Stromal Interaction Molecule 1 (STIM1) Regulates ATP-sensitive Potassium (K_ATP) and Store-operated Ca²⁺ Channels in MIN6 β-Cells

Abstract

Stromal interaction molecule 1 (STIM1) regulates store-operated Ca²⁺ entry (SOCE) and other ion channels either as an endoplasmic reticulum Ca²⁺-sensing protein or when present in the plasma membrane. However, the role of STIM1 in insulin-secreting β-cells is unresolved. We report that lowering expression of STIM1, the gene that encodes STIM1, in insulin-secreting MIN6 β-cells with RNA interference inhibits SOCE and ATP-sensitive K⁺ (K_ATP) channel activation. The effects of STIM1 knockdown were reversed by transduction of MIN6 cells with an adenovirus gene shuttle vector that expressed human STIM1. Immunoprecipitation studies revealed that STIM1 binds to nucleotide binding fold-1 (NBF1) of the sulfonylurea receptor 1 (SUR1) subunit of the K_ATP channel. Binding of STIM1 to SUR1 was enhanced by poly-lysine. Our data indicate that SOCE and K_ATP channel activity are regulated by STIM1. This suggests that STIM1 is a multifunctional signaling effector that participates in the control of membrane excitability and Ca²⁺ signaling events in β-cells.

Introduction

The primary physiological role of pancreatic β-cells is to regulate metabolism by sensing changes in blood glucose concentration and secreting insulin accordingly. Interplay between glucose metabolism, closure of ATP-sensitive K⁺ (K_ATP) channels, and Ca²⁺ signaling evoked by activation of voltage-dependent Ca²⁺ channels (VDCCs)² form the molecular basis of the so-called consensus model of glucose-stimulated insulin secretion. Nevertheless, this simple model does not address the temporal and spatial diversity of Ca²⁺-dependent signal transduction events generated in β-cells consequent to stimulation with secretagogues (1).

Ca²⁺ homeostasis and signal transduction are controlled by multiple ion transport mechanisms and organelles. Store-operated Ca²⁺ entry (SOCE) plays a critical role regulating spatial and temporal changes in cytoplasmic Ca²⁺ concentration ([Ca²⁺]_c), endoplasmic reticulum (ER) Ca²⁺ homeostasis and protein biosynthesis, mitochondrial function, secretion, and cell viability (2, 3). The ER regulates Ca²⁺ signaling by acting as a subcellular store for Ca²⁺ that can be rapidly and transiently released into the cytoplasm, and the ER luminal Ca²⁺ concentration ([Ca²⁺]_ER) regulates gating of store-operated Ca²⁺ (SOC) channels located in the plasma membrane (PM).

The presence of SOCE in β-cells was first reported in 1994 (4–6). Patch clamp electrophysiology has been used to characterize some of the electrical properties of SOC current (I_SOC) in β-cell lines and primary β-cells isolated from rodent islets of Langerhans (5–7). The store-operated conductance in insulin-secreting cells was found to be an inwardly rectifying current carried by a non-selective cation (CRAN) channel rather than a highly Ca²⁺-selective Ca²⁺-release activated Ca²⁺ (CRAC) channel (5, 8). The electrical characteristics of I_SOC in insulin-secreting cells are similar to mammalian transient receptor potential (trp)-related channels, a gene family that includes Ca²⁺ store-operated channels (9, 10), and several trp genes are expressed in β-cells (11, 12). Members of the Orai family of genes (Orai1, Orai2, and Orai3) and some TRPC channels (TRPC1, TRPC2, and TRPC4) form store-operated channels either as homomeric or heteromeric complexes (13). Although Affymetrix GeneChip microarrays show that rodent β-cell lines as well as human islets express Orai genes (T1Dbase), the roles of the proteins encoded by these genes in β-cell store-operated Ca²⁺ entry and β-cell stimulus-secretion coupling remain to be clarified.

Stromal interaction molecule-1 (STIM1), a single transmembrane-spanning Ca²⁺ binding phosphoprotein located in both the ER and PM, functions as a luminal Ca²⁺ sensor of the ER and couples changes in [Ca²⁺]_ER with activation of SOCE (14). STIM1 is activated by cellular stimuli and stress conditions that lower [Ca²⁺]_ER. After a decrease in [Ca²⁺]_ER, STIM1 translocates to plasma membrane-associated membrane (PAM) complexes, specialized regions where the ER and PM are closely apposed. At the PAM, STIM1 directly interacts with plasmalemmal store-operated ion channels and other protein targets that control a wide range of cellular signaling events (15, 16).

