␤-cells. We show here that expression of the Cav1.2 intracellular II-III loop in the pancreatic ␤-cell line INS-1 doesn’t disrupt SUR1 or EPAC2 localization to lipid rafts but does disrupt CICR stimulated by the sulfonylurea tolbutamide, as assessed by several different experimental approaches. In addition to modulation of insulin secretion, increases in intracellular Ca2⫹ concentration in pancreatic


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␤-cells may also modulate electrical activity stimulated by glucose. One potential mechanism for fine tuning of glucose-stimulated action potentials is via activation of the small conductance Ca2⫹-activated K⫹ channel, SK (24). SK subtypes 1–3 were detected in the rat insulinoma cell line INS-1 (25). Activation of SK channels occurs as intracellular Ca2⫹ accumulates at the plasma membrane, and K⫹ efflux via SK channels limits action potential frequency (26). Indeed, block of SK channels with specific pharmacologic agents such as apamin (27) or UCL 1684 (28) can markedly increase glucose-stimulated action potential frequency in pancreatic ␤-cells, but the major source of the Ca2⫹ that activates SK channels in pancreatic ␤-cells is not clear. Here we report that modulation of glucose-stimulated action potential frequency by SK channels is disrupted in both INS-1 cells stably expressing the Cav1.2/II-III loop and primary rat ␤-cells expressing the Cav1.2/II-III loop via adenoviral transduction.

Materials and Methods Chemicals Antibodies were acquired from the following sources: EPAC2 and eukaryotic translation initiation factor 3 subunit e (eIF3e), Santa Cruz Biotechnology; SUR1, Abcam; Kir6.2, Millipore Corp; green fluorescent protein (GFP) and horseradish peroxidase-conjugated secondary antibodies, Bio-Rad Laboratories. 8-pCPT-2⬘-O-Me-cAMP-AM was from Biolog. Apamin was from Tocris Cookson. All other chemicals were from Sigma-Aldrich.

Cell culture INS-1 cells (29) and Cav1.2/II-III cells (30) were cultured in RPMI medium (Sigma-Aldrich) supplemented with 10% fetal bovine serum (HyClone Laboratories), 11 mg/mL sodium pyruvate, 10 mM HEPES, 100 U/mL penicillin, 100 ␮g/mL streptomycin, and 50 ␮M ␤-mercaptoethanol at 37°C, 5% CO2 (INS-1 RPMI). Cav1.2/II-III cells were cultured in 200 ␮g/mL G418 to maintain expression of the Cav1.2 channel II-III loop. INS-1 cells used for current clamp recordings were cultured in low glucose (2.5 mM) for 18 –24 hours prior to experiments.

Sucrose density gradient fractionation INS-1 cells and Cav1.2/II-III cells were lysed in the presence of protease inhibitors (800 nM aprotinin, 50 ␮M leupeptin, 1 ␮g/mL pepstatin, 1 mM benzimidine, 1 mM AEBSF, 10 ␮g/mL calpain I and II inhibitors; all from Sigma) and fractionated over a discontinuous 5%/30%/40% sucrose gradient as described previously (20). Fractions (1 mL) were removed from the top of each gradient, for a total of 11 fractions. Like fractions from multiple tubes were pooled and concentrated using Amicon Ultra-4 concentrator tubes (Millipore Corp). Protein concentrations were determined using the BCA assay (Pierce Chemical Co) before they were analyzed via Western blot.


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Cav1.2 Function in Pancreatic ␤-Cells

Immunoprecipitation assays Experiments in which the Cav1.2/II-III or Cav1.3/II-III loops fused to GFP were immunoprecipitated from INS-1 cell lysates with antibodies against GFP were performed as previously described (20). Immunoprecipitation of eIF3e was performed as previously described except that cell lysates were incubated with antibodies against eIF3e overnight at 4°C, and then with agarose-immobilized Protein A Plus (Thermo Scientific) overnight at 4°C before washing. Protein was eluted by incubating at 80°C in Laemmli buffer.

Western blotting Protein (20 ␮g ) from each fraction of sucrose gradients was separated by SDS-PAGE using 4% acrylamide (29:1 acrylamide-bisacrylamide) stacking gels, and 8% resolving gels for all blots except for Kir6.2, for which 10% acrylamide resolving gels were used. A constant volume of fraction 1 (50 ␮l) was loaded on each gel because it consistently lacked any protein as detected by the BCA assay. Proteins were resolved on polyacrylamide gels in 25 mM Tris, 192 mM glycine, 0.1% sodium dodecyl sulfate, and then transferred to polyvinylidene difluoride membranes using a Bio-Rad Transblot Cell in 10 mM CAPS, 10% methanol, at 70 V for 35 minutes to 2 hours, depending on the size of the protein to be detected. Polyvinylidene difluoride membranes were blocked by soaking in 5% nonfat milk in PBS with 0.1% Tween 20 (PBST) for 2 hours at room temperature. Blocked membranes were incubated with primary antibodies overnight at 4°C at the following dilutions (EPAC2, 1:100; SUR1, 1:500; Kir6.2, 1:200; eIF3e 1:100, GFP 1:1000) and then washed 3 times with PBST and incubated with secondary antibodies conjugated to horseradish peroxidase (1:1000) for 1 hour at room temperature. Excess secondary antibodies were removed by washing membranes 3 times in PBST, and proteins were detected using enhanced chemiluminescence or chemifluorescence (Amshersham Biosciences).

Electrophysiological assays Electrophysiological measurements in INS-1 cells and Cav1.2/II-III cells were recorded at room temperature using an Axopatch 200B amplifier (Molecular Devices) and filtered at 1 kHz (6-pole Bessel filter, ⫺3 dB). Patch pipettes were pulled from borosilicate glass (VWR) and fire polished to resistances of 2–5 M⍀. INS-1 cells used for current clamp recordings were cultured in low glucose (2.5 mM) for 18 –24 hours prior to experiments. For current clamp experiments and voltage clamp experiments to measure KATP channel current, the intracellular solution contained (in mM concentration) 90 K2SO4, 10 NaCl, 1 MgCl2, 1.1 EGTA, 0.1 CaCl2, 5 HEPES, 0.3 ATP, 0.2 GTP. The extracellular solution used for measuring KATP currents contained (in mM concentration) 138 NaCl, 5.6 KCl, 11.1 glucose, 10 HEPES, 1.2 MgCl2, 2.6 CaCl2, and the same solution with 2.5 mM glucose instead of 11.1 mM glucose was used in current-clamp experiments. The pH of solutions was adjusted to 7.4 with NaOH, and the osmolality was adjusted to 290 –300 mOsm. Whole-cell KATP currents were elicited by 1.3-second steps of ⫾ 20 mV from a holding potential of ⫺70 mV. Data were acquired at a sampling frequency of 1 kHz. The membrane potential of INS-1 cells and Cav1.2/II-III cells was measured using gap-free recording at a sampling frequency of 1 kHz in I ⫽ 0 current clamp mode. The KATP channel opener, diazoxide


(300 ␮M), was transiently applied to maximally open KATP channels, before application of tolbutamide. Tolbutamide solutions were prepared from stocks dissolved in 0.1 M NaOH, made fresh daily. Diazoxide solutions were prepared from stocks dissolved in dimethylsulfoxide. For recordings of voltagegated Ca2⫹ channel currents, the bath solution contained (in mM concentration) 150 Tris, 10 BaCl2, 4 MgCl2. The intracellular solution contained (in mM concentration) 130 N-methylD-glucamine, 10 EGTA, 60 HEPES, 2 ATP, and 1 MgCl2. The pH of both solutions was adjusted to 7.3 with methanesulfonic acid, and the osmolality was corrected to 290 –300 mOsM. Current-voltage relationship data were collected by applying 100-millisecond test depolarizations from ⫺50 to ⫹60 mV in 10-mV increments, from a holding potential of ⫺70 mV. Data were acquired at a sampling frequency of 10 kHz and filtered at 1 kHz. For current clamp recordings of membrane depolarization with glucose, the perforated patch technique was used to keep the intracellular signaling environment intact. The bath solution contained (in mM concentration) 138 NaCl, 5.6 KCl, 2.5 glucose, 10 HEPES, 1.2 MgCl2, 2.6 CaCl2. The intracellular solution contained (in mM concentration) 10 KCl, 10 NaCl, 70 K2SO4, 2 MgCl2, 10 HEPES, 100 –120 ␮g/mL amphotericin B. The pH of both solutions was adjusted to 7.4 with NaOH and the osmolality was adjusted to 290 –300 mOsm. A 30 mg/mL amphotericin stock was prepared in dimethylsulfoxide from powder prior to all experiments. The stock solution and the intracellular solution containing amphotericin were sonicated for at least 15 minutes prior to use, were used within 1 hour, and were protected from light. Recording pipettes were front-filled with intracellular solution without amphotericin and were back-filled with intracellular solution containing amphotericin. After formation of the cell-attached patch, the access resistance (Ra) was monitored. After 10 –15 minutes, when Ra was less than 50 M⍀, the patch was considered to be perforated enough to begin current clamp recordings. Upon adequate access, the membrane potential was measured and 300 ␮M diazoxide was briefly applied to fully hyperpolarize the membrane potential. Once a stable baseline membrane potential was achieved, 18 mM glucose was perfused into the recording chamber to stimulate membrane depolarization and action potential spiking. Whole-cell SK channel current recording was performed using a holding potential of ⫺70 mV with steps to ⫺50 mV. The intracellular and extracellular solutions for SK channel whole-cell recordings were made as described previously (25). The intracellular solution consisted of (in mM concentration): 110 potassium-gluconate, 10 KCl, 10 NaCl, 1MgCl2, 3 Mg-ATP, 5 HEPES, 10 EGTA, and 9.6 CaCl2 to achieve 2 ␮M free [Ca2⫹] (MaxChelator http://www.maxchelator.stanford.edu); pH and the osmolality were adjusted to 7.2 and 290 –310, respectively. Extracellular solution contained (in mM concentration): 140 NaCl, 4 KCl, 2 NaHCO3, 1 NaH2PO4, 1 MgSO4, 5 HEPES, 2.5 CaCl2, with pH and osmolality adjusted to 7.4 and 290 –310, respectively.

Insulin secretion assay INS-1 cells or Cav1.2/II-III cells were plated in 24-well tissue culture plates at 50%–70% confluency and incubated overnight in RPMI medium with 2.5 mM glucose at 37°C, 5% CO2. Immediately before the assay, cells were washed twice in PBS (137 mM NaCl, 10 mM phosphate, 2.7 mM KCl [pH 7.4]), and preincubated with a modified Krebs-Ringer buffer (KRBH: 134


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mM NaCl, 3.5 mM KCl, 1.2 mM KH2PO4, 0.5 mM MgSO4, 1.5 mM CaCl2, 5 mM NaHCO3, 10 mM HEPES [pH 7.4]) supplemented with 0.05% fatty acid free BSA (KRBH buffer) alone, or containing indicated concentrations of inhibitors, for 30 minutes at 37°C, 5% CO2. The buffer was decanted and replaced with fresh KRBH buffer alone (basal condition) or KRBH containing the indicated concentrations of tolbutamide or glucose with or without inhibitors for 1 hour at 37°C, 5% CO2. Secreted insulin was assayed using an ELISA for rat insulin (High-Range EIA kit; ALPCO Diagnostics). Cells were lysed in 20 mM Na2HPO4, 150 mM NaCl, 0.1% Triton X-100, 800 nM aprotinin, 50 ␮M leupeptin, 1 ␮g/ml pepstatin, 1 mM benzamidine, 1 mM 4-(2-Aminoethyl) benzenesulfonylfluoride, 10 ␮g/mL calpain inhibitor I, 10 ␮g/mL calpain inhibitor II (pH 7.4), and cellular protein in each well was determined using the BCA assay (Thermo Scientific).


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experiment to elicit CICR. Cells were alternatively excited at 340 nm and 380 nm using a filter wheel (Sutter Instrument), and the images (emission signal at 510 nm) were recorded in time lapse (one ratio every 2 seconds) using a Clara CCD camera (Andor Technology) at the beginning of KRBH application. Single and isolated cells were selected as regions of interest (ROI) and the averaged intensities (340:380 nm ratio) for each ROI at different time points were measured using Metamorph (Molecular Devices). Basal [Ca2⫹]i level of a cell was obtained by averaging the intensities of that cell within the first 30 seconds when only KRBH was present, and the basal was used to normalize the intensities at all time points. A single cell Ca2⫹ transient trace was obtained by plotting all the normalized intensities against lapsed time points. The slope of a peak was used to define a CICR peak. A spike in a trace is considered as a CICR event if the slope of the peak is equal or greater than that of the KCl-induced spike.

Intracellular Ca2ⴙ assays in 96-well plates INS-1 cells or Cav1.2/II-III cells were plated at 100% confluency in black-walled 96-well plates (Corning Life Sciences) in RPMI supplemented as described above and incubated overnight at 37°C, 5% CO2. Cells were washed twice with PBS and incubated with 5 ␮M Fura-2AM (Molecular Probes) diluted in KRBH for 1 hour at 37°C, 5% CO2. The KRBH containing Fura-2AM was then removed, and the cells were washed twice with KRBH, and equilibrated for 30 minutes at 37°C, 5% CO2, in the KRBH alone, or with indicated concentrations of inhibitors or 8-pCPT-2⬘-O-Me-cAMP-AM. Cells were stimulated by injection of the indicated concentration of sulfonylurea or glucose (or buffer control), and changes in intracellular Ca2⫹ concentrations were measured by recording the ratio of fluorescence intensities at 508/20 nm resulting from excitation of Fura-2 at 340/11 nm or 380/20 nm (center/bandpass) using a Synergy 4 multimode microplate reader (BioTek). Ratios were acquired every 0.7 second for 15 seconds before injection and 2 minutes after injection of tolbutamide. In experiments with 18 mM glucose as the stimulus, ratios were acquired every minute for 1 hour. Data were corrected for any injection artifact by subtracting the change in fluorescence ratio measured in cells injected with KRBH alone.

Single-cell intracellular Ca2ⴙ assays Cells were seeded on a poly-D-lysine coated round glass coverslip and incubated overnight in INS-1 RPMI medium. Cells were washed with PBS before loading with 3 ␮M cell-permeant Ca2⫹-sensitive dye Fura-2 (Life Technologies) at room temperature for 1 hour. The loading solution was removed, and cells were washed with KRBH followed by incubation in 0.5% BSA KRBH at room temperature for 30 minutes. The coverslip was mounted on a perfusion chamber attached to the stage of an Olympus, IX50 inverted microscope (Olympus) equipped with a PlanApo 40⫻ objective lens (0.95 na). Solution was perfused to the chamber through an in-line solution heater (Warner Instrument) at a constant flow rate (1 mL/ min) and constant temperature (37 ⫾ 1°C). Cells were first perfused with KRBH for 30 seconds followed by 30 mM KCl to depolarize the cells for 5 minutes. Verapamil (50 ␮M) was added during the final 4 minutes with 30 mM KCl to bring the [Ca2⫹]i to basal. A combination of 10 mM glucose, 10 nM glucagon-like peptide 1 (GLP-1), and 50 ␮M verapamil was applied for the rest of the

Measurement of ER Ca2ⴙ dynamics INS-1 cells or Cav1.2/II-III cells were transiently transfected with the pcDNA3-D1ER plasmid (Addgene) encoding an ERtargeted fluorescence resonance energy transfer (FRET)-based Ca2⫹ sensor in which Ca2⫹ binding enables FRET between cyan fluorescent protein (CFP) and citrine (31). Cells were split 24 – 48 hours after transfection into 4 chambered 35-mm glass bottom dishes (In Vitro Scientific) and imaged 48 –72 hours after transfection. Cell culture media were replaced with KrebsRinger buffer 30 minutes prior to imaging. The basal FRET signal was collected for 1 minute prior to treatment and 10 minutes following the addition of the positive control carbachol (500 ␮M) or tolbutamide (200 ␮M), which were diluted in Krebs-Ringer buffer. To determine the minimum FRET value, or Rmin, cells were treated with thapsigargin (1 ␮M) for 30 minutes prior to imaging and subsequently stimulated with carbachol to exhaust ER calcium stores. The D1ER FRET signal was measured with a ⫻20 objective lens using a Nikon A1 Confocal System with a Perfect Focus Ti-E Inverted Microscope. The CFP fluorophore of the D1ER FRET sensor was excited at 457 nm with an Argon laser. CFP and citrine emission were collected at 482 nm and 525 nm, respectively, using 482/35nm and 525/50 filter cubes. The citrine signal intensity, or FRET signal, was normalized to the intensity of the CFP emission, which served as an expression of ER calcium levels. The time course of the FRET signal was analyzed using NIS Elements software. Only those cells that displayed a high FRET signal were chosen for analysis, and these values were averaged among the chosen cells for each experiment at all time points. Rmin was averaged among several separate experiments and represents the point at which the D1ER sensor no longer detects a decrease in ER calcium. This value was subtracted from each FRET time point in all experiments in order to normalize for the difference in Rmin between INS-1 and Cav1.2/II-III cells. To compare ER calcium changes among experiments for each cell type, the FRET/CFP values over the time course were then normalized to the average FRET-CFP ratio over the first minute of no treatment, yielding a baseline value over the first 60 seconds of 100% or Rmax. The peak change in the FRET-CFP ratio, or decrease in ER calcium levels, was calculated based on the maximum difference from baseline over a variable time span following stimulation. Change in FRET was calculated as a percentage of the


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dynamic ER calcium range, or difference between the Rmax and Rmin, in either INS-1 or Cav1.2/II-III cells.

Isolation and adenoviral transduction of rat pancreatic islets Pancreatic islets were isolated from 2-month-old, male Wistar rats (⬃225 g) via collagenase digestion by the Islet Core Facility of the Indiana Diabetes Research Center, Indiana University School of Medicine. Islets were maintained in islet RPMI medium (RPMI-1640 medium containing 8 mM glucose, 10 mM HEPES, 10% fetal bovine serum (Hyclone), 11 mg/mL sodium pyruvate, 100 U/mL penicillin, and 100 ␮g/mL streptomycin) and kept at 37°C and 5% CO2 for 24 hours. Groups of 150 islets were handpicked into fresh islet RPMI medium for adenoviral transduction. Adenovirus encoding GFP alone or GFP fused to the C-terminal end of the Cav1.2/II-III loop domain (30) was obtained from ViraQuest. Islets were transduced at an MOI of 50 (assuming 1000 cells/islet). After 24 hours with the virus, the islets were washed with PBS and transferred to fresh islet RPMI medium. After another 24 hours, the islets were dissociated into single cells.


EPAC2 and SUR1 are highly concentrated in lipid raft fractions of sucrose gradients (the interface of 5% and 30% sucrose) in both INS-1 cells and Cav1.2/II-III cells (Figure 1). We also assessed the localization of the KATP channel subunit Kir6.2 and found that although it is present at the 5%/30% sucrose interface, it was also distributed throughout the 40% sucrose fractions in both INS-1 cells and Cav1.2/II-III cells (Figure 1). The lipid raft-resident protein caveolin 1 was detected at the 5%/30% sucrose interface but also distributed throughout the sucrose gradient in samples from both INS-1 and Cav1.2/IIIII cells. This distribution of caveolin 1 is similar to that observed in a previous study using the pancreatic ␤-cell line HIT-T15 (32). Thus, the KATP channel subunits SUR1 and Kir6.2, along with the interacting protein EPAC2, are present in lipid rafts in INS-1 cells, and their

Islet dissociation Islets were digested in 1 mL of 0.25% Trypsin-EDTA for every 100 islets and triturated repeatedly for 3 minutes. Trypsin was neutralized with the addition of 3 mL of RPMI medium, and cells were centrifuged at 800 rpm for 3 minutes, resuspended in islet RPMI medium containing 2.5 mM glucose, and plated into poly-D-lysine coated 35-mm tissue culture dishes. Dissociated cells were incubated at 37°C, 5% CO2 until they were used for electrophysiological experiments.

Data analysis Electrophysiological data were analyzed using Clampfit 8 (Molecular Devices). Area-under-the-curve (AUC) analysis and dose-response curve fits were performed using SigmaPlot 11 (Systat Software). Single-cell calcium imaging data were analyzed with Metamorph software (Molecular Devices). Analyzed data are shown as mean values ⫾ SE. Statistical significance was determined using one-way ANOVA and the Holm-Sidak post hoc test unless otherwise indicated. Differences with P values ⬍ .05 were considered significant.