STIM1 is expressed in β-cells, but its role in β-cell Ca²⁺ signaling has not been fully resolved (17). Confocal microscopy demonstrated that subsequent to decreasing [Ca²⁺]_ER with thapsigargin, an inhibitor of sarco(endo)plasmic reticulum calcium ATPase (SERCA), fluorescently-tagged STIM1 expressed in MIN6 β-cells translocates to regions near the plasma membrane (18) consistent with STIM1 regulating β-cell SOC channels. STIM1 translocation and formation of sub-PM puncta or clusters of STIM1 molecules can also occur in response to cAMP elevation but without activation of SOCE (19).

To better understand the role of STIM1 in β-cells, we utilized patch clamp electrophysiology, Ca²⁺ imaging, and RNA interference. We demonstrate for the first time that in insulin-secreting cells STIM1 participates in the activation of store-operated Ca²⁺ current but also directly interacts with K_ATP channels. Our data suggest that STIM1 regulates β-cell excitability by coupling multiple PM signaling mechanisms with changes in [Ca²⁺]_ER.

Results

Store-operated Ca²⁺ Entry in MIN6 Cells

Application of carbachol, a muscarinic receptor agonist, in the absence of external Ca²⁺ evoked a transient increase in [Ca²⁺]_c (Fig. 1A). After discharge of the intracellular Ca²⁺ stores, the addition of extracellular Ca²⁺ caused a biphasic increase in [Ca²⁺]_c characteristic of SOCE found in a wide range of cell types (Fig. 1A). The response of MIN6 cells with depleted stores to Ca²⁺ reintroduction was significantly reduced (peak amplitude reduced by 11% and area under the Ca²⁺ response curve (area-under-curve (AUC)) by 24%) by nimodipine (5 μm), an inhibitor of voltage-gated Ca²⁺ channels (Fig. 1, B and C). These data indicate that Ca²⁺ influx through L-type voltage-gated Ca²⁺ channels makes a small, but significant, contribution to the increase in [Ca²⁺]_c under these conditions. This nimodipine-sensitive influx could result from spontaneous activity of L-type channels at the resting potential (20) or from a small SOCE-induced depolarization and weak activation of VDCCs. The broad spectrum channel inhibitor, 1-[2-(4-methoxyphenyl)-2-[3-(4-methoxyphenyl) propoxy]ethyl-1H-imidazole hydrochloride (SKF 96365), is reported to inhibit SOCE in β cells (8) and was also tested. MIN6 cells were plated into 96-well plates and Fura 2-loaded in Ca²⁺-free solution containing 1 μm thapsigargin (Tg) to deplete ER stores and activate SOCE. The Fura 2 loading solution was replaced with Ca²⁺-free bath solution containing Tg ± SKF96365. Ca²⁺ addition to the bath solution induced SOCE that was dose-dependently inhibited by SKF96365 (Fig. 1D).

FIGURE 1.

Store-operated Ca²⁺ entry in MIN6 cells. A, MIN6 cells were loaded with Fura 2 and perifused with test solutions containing 2 mm glucose at 37 °C. Cells were initially bathed in normal, 2.5 mm Ca²⁺ extracellular solution to establish a baseline. The bath solution was then exchanged for Ca²⁺-free solution (10 μm EGTA), Ca²⁺-free with 250 μm carbachol (CCh) followed by Ca²⁺-free with 10 μm cyclopiazonic acid (CPA) to discharge and prevent refilling of intracellular Ca²⁺ stores. Reintroduction of 2.5 mm Ca²⁺ caused a rise of cytosolic [Ca²⁺] ([Ca²⁺]_c) representing SOCE. B, the addition of 5 μm nimodipine (Nimod) before the reintroduction of extracellular Ca²⁺ caused a small decrease in the peak amplitude. The efficacy of nimodipine to block voltage-gated Ca²⁺ channels was confirmed by applying a pulse of 56 mm KCl solution (indicated by arrow). Whereas under control conditions KCl elicited a large rise in [Ca²⁺]_c, this rise was inhibited by nimodipine. C, the AUC of the SOCE response shown in B was measured for the 160-s period after Ca²⁺ addition and was significantly reduced by nimodipine (52 control cells, 24 nimodipine-treated; *, p = 0.002, ANOVA). D, MIN6 cells in a 96-well plate were loaded with Fura 2 in Ca²⁺-free solution containing 2 mm glucose and 1 μm Tg. The cells were then washed with fresh bath solution containing Tg. Ca²⁺-free bath solution (Cont) or Ca²⁺-containing solution with Tg and SKF96365 (final, 2 mm [Ca²⁺], SKF96365 concentration indicated in μm) was injected at 100 s. SOCE was dose-dependently blocked by SKF96365.Data are plotted as the mean ± S.E., error bars shown are larger than symbols, averaged from 12 wells for each solution and are representative of three independent experiments.