Results Sucrose-density gradient fractionation of proteins involved in depolarization-induced insulin secretion in INS-1 cells and Cav1.2/II-III cells The KATP channel, composed of Kir6.2 and SUR1 subunits, plays a central role in the insulin secretion stimulated by sulfonylureas and glucose. We examined the localization of Kir6.2, SUR1, and EPAC2 in lipid rafts by fractionating the Triton X100-insoluble portion of INS-1 and Cav1.2/II-III cell lysates on discontinuous sucrose gradients. EPAC2 is reported to interact directly with both Piccolo (21) and SUR1 (19), and we found that both

Figure 1. KATP channel subunits and the cAMP effector EPAC2 are present in lipid rafts in both INS-1 cells and Cav1.2/II-III cells. Western blots detecting the indicated proteins are shown for each fraction of the sucrose-density gradients for cell lysates from INS-1 cells or Cav1.2/II-III cells. Fractions 2 and 3, at the interface of 5% and 30% sucrose, contained the Triton X-100 insoluble, low-density material (lipid rafts). Kir6.2, the pore-forming subunit of the KATP channel, is detected in, but not restricted to, lipid raft fractions. This distribution is not affected by expression of the Cav1.2/II-III loop. The dashed line across the Kir6.2 blots indicates the boundary between 2 separate polyacrylamide gels containing fractions 1–9 (top) and fractions 10 and 11 (bottom) that were simultaneously transferred to a single polyvinylidene difluoride membrane before western blotting. EPAC2 and SUR1 are highly localized to the lipid raft fractions, and this localization was not affected by expression of the Cav1.2/II-III loop. Caveolin 1 is present in lipid raft fractions but also distributed into the 40% sucrose fractions in both INS-1 cells and Cav1.2/ II-III cells. Fractions 10 and 11 are not shown for the SUR1, EPAC2, and caveolin 1 blots because they contained little or none of the indicated proteins. Molecular weight standards are shown. Each blot is representative of at least 3 independent experiments.


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ureas; therefore, we compared the modulation of electrical activity in INS-1 cells and Cav1.2/II-III cells by tolbutamide. Figure 2A shows a whole-cell voltage-clamp experiment with a Cav1.2/II-III cell held at ⫺70 mV, with alternating steps to ⫺50 and ⫺90 mV. Application of tolbutamide via external perfusion blocked both the inward and outward K⫹ current in a dose-dependent manner. Plots of the percent current blocked by tolbutamide concentrations between 100 nM and 500 ␮M are shown in Figure 2A. Fits to these plots yielded EC50 values for tolbutamide of 2.6 ⫾ 0.7 ␮M and 3.8 ⫾ 0.2 ␮M for INS-1 cells and Cav1.2/II-III cells, respectively. Because block of KATP channels by tolbutamide leads to membrane depolarization in pancreatic ␤-cells, we performed current clamp experiments to compare the potency of tolbutamide depolarizing the membrane potential in INS-1 cells and in Cav1.2/II-III cells. As shown in Figure 2B, 200 ␮M tolbutamide elicited strong membrane depolarization, Figure 2. Tolbutamide induces similar KATP block and membrane depolarization in INS-1 and leading to initiation of action potenCav1.2/II-III cells. A (left), representative trace of the dose-dependent tolbutamide block of KATP current measured in Cav1.2/II-III cells using whole-cell voltage clamp. Whole-cell current was tials in both INS-1 cells (left panel) elicited by a ⫾ 20 mV step from a holding potential of ⫺70 mV. A (right), percentage of the and Cav1.2/II-III cells (right panel). whole-cell KATP current blocked by tolbutamide in INS-1 cells (open circles, n ⫽ 4 –11) and Neither the resting membrane poCav1.2/II-III cells (closed circles, n ⫽ 5–7). IC50 values were determined by fitting the data using the equation % Current Blocked ⫽ min ⫹ (max ⫺ min)/(1 ⫹ 10ˆ(logIC50 ⫺ [tolbutamide])). B, tential, nor the membrane depolarRepresentative traces of the change in membrane potential elicited by 200 ␮M tolbutamide in ization elicited by 10, 50, 200, or INS-1 cells and Cav1.2/II-III cells. C, dose-dependent effect of tolbutamide on membrane 500 ␮M tolbutamide were signifidepolarization in INS-1 cells (open bars, n ⫽ 8 –18) and Cav1.2/II-III cells (closed bars, n ⫽ 7–13). cantly different in INS-1 cells or EC50 values were determined by fitting the data using the equation membrane potential ⫽ 1/(1 ⫹ ([tolbutamide]/EC50)n). D, Whole-cell current density comparison of voltage-gated calcium Cav1.2/II-III cells (Figure 2C). Fichannels (Cav) and KATP channels in INS-1 cells (open bars) and Cav1.2/II-III cells (closed bars). IBa nally, we compared the density of density (pA/pF) was measured at 0 mV from a holding potential of ⫺70 mV. The difference in Ba2⫹ current conducted by voltagetotal whole-cell IBa density for Cav1.2/II-III cells compared with INS-1 cells is statistically significant (***, P ⬍ .001; n ⫽ 13–15). Current density of KATP channels was determined by measuring gated Ca2⫹ channels and the density whole-cell current elicited by a ⫾ 20 mV step from a holding potential of ⫺70 mV in the of tolbutamide-sensitive K⫹ current presence of 300 ␮M diazoxide to maximally open KATP channels. There is no significant in INS-1 cells and Cav1.2/II-III cells. difference in KATP channel current density between INS-1 cells and Cav1.2/II-III cells (n ⫽ 20). s, seconds; Tolb, tolbutamide. As we reported previously, the IBa density is markedly increased in distribution on discontinuous sucrose gradients is not Ca 1.2/II-III cells compared with INS-1 cells (20); howv perturbed by expression of the Cav1.2 intracellular II-III ever, the KATP channel current density is not significantly loop. different (Figure 2D). Thus, our data demonstrate that expression of the Cav1.2 II-III loop increases voltageElectrophysiological characterization of gated Ca2⫹ channel current density but does not affect Cav1.2/II-III cells Cav1.2 is reported to exist in a complex with proteins either KATP channel current density, the sensitivity of essential for stimulation of pancreatic ␤-cells by sulfonyl- KATP channels to tolbutamide block, or the ability of tol-


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amide-stimulated insulin secretion in INS-1 cells and Cav1.2/II-III cells using the static incubation method (Figure 3). As expected from the electrophysiological characterization of tolbutamide in each cell line, 200 ␮M tolbutamide stimulated insulin secretion that was significantly different from basal secretion (Figure 3A). The L-type-selective Ca2⫹ channel blocker, nicardipine (2 ␮M), completely inhibited secretion stimulated by tolbutamide in each cell line (Figure 3A). To assess the role of internal stores of Ca2⫹ in tolbutamide-stimulated insulin secretion, the ability of 1 ␮M thapsigargin, an inhibitor of the sarcoplasmic/ER Ca2⫹-ATPase, to inhibit insulin secretion stimulated by tolbutamide was determined. In INS-1 cells, we found that thapsigargin significantly, but incompletely, blocked insulin secretion stimulated by 200 ␮M tolbutamide (Figure 3A), suggesting that both influx of Ca2⫹ via L-type Ca2⫹ channels and release of internal stores of Ca2⫹ are required for maximal insuFigure 3. Insulin secretion and Ca2⫹ transients stimulated by tolbutamide are sensitive to lin secretion stimulated by tolbutthapsigargin in INS-1 cells but not Cav1.2/II-III cells. A, Insulin secretion stimulated by 200 ␮M amide in INS-1 cells. In sharp contolbutamide is completely blocked by 2 ␮M nicardipine (black bars) in both INS-1 cells and Cav1.2/II-III cells. (***, P ⬍ .001 compared with basal; ###, P ⬍ .001 compared with nicardipine (n ⫽ 9 in 3 trast, thapsigargin treatment does separate experiments). Insulin secretion stimulated by 200 ␮M tolbutamide is partially inhibited by not inhibit tolbutamide-stimulated pretreatment with 1 ␮M thapsigargin (white bars) in INS-1 cells, but not in Cav1.2/II-III cells (***, P ⬍ secretion in Cav1.2/II-III cells (Fig.001; **, P ⬍ .01; *, P ⬍ .05 compared with basal; ##, P ⬍ .01 compared with tolbutamide alone ure 3A). Therefore, our results sug(n ⫽ 9 in 3 separate experiments)). B, Ca2⫹ transients stimulated by 200 ␮M tolbutamide are mediated by L-type Ca2⫹ channels in both INS-1 cells and Cav1.2/II-III cells. Cells pretreated with 500 gest that release of internal Ca2⫹ nM FPL 64176 (black circles) exhibit markedly greater increases in intracellular Ca2⫹ concentration in stores is not required for maximal response to tolbutamide stimulation than cells pretreated with KRBH buffer only (white circles). insulin secretion in response to tolbPretreatment of INS-1 cells or Cav1.2/II-III cells with 2 ␮M nicardipine (white diamonds) completely inhibits the Ca2⫹ transients elicited by 200 ␮M tolbutamide ⫹ 500 nM FPL 64176. Data shown are utamide in Cav1.2/II-III cells. mean ⫾ SE from experiments representative of 3 performed in quadruplicate. C, The rapid, initial To compare changes in intracelphase of Ca2⫹ transients stimulated by 200 ␮M tolbutamide is inhibited by pretreatment with 1 ␮M lular Ca2⫹ concentration induced thapsigargin in INS-1 cells but not in Cav1.2/II-III cells. White circles represent the thapsigarginsensitive portion of the tolbutamide response. Black circles represent the response to tolbutamide in by tolbutamide, INS-1 cells and the absence of thapsigargin. In contrast, the rapid peak in [Ca2⫹]i stimulated by tolbutamide is greatly Cav1.2/II-III cells in 96-well plates amplified in both cell lines by replacing extracellular Ca2⫹ with 2 mM Sr2⫹ (open diamonds). Data were loaded with the Ca2⫹ indicator shown are mean ⫾ SE from experiments representative of 3 performed in quadruplicate. Tolb, Fura-2AM, and changes in the ratio tolbutamide; Thaps, thapsigargin. of fluorescence intensity at 510 nm butamide to depolarize the membrane potential in INS-1 after excitation at 340 nm and 380 nm (340-nm to cells in a dose-dependent manner. 380-nm ratio) were measured upon injection of tolbutamide (final concentration of 200 ␮M). The changes in Insulin secretion and intracellular Ca2ⴙ transients 340-nm to 380-nm ratio upon injection of tolbutamide stimulated by tolbutamide in Cav1.2/II-III cells were corrected by subtracting the changes in 340-nm to don’t require intact intracellular stores of Ca2ⴙ Given the central role of the L-type channel Cav1.2 in 380-nm ratio measured in replicate wells of cells upon the stimulation of insulin secretion, we examined tolbut- injection of buffer only. The net change in Fura-fluores-


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cence ratio, reflecting the net change in intracellular Ca2⫹ concentration stimulated by 200 ␮M tolbutamide, is shown in Figure 3. Tolbutamide induced a biphasic rise in intracellular Ca2⫹ concentration in INS-1 cells and Cav1.2/II-III cells. In INS-1 cells, a rapid peak in intracellular Ca2⫹ concentration is followed by decay to an elevated plateau level that persisted for the duration of the 2-minute measurement (white circles: tolbutamide, Figure 3B). Tolbutamide induced Ca2⫹ transients in Cav1.2/ II-III cells that rose to a similar peak as in INS-1 cells, but Cav1.2/II-III transients appeared to decay more slowly (white circles: tolbutamide, Figure 3B). Ca2⫹ transients induced by tolbutamide in INS-1 cells and Cav1.2/II-III cells are mediated by L-type Ca2⫹ channels because pretreatment with 500 nM FPL-64176, an L-type-specific Ca2⫹ channel agonist, greatly enhanced the level of intracellular Ca2⫹ in response to tolbutamide stimulation (black circles: tolbutamide ⫹ FPL-64176, Figure 3B). When cells were pretreated with 2 ␮M nicardipine, the Ca2⫹ transient stimulated by tolbutamide and FPL64176 was greatly reduced (white diamonds: tolbutamide ⫹ FPL-64176 ⫹ nicardipine, Figure 3B). Thus, in both INS-1 cells and Cav1.2/II-III cells, the increase in intracellular Ca2⫹ concentration stimulated by tolbutamide is controlled by influx of Ca2⫹ via L-type Ca2⫹ channels. Because insulin secretion stimulated by tolbutamide was differentially sensitive to thapsigargin in INS-1 and Cav1.2/II-III cells, we examined the effect of thapsigargin on changes in intracellular Ca2⫹ concentration stimulated by tolbutamide in these 2 cell lines (Figure 3C). Pretreatment of INS-1 cells with 1 ␮M thapsigargin before stimulation with tolbutamide selectively inhibited the rapid peak in intracellular Ca2⫹ concentration but had a minimal effect on the sustained plateau phase of the transient. In the Cav1.2/II-III cells, pretreatment with thapsigargin was much less effective in inhibiting the peak in Ca2⫹ concentration. The portion of the Ca2⫹ response to tolbutamide that was thapsigargin sensitive [tolbutamide response ⫺ (tolbutamide ⫹ thapsigargin response)] is shown as white circles in Figure 3C for INS-1 and Cav1.2/II-III cells. The insensitivity of the Cav1.2/IIIII cells to pretreatment with thapsigargin could indicate either an uncoupling of Ca2⫹ influx via L-type Ca2⫹ channels from CICR or a decreased sensitivity of the Ca2⫹ release channel RYR2 to gating by Ca2⫹. To test these possibilities, we measured the tolbutamide-stimulated rise in intracellular Ca2⫹ concentration with extracellular Ca2⫹ replaced with 2 mM Sr2⫹. Sr2⫹ binds to Fura-2 with a higher KD value than Ca2⫹ (33), is able to stimulate release of Ca2⫹ from internal stores via activation of RYR2 in ␤-cells (14), and is not buffered as strongly in the cytoplasm as Ca2⫹ (34). We found that in the presence of


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2 mM extracellular Sr2⫹, 200 ␮M tolbutamide stimulated marked transient elevations of Fura-2 340-nm to 380-nm fluorescence ratio in both INS-1 cells and Cav1.2/II-III cells (Figure 3C, open diamonds). Taken together, the results presented in Figure 3 suggest that influx of Ca2⫹ via L-type channels in Cav1.2/II-III cells does not couple efficiently to release of Ca2⫹ from internal stores, but that Sr2⫹ influx strongly stimulates release of Ca2⫹ from internal stores in both INS-1 cells and Cav1.2/II-III cells. Potentiation of tolbutamide-stimulated intracellular Ca2ⴙ transients and insulin secretion by 8-pCPT-2ⴕ-O-Me-cAMP-AM Because tolbutamide-stimulated Ca2⫹ transients were not sensitive to thapsigargin in Cav1.2/II-III cells, we asked whether Ca2⫹ transients stimulated by tolbutamide would be differentially potentiated by the EPAC2-selective cAMP analog 8-pCPT-2⬘-O-Me-cAMP-AM in INS-1 and Cav1.2/II-III cells. We tested the ability of 1 ␮M and 5 ␮M 8-pCPT-2⬘-O-Me-cAMP-AM to potentiate tolbutamide-stimulated Ca2⫹ transients and insulin secretion in INS-1 and Cav1.2/II-III cells. Cells were loaded with Fura-2AM and were pretreated for 30 minutes with 0, 1, or 5 ␮M 8-pCPT-2⬘-O-Me-cAMP-AM prior to injection of either buffer only or 200 ␮M tolbutamide. Figure 4A shows the net change in Fura-2 fluorescence ratio (tolbutamide response ⫺ buffer-only response) for representative experiments in INS-1 cells and Cav1.2/II-III cells. Pretreatment with 5 ␮M 8-pCPT-2⬘-O-Me-cAMP-AM increased the early peak of the Ca2⫹ transient over pretreatment with buffer alone or 1 ␮M 8-pCPT-2⬘-O-MecAMP-AM in both INS-1 cells and Cav1.2/II-III cells. AUC analysis for the entire postinjection time course of 3 independent experiments in INS-1 cells revealed that 5 ␮M 8-pCPT-2⬘-O-Me-cAMP-AM, but not 1 ␮M, significantly increased the Ca2⫹ transient induced by tolbutamide (Figure 4B). However, AUC analysis in Cav1.2/IIIII cells revealed that 1 ␮M and 5 ␮M 8-pCPT-2⬘-O-MecAMP-AM significantly increased the Ca2⫹ transient induced by tolbutamide (Figure 4B). Because tolbutamide-stimulated Ca2⫹ transients in INS-1 cells and Cav1.2/II-III cells are potentiated by 5 ␮M 8-pCPT-2⬘-OMe-cAMP-AM and 1 and 5 ␮M 8-pCPT-2⬘-O-MecAMP-AM, respectively, we next asked whether tolbutamide-stimulated insulin secretion would be similarly potentiated by the same concentrations of 8-pCPT-2⬘-OMe-cAMP-AM. Insulin secretion was stimulated with 200 ␮M tolbutamide in the absence or presence of 0, 1, or 5 ␮M 8-pCPT-2⬘-O-Me-cAMP-AM. Figure 4C shows that 5 ␮M 8-pCPT-2⬘-O-Me-cAMP-AM significantly potentiated tolbutamide-stimulated insulin secretion in INS-1 and Cav1.2/II-III cells.


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basal FRET value was taken to be the FRET ratio measured under basal conditions (Rmax), and the minimum FRET value was taken as the FRET ratio measured in cells pretreated for 30 minutes with thapsigargin and further stimulated with 500 ␮M carbachol (Rmin). The FRET window for each cell line was taken as Rmax ⫺ Rmin. The FRET window for Cav1.2/ II-III cells was significantly lower than that measured for INS-1 cells (Figure 5A). To normalize for this difference, changes in FRET ratios in subsequent experiments were expressed as a percentage of the FRET window for that cell line. In both INS-1 cells and Cav1.2/II-III cells, application of 500 ␮M carbachol triggered a rapid and persistent reduction in FRET, indicating a decrease in ER [Ca2⫹] (Figure 5B). In contrast, stimulation of both INS-1 cells and Cav1.2/II-III cells with 200 ␮M tolbutamide resulted in a rapid but transient change in FRET ratio that returned to basal levels approximately 1 minute after stimulation (Figure 5C). Comparison of the average maximum decrease in FRET Figure 4. The EPAC-selective cAMP analog 8-pCPT-2⬘-O-Me-cAMP (ESCA) potentiates insulin secretion and Ca2⫹ transients stimulated by tolbutamide in both INS-1 cells and Cav1.2/II-III cells. A, Ca2⫹ transients ratio upon stimulation over 7– 8 exstimulated by 200 ␮M tolbutamide in both INS-1 cells and Cav1.2/II-III cells are potentiated by pretreatment periments (246 –271 cells for each of cells with ESCA. Data shown are mean ⫾ SE from representative experiments performed in condition) revealed that the decrease quadruplicate. B, Area under the curve (AUC: 2 minutes post injection) analysis of 9 (INS-1 cells) or 3 (Cav1,2.II-III cells) independent experiments in which cells were pretreated with 0, 1, or 5 ␮M ESCA before in ER [Ca2⫹] in response to carbachol stimulation with 200 ␮M tolbutamide. ***, P ⬍ .001; **, P ⬍ .01; *, P ⬍ .05 compared with tolbutamide was not different between INS-1 cells alone; ###, P ⬍ .001 compared with tolbutamide ⫹ 1 ␮M ESCA. C, Tolbutamide-stimulated insulin and Cav1.2/II-III cells, but that the desecretion is potentiated by 5 ␮M ESCA in INS-1 and Cav1.2/II-III cells (***, P ⬍ .001; **, P ⬍ .01 compared crease in ER [Ca2⫹] in response to tolwith basal; ###, P ⬍ .001 compared with 1 ␮M ESCA ⫹ tolbutamide). Tolb, tolbutamide. butamide was significantly reduced in Tolbutamide stimulation reduces ER Ca2ⴙ Cav1.2/II-III cells compared with INS-1 cells (Figure 5D). concentration to a greater extent in INS-1 cells Thus, the data in Figure 5C show a rapid, transient decrease than in Cav1.2/II-III cells in ER [Ca2⫹] corresponding with the rapid, transient thapTo further substantiate that CICR is disrupted in Cav1.2/ sigargin-sensitive rise of [Ca2⫹]in stimulated by tolbutamide II-III cells, we directly examined the change in ER [Ca2⫹] in shown in Figure 3C. Further, the tolbutamide-stimulated response to tolbutamide using the ER-targeted calmodulin- transient drop in ER [Ca2⫹] and the corresponding rise of based Ca2⫹ sensor D1ER (31). D1ER comprises the fluores- cytosolic [Ca2⫹] detected with Fura-2AM in INS-1 cells are cent proteins CFP and citrine connected by calmodulin and both markedly reduced in Cav1.2/II-III cells. a mutant calmodulin-binding peptide in such a way that Ca2⫹ binding permits FRET between CFP and citrine. Thus, Glucose and GLP-1 in the presence of verapamil in the high [Ca2⫹] environment of the ER, FRET is detected, stimulate Ca2ⴙ-induced Ca2ⴙ release events at a and stimuli that reduce ER Ca2⫹ concentration will reduce lower frequency in Cav1.2/II-III cells than in INS-1 D1ER FRET. INS-1 cells or Cav1.2/II-III cells were tran- cells To further examine the ability of Ca2⫹ influx across siently transfected with the pcDNA3-D1ER plasmid, and FRET between CFP and citrine was clearly detected. The the membrane to activate Ca2⫹ release from internal