Knockdown of STIM1 Reduces SOCE in MIN6 Cells

STIM1 functions as a sensor of [Ca²⁺]_ER, coupling ER Ca²⁺ store content with generation of SOCE and cytosolic Ca²⁺ oscillations, but knowledge of the role of STIM1 in β-cell Ca²⁺ signaling is limited. We employed a knockdown strategy using viral transduction with short-hairpin RNAs (shRNA) directed against STIM1. Cells expressing a scrambled sequence with no known target (shRNA-scr, ss) were used as controls. By qPCR screening of RNA isolated from cells expressing different shRNA constructs, we identified a construct (shRNA-STIM1, sh) that significantly reduced STIM1 mRNA expression and protein levels by 84 and 70%, respectively (Fig. 2A). In shRNA-STIM1 cells, SOCE peak amplitude and AUC were 67 and 61%, respectively, lower than shRNA-scr control cells (Fig. 2, B–D). The specificity of shRNA construct targeting and absence of off-target effects were confirmed by transducing shRNA-STIM1 cells with an adenovirus encoding the human isoform of STIM1 (AdV-hSTIM1, av). Reconstituting STIM1 protein expression (Fig. 2A) in this way partially restored SOCE (Fig. 2, B–D). We also used PCR with primers spanning exon 11 to test for the expression of long and short isoforms of STIM1 (21, 22) and found no evidence for expression of the long isoform in MIN6 cells (data not shown).

FIGURE 2.

Store-operated Ca²⁺ entry in MIN6 cells is regulated by STIM1. A, Western blotting shows expression of STIM1 in MIN6 cells and knockdown of STIM1 protein expression by shRNA-STIM1 (sh) relative to either wild-type MIN6 cells (wt) or cells with shRNA-scr (ss). Expression of STIM1 was reconstituted in the shRNA-STIM1 cells by transduction with an adenovirus expression vector encoding the human isoform of STIM1 (av). The left lane shows M_r markers and then duplicate lanes for each cell line. Anti-STIM1 is shown in the top part of the gel and anti-actin as a loading control in the lower part. B, stable MIN6 cell lines were treated using the protocol shown in Fig. 1A. Traces show only the records immediately before and after reintroduction of Ca²⁺ (indicated by open bar). SOCE in shRNA-STIM1 cells (sh) is inhibited relative to shRNA-scr (ss) and was partially rescued by adenoviral transduction with human STIM1 (av). Traces show the mean ± S.E. The peak SOCE amplitude (C) and AUC for the first 100 s after Ca²⁺ addition (D) were quantified for ss, sh, and av cells (28, 26 and 139 cells, respectively). Both peak and AUC were significantly reduced for sh cells relative to ss cells and were significantly recovered by av (*, p = 0.05; **, p < 0.01, ANOVA).

Characterization of Inward Currents Activated by STIM1 in β-Cells

After confirming that SOCE occurs in MIN6 cells by Ca²⁺ imaging, experiments to investigate ion currents were performed in the whole cell configuration using the standard bath solution and Cs⁺ pipette solution. Cells were transduced with an ER-targeted Ca²⁺ reporter to confirm that ER stores were depleted by dialysis with the base pipette solution. Under these conditions, an inward current was activated (Fig. 3A, upper trace) that followed depletion of ER Ca²⁺ stores (Fig. 3A, lower trace). This current reached peak amplitude within 5 min after break-in and showed slow inactivation. The inward current displayed a linear current-voltage relation that reversed at −12.7 ± 2.3 mV (n = 27 cells, Fig. 3B). This property is consistent with previous reports of a store-operated non-selective cation current in β-cells (5). A non-selective blocker of SOCE, SKF96365 (2, 8), reversibly inhibited this inward current (Fig. 3C) by 92.3 ± 2.6% (100 μm, n = 6 cells).

FIGURE 3.

Store depletion and activation of an inward current in MIN6 cells. A, membrane current and [Ca²⁺]_ER were measured simultaneously in cells expressing D1ER, a FRET-based genetically encoded indicator of ER Ca²⁺. Whole cell recording of membrane current with normal bath solution and Cs⁺ pipette solution designed to passively deplete ER stores is shown. The FRET ratio decreased, proportional to [Ca²⁺]_ER, immediately after break-in to the whole cell configuration. An inward membrane current started to develop within a few seconds and reached a peak after ∼1 min in this cell. The whole cell capacitance (C_m) of the cell illustrated was 4.2 pF. B, the current-voltage relation of the current after a voltage ramp from the holding potential of −60 mV to +40 mV at 1 V/s. This inward current has a linear current-voltage relation, and the mean reversal potential was −12.7 ± 2.3 mV (n = 27, −13.3 mV in the cell illustrated). C, extracellular application of the channel blocker SKF96365 (100 μm) reversibly inhibits the inward current. Break-in for this whole cell record occurred at the start of the trace. Shown is a representative example of six cells with a mean current inhibition of 92.3 ± 2.6% induced by SKF96365.