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in KRBH for 5 minutes in the absence of glucose, resulting in a large, transient rise in intracellular Ca2⫹ concentration. One minute into the 30 mM KCl depolarization, perfusion of 50 ␮M verapamil was initiated and maintained for the duration of the experiments. Four minutes after verapamil perfusion was initiated, 30 mM KCl perfusion was terminated, and perfusion with 10 mM glucose and 10 nM GLP-1 was initiated and maintained for the duration of the experiments. Perfusion with glucose and GLP-1 stimulated discrete, transient spikes in intracellular Ca2⫹ concentration that arose from, and resolved to, baseline levels in INS-1 cells. Figure 6B shows 5 examples of the same single-cell experiment performed with 2⫹ Figure 5. The drop in ER [Ca ] in response to tolbutamide stimulation is diminished in Cav1.2/II-III Cav1.2/II-III cells. In both INS-1 cells compared with INS-1 cells. A, Quantification of the D1ER FRET window (difference between the Rmax and Rmin) in INS-1 and Cav1.2/II-III cells. INS-1 cells have a significantly greater dynamic range of cells and Cav1.2/II-III cells, some ER calcium than Cav1.2/II-III cells. The Rmax was calculated prior to any type of stimulation and cells displayed a gradual increase represents the average of 26 experiments (total of 799 cells) in INS-1 cells and 21 experiments (total in baseline Ca2⫹ concentration, of 727 cells) in Cav1.2/II-III in cells (INS-1 cells: 2.81 ⫾ 0.04; Cav1.2/II-III: 2.33 ⫾ 0.04). The Rmin was calculated through depletion of ER calcium stores using 30-minute treatment with thapsigargin whereas other cells displayed little, if followed by stimulation with carbachol. The minimal FRET ratio represents the average of 6 any, persistent increase in intracelluexperiments (total of 259 cells) in INS-1 cells and 4 experiments (total of 139 cells) in Cav1.2/II-III in lar Ca2⫹ concentration. However, cells (INS-1 cells: 2.11 ⫾ 0.07; Cav1.2/II-III: 1.99 ⫾ 0.03). The D1ER window was calculated by intermittent Ca2⫹ spikes were obsubtracting the minimal FRET value from the basal FRET value (***, P ⬍ .001; Student’s unpaired t test). B, Stimulation of INS-1 and Cav1.2/II-III cells with carbachol (500 ␮M; added at the 60-seconds served in both backgrounds. These time point) causes a rapid and prolonged decrease in the D1ER FRET signal. The INS-1 trace is a intermittent Ca2⫹ spikes stimulated representative experiment that includes 49 cells. The Cav1.2/II-III trace is a representative experiment, by glucose and GLP-1 in the preswhich includes 83 cells. C, Stimulation of INS-1 and Cav1.2/II-III cells with tolbutamide (200 ␮M; added at the 60-seconds time point) causes a transient decrease in the D1ER FRET signal followed by ence of verapamil were significantly a rapid recovery to baseline. The peak reduction in ER calcium levels is greater in INS-1 cells than more frequent in INS-1 cells than in Cav1.2/II-III cells. Data shown are representative experiments including 78 INS-1 cells, and 61 Cav1.2/ Cav1.2/II-III cells (Figure 6C; P ⬍ II-III cells. D, Quantification of the decrease in D1ER FRET signal in INS-1 and Cav1.2/II-III cells following stimulation with carbachol or tolbutamide. Stimulation of ER calcium release by tolbutamide but not .05). In addition, the average spike carbachol is significantly greater in INS-1 cells than in Cav1.2/II-III cells. Percent decrease in basal FRET amplitude and slope were both sigratio was calculated by dividing the change in FRET ratio by (Rmax ⫺ Rmin), and this was averaged nificantly reduced in Cav1.2/II-III among all experiments for both cell lines (carbachol: INS-1 cells 22.29 ⫾ 1.79; Cav1.2/II-III cells cells compared with INS-1 cells (Fig23.18 ⫾ 2.39; Tolbutamide: INS-1 cells 8.43 ⫾ 1.06; Cav1.2/II-III cells: 4.01 ⫾ 1.76)( *, P ⬍ .05, INS-1 cells compared with Cav1.2/II-III cells, Student’s unpaired t test). Carb, carbachol; Tolb, tolbutamide. ure 6, D and E; P ⬍ .001 and P ⬍ .05, respectively). Taken together, stores in INS-1 cells and Cav1.2/II-III cells, we measured the data in Figure 6 suggest that influx of Ca2⫹ in rechanges in intracellular Ca2⫹ concentration in single cells sponse to glucose and GLP-1 stimulation in Cav1.2/II-III stimulated with 10 mM glucose and 10 nM GLP-1 in the cells is less efficiently coupled to release of Ca2⫹ from presence of 50 ␮M verapamil (2). The rationale for this internal stores than in INS-1 cells. protocol is that glucose and GLP-1 potentiate CICR, whereas verapamil decreases the frequency of L-type Glucose stimulates action potentials at a greater channel openings on the plasma membrane, allowing ob- frequency in INS-1 cells and primary rat ␤-cells servation of discrete CICR events (35). Figure 6A shows 5 expressing the Cav1.2/II-III loop Because glucose uptake and metabolism stimulate examples of INS-1 cells in which changes in intracellular 2⫹ Ca concentration were measured using Fura-2. Cells membrane depolarization and action potential firing in were initially depolarized with application of 30 mM KCl pancreatic ␤-cells, we measured changes in membrane


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mM glucose) nor the average baseline membrane potential after application of 18 mM glucose (ie, the membrane potential from which the action potentials arose) were different between INS-1 cells and Cav1.2/ II-III cells (Figure 7C). Activation of the small conductance, calcium-activated potassium channel (SK channel), is known to regulate action potential frequency in pancreatic ␤-cells (27). Therefore, we examined whether or not dysregulation of SK channel activity might play a role in the increased action potential frequency observed in Cav1.2/II-III cells. In the presence of the SK channel blocker apamin (1 ␮M), the frequency of 18 mM glucose-stimulated action potentials in INS-1 cells is significantly increased (Figure 7, B and D) but is not different from the frequency measured in Cav1.2/II-III cells in the absence of apamin (Fig2⫹ ure 7D), suggesting that Ca2⫹ actiFigure 6. Discrete spikes in intracellular Ca concentration in response to glucose and GLP-1 in the presence of verapamil are less frequent in Cav1.2/II-III cells than in INS-1 cells. vation of SK channels in Cav1.2/IIRepresentative single cell Ca2⫹ traces of INS-1 cells (A) or Cav1.2/II-III cells (B). Cells were loaded III cells is aberrant. Indeed, in with 3 ␮M Fura-2AM, and perfused with KRBH for 30 seconds (basal) followed by 30 mM KCl Cav1.2/II-III cells, application of 1 for 1 minute. Coapplication of 50 ␮M verapamil for 4 minutes was used to bring the intracellular Ca2⫹ level ([Ca2⫹]i) to basal; 10 mM glucose (Glu) ⫹10 nM GLP-1⫹ 50 ␮M verapamil (verap) ␮M apamin doesn’t significantly alwas perfused for the rest of the experiment to elicit CICR. C, The CICR peak numbers per cell for ter the frequency of action potentials INS-1 and Cav1.2/II-III ( 2.261 ⫾ 0.509 (n ⫽ 46) and 0.925 ⫾ 0.290 (n ⫽ 40), respectively; *, P ⬍ stimulated by 18 mM glucose (Fig.05, one-way ANOVA with the Tukey post hoc test). D, The average amplitude of CICR peaks ure 7, B and D). To further under(⌬340/380) is significantly greater in INS-1 cells than in Cav1.2/II-III cells (0.065 ⫾ 0.003 [n ⫽ 108] and 0.03 ⫾ 0.003 [n ⫽ 40], respectively; ***, P ⬍ .001, one-way ANOVA with the Tukey stand the mechanism leading to inpost hoc test). E, The initial slopes of CICR peaks in INS-1 cells are significantly greater than those creased glucose-stimulated action of Cav1.2/II-III cells. The CICR peak slope is calculated as dividing the peak amplitude (⌬340/380) potential frequency in Cav1.2/II-III by the time (sec) from the lowest point to the highest point. The CICR peak slopes in INS-1 and Cav1.2/II-III cells are 0.02 ⫾ 0.002 (n ⫽ 108) and 0.012 ⫾ 0.001 (n ⫽ 40), respectively (*, P ⬍ cells, we compared the magnitude of .05, one-way ANOVA with the Tukey post hoc test). Glu, glucose; Verap, verapamil. the action potential after hyperpolarizations (AHPs) measured in potential in intact INS-1 cells and Cav1.2/II-II cells using INS-1 cells and Cav1.2/II-III cells during trains of action the perforated patch configuration of the current clamp. potentials stimulated with 18 mM glucose. Figure 7E Figure 7A shows representative traces of membrane de- shows example traces of action potentials recorded from polarization stimulated by application of 18 mM glucose an INS-1 cell or a Cav1.2/II-III cell. Even though the initial (arrows indicate time at which glucose was applied) to an action potentials both arise from a membrane potential of INS-1 cell and a Cav1.2/II-III cell. The higher frequency of approximately ⫺40 mV, the AHP in the Cav1.2/II-III cell action potentials stimulated by glucose in the Cav1.2/II-III is of lower amplitude (ie, less negative) than the INS-1 cells is clearly visible in these traces. Indeed, analysis of ⱖ cell. Consequently, within the time frame of the traces, 20 cells of each type revealed that glucose stimulated ac- another action potential is generated in the Cav1.2/II-III tion potentials at a significantly higher frequency in cells, but not in the INS-1 cell. The mean AHP amplitude Cav1.2/II-III cells than in INS-1 cells (P ⬍ .01) (Figure was significantly reduced in Cav1.2/II-III cells (⫺11.4 7D). This increase in action potential frequency was not mV ⫾ 0.1 mV; n ⫽ 16 cells, 1564 AHPs) compared with the result of a stronger depolarization in response to glu- INS-1 cells (⫺13.5 mV ⫾ 0.1 mV; n ⫽ 18 cells, 1087 cose because neither the resting membrane potential (2.5 AHPs; P ⬍ .001). Application of the SK channel blocker