We determined ion selectivity of the inward current by substituting cations present in the extracellular solutions. Replacement of extracellular Na⁺ with N-methyl-d-glucamine (NMDG⁺) had no effect on the inward current amplitude (Fig. 4A), suggesting that the current is not carried through non-selective cation channels. Given that store depletion activates Ca²⁺ influx, we applied a Ca²⁺-free extracellular solution. This Ca²⁺-free solution increased, rather than inhibited, this current (Fig. 4B), suggesting that the current activated under these conditions is not a pure Ca²⁺ current but is consistent with extracellular Ca²⁺ inhibiting influx through K⁺ channels (23, 24).

FIGURE 4.

Inward currents through K_ATP channels. MIN6 cells were voltage-clamped at −60 mV in the whole cell configuration with a Cs⁺ pipette solution containing 200 μm EGTA. Under these recording conditions, with normal bath solution, an inward current developed, and the effects of ion substitution were tested. A, the current was not inhibited by substituting extracellular Na⁺ with NMDG⁺ (NMG, application indicated by filled bar). B, the inward current amplitude increased upon removal of extracellular Ca²⁺ (Ca-free, filled bar). C, raising extracellular K⁺ from 5.9 to 56 mm (56 K, filled bar) produced an increase in current amplitude (−30.9 ± 9.1 pA/pF at normal extracellular K⁺ and −247.2 ± 75.0 pA/pF at 56 mm K⁺, n = 7). D, increasing K⁺ produced a positive shift of the reversal potential for the current. In the cell illustrated, the reversal potential shifted from −16.1 mV to +15.9 mV and recovered to −14.3 mV, the mean change was from −17.6 ± 2.5 mV at normal (5.9 mm K⁺) to +13.0 ± 3.1 mV at 56 mm K⁺ and −15.2 ± 2.2 after returning to normal K⁺. I_m, inward current. E, substitution of extracellular Ca²⁺ for Ba²⁺ also inhibited the inward current. Current inhibition caused by a 10-s pulse of Ba²⁺ was 97.1 ± 1.2% (n = 7). F, the inward current was reversibly inhibited by 100 μm tolbutamide (Tolbut). G, increasing concentrations of ATP in the pipette solution inhibited the inward current (I_m). Data points were curve-fitted indicating an IC₅₀ of 0.6 mm (data from 9–13 cells at each concentration). H, inhibition of inward current (I_m) by 1 mm ATP was reversed by 1 mm ADP (C, control (no ATP), n = 13 cells; ATP, n = 9 cells; ATP+ADP, n = 9 cells).

The base pipette solution used in these studies is K⁺-free, and the extracellular K⁺ is 5.9 mm. Under these conditions the predicted K⁺ reversal potential is expected to approximate the observed reversal potential of the inward current. To determine whether the observed inward current could be carried by K⁺, we tested the effects of raising extracellular K⁺ to 56 mm. This increase of extracellular K⁺ induced a large increase in current amplitude (Fig. 4C) accompanied by a positive shift in the reversal potential of the current (Fig. 4D). The reversal potential shifted from −17.6 ± 2.5 mV to +13.0 ± 3.1 mV and recovered to −15.2 ± 2.2 mV (n = 7, measured using voltage ramps). This inward current was reversibly inhibited by substitution of extracellular Ca²⁺ with Ba²⁺ (Fig. 4E) and by 100 μm tolbutamide (Fig. 4F), suggesting involvement of K_ATP channels. This hypothesis was tested by adding different concentrations of ATP to the patch pipette solutions. The addition of ATP dose-dependently reduced inward current amplitudes with an IC₅₀ of 0.6 mm (Fig. 4G). The inhibitory effect of ATP (1 mm) was reversed by simultaneous addition of an equal concentration of ADP (Fig. 4H). These data suggest that this current is an inward current passing through K_ATP channels. The inhibitory effect of SKF96365 (Fig. 3C) is consistent with this suggestion as this compound was recently shown to reversibly inhibit K_ATP channels in smooth muscle cells (25).