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Taken together, the data in Figure 7 suggest that normal regulation of action potential frequency is disrupted in Cav1.2/II-III cells and that loss of SK channel activation could account for the significant increase in glucose-stimulated action potential frequency in these cells. Given the marked effects of Cav1.2/II-III loop expression on glucose-stimulated electrical activity observed in INS-1 cells, we asked whether this could be recapitulated in primary rat ␤-cells. To this end, we constructed an adenovirus encoding the same Cav1.2/II-III loopGFP fusion expressed in Cav1.2/IIIII cells and used it to transduce primary rat ␤-cells. Figure 8A shows representative current-clamp traces recorded from primary rat ␤-cells transduced with either a control adenovirus encoding GFP only (left panel) or with the adenovirus encoding the Cav1.2/II-IIII loop-GFP fusion (right panel). The arrows indicate the time at which the perfusate was switched from 2.5 mM glucose Figure 7. SK channel modulation of glucose-stimulated action potentials is disrupted in Cav1.2/ to 18 mM glucose. Neither the restII-III cells. A, Representative current-clamp traces from an INS-1 cell and a Cav1.2/II-III cell showing the development of membrane depolarization and action potentials in response to perfusion ing membrane potential (measured with18 mM glucose (arrow). B, Representative current-clamp traces from an INS-1 cell and a at 2.5 mM glucose) nor the baseline Cav1.2/II-III cell showing the effect of bath application of 1 ␮M apamin (arrow) on a train of membrane potential measured at 18 action potentials stimulated with 18 mM glucose. C, Membrane depolarization induced by 18 mM glucose in cells transduced with mM glucose is similar in INS-1 cells (white bar; n ⫽ 40) and Cav1.2/II-III cells (black bar; n ⫽ 23). D, Action potentials induced by 18 mM glucose in Cav1.2/II-III cells (black bar) have a frequency the Cav1.2/II-III-GFP encoding virus significantly greater than the frequency measured in INS-1 cells (white bar) (**, P ⬍ .01; n ⫽ 20 – were different from those measured 22). The SK channel blocker apamin (1 ␮M) increases the frequency of action potentials in INS-1 in cells transduced with the control cells (*, P ⬍ .05; n ⫽ 8 –22). The frequency of glucose-stimulated action potentials in Cav1.2/II-III cells is not significantly different in the presence or absence of apamin (n ⫽ 7–20). E, AHPs of virus (Figure 8B). However, cells glucose-stimulated action potentials are reduced in Cav1.2/II-III cells compared with INS-1 cells. transduced with the Cav1.2/II-IIIExample action potentials stimulated by glucose in an INS-1 cell (black trace) or a Cav1.2/II-III cell GFP virus fired action potentials in (gray trace) are shown. Analysis of 18 INS-1 cells (1087 AHPs) and 16 Cav1.2/II-III cells (1564 AHPs) revealed a significant decrease in the amplitude of the AHP of glucose-stimulated action response to 18 mM glucose at a sigpotentials in Cav1.2/II-III cells compared with INS-1 cells (***, P ⬍ .001). Apamin (1 ␮M) nificantly greater frequency than significantly reduces AHP amplitude in INS-1 cells (8 cells, 430 AHPs; ***, P ⬍ .001), but not in cells transduced with the control viCav1.2/II-III cells (7 cells, 470 AHPs). rus encoding GFP alone (Figure apamin (1 ␮M) significantly reduced the AHP amplitude 8C)(P ⬍ .01). Analysis of AHP amplitude in GFP and of glucose-stimulated action potentials in INS-1 cells Cav1.2/II-III-GFP-transduced ␤-cells (Figure 8D) revealed (⫺10.8 ⫾ 0.1 mV; n ⫽ 8 cells, 430 AHPs; P ⬍ .001 that the AHP amplitude of glucose-stimulated action pocompared with control INS-1 cells), but didn’t signifi- tentials was significantly reduced in cells expressing cantly affect the AHP amplitude in Cav1.2/II-III cells Cav1.2/II-III-GFP (⫺8.6 ⫾ 0.1 mV; n ⫽ 9 cells, 432 (⫺11.1 ⫾ 0.1 mV; n ⫽ 7 cells, 476 AHPs). Interestingly, AHPs) compared with controls expressing GFP only the average AHP amplitude in Cav1.2/II-III cells in the (⫺9.8 ⫾ 0.2 mV; n ⫽ 6 cells, 71 AHPs) (P ⬍ .001, Stuabsence of apamin was not significantly different from dent’s unpaired t test). Thus, consistent with what was that observed in INS-1 cells in the presence of apamin. observed in the INS-1 and Cav1.2/II-III cell lines, acute


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II-III cells (Figure 9A). Application of 100 nM apamin did not block a significant fraction of this current in either cell line if the pipette solution did not contain free Ca2⫹ (Figure 9B). In contrast, if 2 ␮M free Ca2⫹ was included in the pipette solution, 100 nM apamin blocked a significant portion of the K⫹ currents measured using this protocol (Figure 9B). The density of apamin-sensitive outward K⫹ current measured under these conditions was not different between INS-1 and Cav1.2/II-III cells (Figure 9C). Thus, the lack of modulation of glucose-stimulated AP frequency by apamin in Cav1.2/ II-III cells shown in Figures 7 and 8 is not the result of the absence of functional SK channels. We next asked whether block of SK channels with apamin could modulate glucosestimulated insulin secretion in either Figure 8. Glucose-induced action potential frequency is greater in rat ␤-cells expressing Cav1.2/ INS-1 cells or Cav1.2/II-III cells, reaII-III compared with controls expressing GFP alone. A, Representative current clamp traces from a rat primary ␤-cell transduced with adenovirus encoding GFP only (left panel) and a rat primary soning that it may also be differen␤-cell transduced with adenovirus encoding Cav1.2/II-III-GFP (right panel) in response to perfusion tially regulated by apamin in INS-1 with 18 mM glucose (arrow), which triggers membrane depolarization and action potentials. B, cells and Cav1.2/II-III cells. HowResting membrane potential and membrane depolarization induced by 18 mM glucose are not different in ␤-cells transduced with the GFP adenovirus (white bar; n ⫽ 26) or the Cav1.2/II-IIIever, we found that glucose-stimuGFP adenovirus (black bar; n ⫽ 20). C, Glucose stimulates action potentials at a greater lated insulin secretion was signififrequency in ␤-cells transduced with the Cav1.2/II-III-GFP adenovirus (black bar; n ⫽ 9) than in cantly potentiated by apamin in the control ␤-cells transduced with the GFP adenovirus (white bar; n ⫽ 6) (**, P ⬍ .01). D, The both cell types (Figure 9D). To unamplitude of AHPs measured during trains of glucose-stimulated action potentials were significantly reduced in primary rat ␤-cells transduced with the Cav1.2/II-III-GFP adenovirus derstand this difference in apamin compared with cells transduced with adenovirus encoding GFP alone (***, P ⬍ .001, Student’s t modulation of glucose-dependent test). events, we examined the time course of the rise in intracellular [Ca2⫹] expression of the Cav1.2/II-III loop in primary rat ␤-cells results in an increase in the frequency of glucose-stimu- stimulated by 18 mM glucose in INS-1 cells and Cav1.2/ lated action potentials. II-III cells using Fura-2. As shown in Figure 9E, the rise in intracellular [Ca2⫹] was delayed over the first several SK-channel current is detected in both INS-1 cells minutes of glucose stimulation in Cav1.2/II-III cells comand Cav1.2/II-III cells pared with INS-1 cells. Taken together, the data in Figure Given the apparent lack of SK channel regulation of 9 suggest that apamin does not modulate glucose-stimuglucose-stimulated action potentials in ␤-cells expressing lated action potentials in Cav1.2/II-III cells because the the Cav1.2/II-III loop, we asked whether or not SK changlucose-dependent rise in cytoplasmic [Ca2⫹] is not suffinel activity could be detected in Cav1.2/II-III cells. Wholecient to activate SK channels within the time frame of cell K⫹ currents were measured in INS-1 cells and Cav1.2/ II-III cells using steps to ⫺50 mV from a holding potential current-clamp experiments (ie, several minutes after adof ⫺70 mV, with a calculated free Ca2⫹ concentration of dition of glucose). However, within the longer time frame 2 ␮M in the recording pipette. Under these conditions, the of the glucose-stimulated insulin secretion assay (ie, 60 major K⫹ channels available to conduct current are the minutes of glucose stimulation) SK channels are eventuKATP channel and SK channels, because voltage-depen- ally activated in Cav1.2/II-III cells and exert the expected dent channels are not activated. This protocol elicited an inhibitory effect on secretion, which is relieved by apamin outward current at ⫺50 mV in both INS-1 and Cav1.2/ blockade of the channels.


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Figure 9. SK channels are activated by intracellular Ca2⫹ in both INS-1 and Cav1.2/II-III cells. A, Representative whole-cell voltage clamp traces of INS-1 (right panel) and 1.2/II-III (left panel) cells demonstrating the Ca2⫹-dependent K⫹ current blocked by apamin. Cells were depolarized to ⫺50 mV for 1 second from a holding potential of ⫺70 mV. When the pipette solution contained 0 ␮M free Ca2⫹, no current was blocked by 100 nM apamin. Tolbutamide block of KATP channel current was detected both in the absence and presence of free Ca2⫹ in the pipette, and was readily reversed upon washout. B, Comparison of fraction current remaining after 100 nM apamin between INS-1 (n ⫽ 8 at 2 ␮M [Ca2⫹]in and n ⫽ 5 at 0 [Ca2⫹]in) and 1.2/II-III (n ⫽ 8 at 2 ␮M [Ca2⫹]in and n ⫽ 6 at 0 [Ca2⫹]in) cells. Data are shown as mean values ⫾ SEM. (0.65 ⫾ 0.08 (n ⫽ 8) and 1.05 ⫾ 0.08 (n ⫽ 5) at 2 ␮M [Ca2⫹]in and 0 ␮M [Ca2⫹]in, respectively) and in 1.2/II-III cells (0.69 ⫾ 0.05 (n ⫽ 8) and 0.920 ⫾ 0.0651 (n ⫽ 6) at 2 ␮M [Ca2⫹]in and 0 ␮M [Ca2⫹]in, respectively). The fraction of current remaining in the presence of 100 nM apamin was significantly lower with 2 ␮M free Ca2⫹ in the pipette than in the absence of Ca2⫹ (*, P ⬍ .05, one-way ANOVA with Tukey post -hoc test). C, Comparison of SK current density between INS-1 (n ⫽ 8) and 1.2/II-III (n ⫽ 8) cells. Bar graphs represent the mean current density of apamin-sensitive current with 2 ␮M Ca2⫹ in the pipette solution expressed as pA/pF ⫾ SEM (INS-1 cells: 1.280 ⫾ 0.314, n ⫽ 8; 1.2/II-III cells: 0.993 ⫾ 0.381, n ⫽ 8)). The SK current density in Cav1.2/II-III cells is not different from that measured in INS-1 cells (P ⫽ .234, Mann-Whitney Rank Sum Test). D, Glucose-stimulated insulin secretion is potentiated by 100 nM apamin in both INS-1 and Cav1.2/ II-III cells. Cells were stimulated with 10 mM glucose for 1 hour, and insulin secretion was assayed as described in Materials and Methods. The addition of apamin significantly increased insulin secretion compared with glucose alone (P ⬍ 0.05; Student’s unpaired t test). E, The rise in intracellular Ca2⫹ in response to 18 mM glucose is delayed in Cav1.2/II-III cells compared with INS-1 cells. Cells were loaded with Fura-2, and stimulated by injection of 18 mM glucose. The 340 nm to 380 nm ratio was measured every minute for 1 hour. Control experiments were performed with injection of KRBH containing no glucose. The data shown are corrected by subtraction of the slow, linear increase in 340:380 ratio observed in the absence of glucose. Data shown are the means ⫾ SEM of an experiment done in triplicate and are representative of 3 independent experiments. ms, milliseconds; tolb, tolbutamide; wash, washout.


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IQGAP1 and eIF3e coimmunoprecipitate with the Cav1.2/II-III loop Given the marked effects of the expression of the Cav1.2/II-III loop in INS-1 cells and primary ␤-cells on Ca2⫹ signaling, we examined the ability of the Cav1.2/IIIII loop to interact with proteins in INS-1 cells using coimmunoprecipitation experiments. The Cav1.2/II-III loop was previously reported to bind directly to the C2 domains of RIM2 (36) and Piccolo (19) and to coimmunoprecipitate RIM2 from INS-1 cell lysates (20). The GTPbinding protein CDC42 is reported to play a key role in regulating actin polymerization during glucose-stimulated insulin secretion (37) (38). IQGAP1 is a scaffold protein that binds CDC42 in a manner that is regulated by Ca2⫹-calmodulin (39). Further, the C-terminal regions of IQGAP1and -2 were recently found to form a pseudo-C2 fold, capable of binding phosphatidylinositol 3,4,5,-trisphosphate (40). Therefore, we examined whether the Cav1.2/II-III loop could interact with IQGAP1 by immunoprecipitating the loop with antibodies against GFP and blotting for IQGAP1. As shown in Figure 10A, IQGAP1 coimmunoprecipitated with neither GFP transiently expressed in INS-1 cells, nor with the GFP-tagged Cav1.3/II-III loop stably expressed in INS-1


cells. In contrast, immunoprecipitation of the GFP-tagged Cav1.2/II-III loop from Cav1.2/II-III cell lysates coimmunoprecipitated IQGAP1. The translation initiation factor eIF3e was recently reported to mediate Ca2⫹-dependent internalization of Cav1.2 in INS-1 cells (41). eIF3e was previously reported to bind directly to a 3-amino acid sequence toward the C-terminal end of the II-III loop of Cav1.1, 1.2, 1.3, 2.2, and 2.3 (42). Disruption of a mechanism leading to internalization of Cav channels could account for the marked increase in Cav current density that we observed in Cav1.2/II-III cells (Figure 2D). We therefore asked whether the Cav1.2/II-III loop was interacting with eIF3e using immunoprecipitation of eIF3e and Western blotting with antibodies to GFP to detect the Cav1.2/II-III loop fusion. As shown in Figure 10B, immunoprecipitation of eIF3e did coimmunoprecipitate the Cav1.2/II-III loop. We were unable to detect coimmunoprecipitation of the Cav1.2/II-III loop and eIF3e using antibodies against GFP (data not shown). It is possible that eIF3e binding to the Cav1.2/II-III loop/GFP fusion hinders binding of antibodies to GFP because the eIF3e binding motif and the GFP fusion site are both on the C-terminal end of the loop. These data suggest that eIF3e and the Cav1.2/II-III loop interact in Cav1.2/II-III cells in a manner that may result in accumulation of Cav channels in the plasma membrane of these cells.

Discussion

Figure 10. The Cav1.2/II-III loop interacts with IQGAP1 and eIF3e in INS-1 cells. A, IQGAP1 coimmunoprecipitates with the Cav1.2/II-III loop/GFP fusion peptide, but not the Cav1.3/II-III loop fusion peptide, from INS-1 cell lysates. Cell lysates were incubated with anti-GFP antibodies coupled to agarose, washed, and eluted by incubation at 80°C in Laemmli buffer. Ly, 100 ␮g of cell lysate; Ub, unbound proteins removed after incubation with agarose-immobilized antibodies; W, protein in the last wash fraction before elution; E, protein eluted by incubation at 80°C in Laemmli buffer. Samples were blotted with antibodies against IQGAP1 (upper panel) or GFP (lower panel). Blots shown are representative of 3 separate experiments. B, The Cav1.2/II-III loop coimmunoprecipitates with eIF3e. Cav1.2/II-III lysates were incubated with antibodies to eIF3e and protein Asepharose, washed, and eluted with Laemmli buffer. Lanes are labeled as described above. Samples were blotted with antibodies against GFP to detect the Cav1.2/II-III loop-GFP fusion peptide (upper panel) or with antibodies against eIF3e (lower panel). Blots shown are representative of 4 separate experiments. IB, immunoblotting; IP, immunoprecipitation.

Key proteins involved in triggering insulin secretion are present in lipid rafts in INS-1 cells Lipid raft localization of Cav1.2 has been reported in insulin-secreting cells (20, 32), but the functional significance of this localization is largely unclear. However, the scaffolding proteins RIM2 and Piccolo were also detected in lipid rafts prepared using sucrose gradient centrifugation (20) and are reported to bind directly to the Cav1.2 II-III loop via their C2 domains (19). Moreover, RIM2 is coimmunoprecipitated from INS-1 cell lysates by the II-III loop of Cav1.2 but not that of Cav1.3 (20). Further, the KATP channel subunit SUR1 binds to the cAMP effector EPAC2 (19). Activation of EPAC2 by cAMP in pancreatic ␤-cells is linked to amplification of CICR via activation of PLC-⑀ (43) and a 2-APB-sensitive Ca2⫹ influx (44). Here we show that SUR1 and EPAC2 are highly enriched in lipid rafts prepared from INS-1 cell lysates (Figure 1), suggesting that these proteins, along with Cav1.2, may indeed colocalize to lipid rafts on the membrane of INS-1 cells. In contrast, Kir6.2, although present in lipid rafts, is more widely distributed throughout sucrose density gra-


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dients, consistent with reports that it also resides within secretory granules in ␤-cells (45, 46), and that Kir6.2 is present in secretory granules in ␤-cells from Sur1⫺/⫺ mice (47). Interestingly, expression of the Cav1.2/II-III loop did not appear to affect the distribution of Piccolo and RIM2 (20), or SUR1, EPAC2, and Kir6.2 (this study) on sucrose density fractions, in contrast to the marked shift of Cav1.2 to higher density fractions in Cav1.2/II-III cells (20) reported previously. Our finding that SUR1 and Kir6.2 are found in lipid rafts in INS-1 cells contrasts with a previous study, using HIT-T15 cells, which found them to be excluded from lipid rafts, whereas Cav1.2 was found to be highly localized to lipid rafts (32). The reason for this discrepancy is not clear; however, the colocalization of Cav1.2, SUR1, Kir6.2, Piccolo, RIM2, and EPAC2 in lipid rafts is consistent with reports that these proteins interact physically and functionally (19, 21). Expression of the Cav1.2/II-III loop does not disrupt KATP channel activity but increases Cav channel current density We compared the electrophysiological properties of the Cav1.2/II-III cells with those of INS-1 cells (Figure 2). We found no difference between Cav1.2/II-III cells and INS-1 cells in regard to either whole-cell KATP channel current density and sensitivity to block by the sulfonylurea tolbutamide, or to the potency of membrane depolarization and action potential frequency (data not shown) stimulated by tolbutamide in current clamp experiments conducted using the whole-cell configuration. However, as reported previously (20), we did find that whole-cell Cav current density (measured using Ba2⫹ as the permeant ion) was significantly increased in Cav1.2/ II-III cells compared with INS-1 cells. This increase in Cav current density was not confined to L-type channels because the portion of current blocked by 10 ␮M nifedipine was not different in INS-1 cells and Cav1.2/II-III cells (20). The mechanism for this increase in Cav current density is suggested by the finding that the translation initiation factor eIF3e binds to the II-III loop of multiple Cav channels in a Ca2⫹-dependent manner to mediate internalization from the plasma membrane (42). Very recently, this mechanism has been reported to regulate the density of Cav1.2 channel on the plasma membrane of INS-1 cells (41). The 3-amino acid motif in the Cav1.2 II-III found to be critical for interaction with eIF3e (33) is present in the Cav1.2/II-III loop-GFP fusion used in this study. Indeed, we found that antibodies to eIF3e could coimmunoprecipitate the Cav1.2/II-III loop. Thus, the marked increase in Cav current density is likely driven, at least in part, by Cav1.2/II-III/GFP fusion interference with retrieval of Cav channels from the plasma membrane via


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the eIF3e-dependent mechanism. However, even with the increase in Cav current density, CICR is diminished in Cav1.2/II-III cells. It is tempting to speculate that interruption of the Cav1.2 channel-piccolo and or -RIM2 interactions by expression of the Cav1.2/II-III loop excludes Cav1.2 from a signaling complex critical for CICR, but does not otherwise interfere with insertion of Cav1.2 into the plasma membrane. However, this study cannot exclude the possibility that loss of eIF3e modulation of Cav1.2 plasma membrane expression, or interruption of a Cav1.2/IQGAP1 interaction, is contributing to the deficits in CICR and SK channel activation observed in Cav1.2/II-III cells. Interestingly, IQGAP1 is an essential component of the Wnt-receptor-actin-myosin-polarity structure, which recruits cortical ER to the plasma membrane during directional cell movement (48). It will be of interest to determine the relative roles of Cav1.2 interactions with scaffolding proteins and/or eIF3e in CICR and SK channel activation in pancreatic ␤-cells. Ca2ⴙ-induced Ca2ⴙ release is diminished in Cav1.2/ II-III cells compared with INS-1 cells Given that we detected no deficits in KATP channel function or sensitivity to tolbutamide in Cav1.2/II-III cells compared with INS-1 cells, we examined both insulin secretion and Ca2⫹ transients simulated by tolbutamide in these cell lines (Figure 3). Secretion stimulated by tolbutamide in INS-1 cells is completely inhibited by nicardipine and partially inhibited by pretreatment with thapsigargin. This pharmacologic profile is consistent with contributions by both Ca2⫹ flux across the membrane through L-type channels (thapsigargin-resistant portion) as well as from Ca2⫹-induced release of Ca2⫹ from internal stores (thapsigargin-sensitive portion) to tolbutamide-stimulated insulin secretion. In contrast, insulin secretion stimulated by tolbutamide in Cav1.2/II-III cells was completely blocked by nicardipine, but completely resistant to pretreatment with thapsigargin, suggesting that release of Ca2⫹ from internal stores does not contribute to tolbutamide-stimulated insulin secretion in these cells. Indeed, the strong inhibition of the rapid tolbutamide-stimulated peak in intracellular Ca2⫹ concentration by thapsigargin in INS-1 cells was markedly reduced in Cav1.2/II-III cells, supporting the conclusion that Ca2⫹-induced Ca2⫹ release from internal stores is disrupted in these cells. This conclusion was strongly supported by measurements of ER [Ca2⫹] changes with the D1ER Ca2⫹ indicator (Figure 5). Indeed, the kinetics of the decrease in ER [Ca2⫹] upon tolbutamide stimulation closely reflected the corresponding increase in cytosolic Ca2⫹ measured with Fura-2 in INS-1 cells, and both responses were similarly inhibited in Cav1.2/II-III cells. In-


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terestingly, the observation that the D1ER FRET window was significantly lower in Cav1.2/II-III cells compared with INS-1 cells suggests that localization of Cav1.2 via the II-III loop may contribute to maintenance of ER Ca2⫹ stores. To determine whether the above observations were unique to sulfonylurea stimulation, we compared CICR in INS-1 and Cav1.2/II-III cells stimulated with GLP-1 and glucose in the presence of 50 ␮M verapamil (Figure 6). The rationale behind these experiments is to enhance the sensitivity of RYR2-mediated release of Ca2⫹ from stores via the actions of GLP-l (49, 50), but minimize the Ca2⫹ influx via L-type Ca2⫹ channels, which is also enhanced under these conditions (51), using the Ca2⫹ channel blocker verapamil to reduce the probability of channel opening (52). This protocol is reported to generate discrete CICR events in pancreatic ␤-cells (2), and indeed, we observed, discrete, intermittent Ca2⫹ spikes using this protocol with INS-1 cells. When this protocol was applied to Cav1.2/II-III cells, these discrete Ca2⫹ spikes were detected less frequently and were of a significantly lower amplitude and slope compared with those detected in INS-1 cells. These data further support the conclusion that Ca2⫹ influx via Cav1.2 channels is not efficiently coupled to release of Ca2⫹ from internal stores in Cav1.2/ II-III cells during glucose ⫹ GLP-1 stimulation. Given the increased Cav current density in Cav1.2/II-III cells, we propose that the observed disruption of CICR is the result of a spatial uncoupling of Cav1.2 and RYR2 in Cav1.2/II-III cells. An alternative explanation might be a deficit in the RYR2 response to increases in intracellular Ca2⫹ concentration in Cav1.2/II-III cells. However, our findings that replacement of extracellular Ca2⫹ with Sr2⫹ restored the thapsigargin-sensitive, tolbutamide-stimulated early phase of the Ca2⫹ transient in Cav1.2/II-III cells (Figure 3), and that the EPAC2-selective cAMP analog 8-pCPT-2⬘-O-Me-cAMP-AM strongly potentiated both tolbutamide-stimulated insulin secretion and Ca2⫹ transients in Cav1.2/II-III cells (Figure 4) argue against this interpretation. Cav1.2/II-III cells exhibit an increased frequency of action potentials in response to glucose If coupling of Ca2⫹ influx via Cav1.2 to intracellular signaling has, in fact, been disrupted in Cav1.2/II-III cells, then electrical excitability in response to glucose could be enhanced in Cav1.2/II-III cells compared with INS-1 cells, if it contributes to activation of small-conductance Ca2⫹activated K⫹ (SK) channels in pancreatic ␤-cells. SK channels are expressed in mouse ␤-cells (53), INS-1 cells (25), and human ␤-cells (27). Pancreatic ␤-cells from SK-4 channel knockout mice exhibited an increase in glucose-


stimulated action potential frequency (54), and block of SK channels in human ␤-cells with the toxin apamin increases glucose-stimulated action potential frequency, reflecting the role of SK channels in mediating the AHP (25). Our finding that glucose-stimulated action potentials in both Cav1.2/II-III cells and in primary rat ␤-cells transduced with an adenovirus encoding the Cav1.2/II-III loop occurred at a frequency approximately twice that of their appropriate controls (Figures 7 and 8), supports the conclusion that interaction of Cav1.2 with other proteins via the II-III loop is required for normal regulation of SK channels. Loss of SK channel activation as the mechanism for this increase in glucose-stimulated excitability is strongly supported by the insensitivity of glucose-stimulated action potentials in Cav1.2/II-III cells to apamin and the highly significant decrease in the magnitude of the AHP during trains of glucose-stimulated action potentials in Cav1.2/II-III cells and primary ␤-cells transduced with the Cav1.2/II-III-GFP virus. Further, the data in Figure 9 demonstrate that SK channels are activated in Cav1.2/IIIII cells if 2 ␮M Ca2⫹ is provided in the pipette solution. Thus, the primary deficit leading to dysregulation of SK channels in Cav1.2/II-III cells is likely the prolonged delay in the rise of [Ca2⫹]in in response to glucose. In conclusion, this study provides evidence that the Cav1.2 II-III loop is critical for efficient CICR and regulation of SK channel activity in pancreatic ␤-cells. Specific interactions between Cav1.2 and RIM2, Piccolo, IQGAP1, and eIF3e may contribute to maintenance of CICR and/or SK channel regulation, but the role that each of these individual proteins plays remains to be determined. In type 2 diabetes, defects in ␤-cell Ca2⫹ responses to glucose are commonly observed (55). A recent study reported that transgenic mice expressing a phosphomimic, gain-of-function mutant of RYR2 (S2814D) had diminished Ca2⫹ responses to glucose and KCl and developed glucose intolerance (56). Thus, it is conceivable that derangements in CICR could contribute to type 2 diabetes. Interestingly, exposure of INS-1 cells to 28 mM glucose for 72 hours disrupted lipid rafts and inhibited insulin secretion in response to both KCl and glucose (57), an effect that was attributed to glucose inhibition of cholesterol synthesis. In addition, exposure of pancreatic islets to palmitate inhibited glucose-stimulated insulin secretion and dispersed microdomains of elevated intracellular Ca2⫹ concentration in response to brief episodes of membrane depolarization in pancreatic ␤-cells (58). It will be of interest to examine whether disruption of Cav1.2 localization, and the loss of CICR efficiency and regulation of glucose-stimulated electrical excitability described herein, plays a role in pancreatic ␤-cell dysfunction induced by glucotoxicity or lipotoxicity.


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Acknowledgments We thank Dr Li Li for assistance in analyzing the FRET data and Natalie Stull of the Indiana Diabetes Research Center Islet Core for isolation of rat pancreatic islets. Address all correspondence and requests for reprints to: Gregory H. Hockerman, 575 Stadium Mall Drive, West Lafayette, IN 47906. E-mail: [email protected]. This work was supported by a grant from the National Institute of Diabetes and Digestive and Kidney Diseases (R01 DK064736) (to G.H.H.). Disclosure Summary: The authors declare no conflicts of interest.

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Regulation of amylase release from dispersed pancreatic acinar cells.

Regulation of Ca2+ release from sarcoplasmic reticulum in skeletal muscles.

Age-dependent uncoupling of mitochondria from Ca2⁺ release units in skeletal muscle.

Effect of GTP and Ca2+ on inositol 1,4,5-trisphosphate induced Ca2+ release from permeabilized rat exocrine pancreatic acinar cells.

Ca2+-release channel RyR1 by S-palmitoylation.

Gating of the cardiac Ca2+ release channel: the role of Na+ current and Na(+)-Ca2+ exchange.

Sarcoplasmic reticulum Ca²⁺ release is both necessary and sufficient for SK channel activation in ventricular myocytes.

Effect of perfusate [Ca2+] on cardiac sarcoplasmic reticulum Ca2+ release channel in isolated rat hearts.

Cytoplasmic Ca2+ oscillations in pancreatic beta-cells.

Neuronal Na+ channel blockade suppresses arrhythmogenic diastolic Ca2+ release.

Involvement of Ca2(+)-induced Ca2+ release in the volume regulation of human epithelial cells exposed to a hypotonic medium.

Ca2+ release from isolated sarcoplasmic reticulum of guinea-pig psoas muscle induced by K(+)-channel blockers.

Rose bengal activates the Ca2+ release channel from skeletal muscle sarcoplasmic reticulum.

Ca2+ and vasopressin release in isolated rat neurohypophysis: differential effects of four classes of Ca2+ channel ligands.

Regulation of Ca2+ release from mitochondria by the oxidation-reduction state of pyridine nucleotides.

NPY-stimulated Ca2+ mobilization in SK-N-MC cells is enhanced after isoproterenol treatment.

Antiarrhythmic Mechanisms of SK Channel Inhibition in the Rat Atrium.

Osteopontin Affects Insulin Vesicle Localization and Ca2+ Homeostasis in Pancreatic Beta Cells from Female Mice.

K+ channel openers, cromakalim and Ki4032, inhibit agonist-induced Ca2+ release in canine coronary artery.

Regulation of acetylcholine release from neuroblastoma x glioma hybrid cells.

Regulation of nicotine-evoked dopamine release from PC12 cells.

paracrine regulation of prolactin release from human decidual cells.

Measurement of intracellular Ca2+ release in intact cells using 45Ca2+.

Measurement of intracellular Ca2+ release in permeabilized cells using 45Ca2+.