Several studies have described SOCE in β-cells (17), suggesting that a SOC current is activated in MIN6 cells after ER Ca²⁺ store depletion. To unmask the potential contribution of a SOC current, we used a Cs⁺ pipette solution in combination with bath solutions containing either glyburide (1 μm), a sulfonylurea that blocks K_ATP channels, or with tetraethylammonium chloride (TEA; 140 mm, substituted for Na⁺) to block inward currents through K⁺ channels. In the presence of extracellular glyburide, an inward current was observed (Fig. 5A) with a reversal potential similar to that measured in the absence of glyburide (Fig. 5B). These data are inconsistent with a highly Ca²⁺-selective current conducted by Orai channels and suggest that if a SOC current is present, it was not adequately isolated by adding extracellular glyburide. However, in the presence of extracellular TEA, dialysis of wild-type MIN6 cells to deplete ER stores generated a small inward current with a mean current density of 0.5 ± 0.1 pA/pF (n = 7 cells); this current was inhibited by exposing the cells to a Ca²⁺-free bath solution (Fig. 5C). This small Ca²⁺ current is likely to represent activation of SOC channels in β-cells. Under these recording conditions, activation of the store-independent arachidonic acid-regulated Ca²⁺ (ARC)-channel current present in β-cells (26) is not expected to occur.

FIGURE 5.

Store-operated Ca²⁺ current in MIN6 cells. A, glyburide (1 μm) and nimodipine (10 μm) were added to the bath solution to block K_ATP and L-type VDCCs, respectively, in cells dialyzed with Cs⁺ pipette solution containing 5 mm EGTA. Under these conditions a small inward current developed. B, measurement of the reversal potential of the current shown in A indicates that this current is not Ca²⁺-selective. I_m, membrane current; V_m, membrane potential. C, using an extracellular solution containing TEA (substituted for Na⁺) to block inward K⁺ currents through K_ATP channels, a small inward current carried by Ca²⁺ was revealed. This current was inhibited by transient puffer applications of Ca²⁺-free bath solution, indicated by filled bars. The current illustrated is representative of seven cells with a mean current density of 0.5 ± 0.1 pA/pF.

The store-operated Ca²⁺ current (Fig. 5C) in shRNA-scr MIN6 cells was 0.2 ± 0.02 pA/pF (n = 11 cells). Knockdown of STIM1 expression in shRNA-STIM1 cells suppressed this current below the limit for reliable measurement. These data are consistent with the expression of a STIM1-regulated store-operated Ca²⁺-influx pathway in MIN6 cells similar to that described in other types of cells (27).

STIM1 Regulates K_ATP Currents in MIN6 Cells

Ca²⁺ imaging and electrophysiology data shows that knockdown of STIM1 using shRNA results in the predicted reduction of SOCE in MIN6 cells. However, STIM1 has previously been shown to interact with other target proteins including an inhibitory effect on Ca_V1.2 L-type Ca²⁺ channels (28–30) that is important for triggering insulin secretion (31, 32). To test for the potential effects of STIM1 on alternative targets important for regulating insulin secretion from β-cells, we measured membrane currents in cells stably expressing shRNA-STIM1 or control shRNA-scr. Knockdown of STIM1 reduced the amplitude of the K_ATP current by 54% (11.1 ± 1.6 pA/pF, n = 18 cells and 5.1 ± 1.1 pA/pF, n = 30 cells) for shRNA-scr (ss) and shRNA-STIM1 cells (sh), respectively (Fig. 6A). Reconstitution of STIM1 expression by AdV-hSTIM1 transduction restored this current, and current densities in shRNA-STIM1 cells and AdV-hSTIM1/shRNA-STIM1 (av) cells were 2.8 ± 1.3 pA/pF (n = 6 cells) and 18.8 ± 2.4 pA/pF (n = 8 cells), respectively (Fig. 6A). We detected no effect of shRNA-Stim1 on voltage-dependent Ca²⁺ currents (Fig. 6B) or Ca²⁺-dependent exocytosis (Fig. 6C).

FIGURE 6.

Effects of STIM1 knockdown on MIN6 cell membrane currents and Ca²⁺-dependent exocytosis. A, ShRNA-STIM1 (sh) reduced the amplitude of the inward current recorded with normal bath and Cs⁺ pipette solutions, as shown in Fig. 3, by ∼50% relative to shRNA-scr (ss) controls (*, p = 0.014; ss, n = 18 cells; sh, n = 24 cells), and this effect was reversed by adenoviral transduction of sh cells with human STIM1 (sh, n = 6 cells; sh + av, n = 8 cells; **, p < 0.001, ANOVA). B and C, STIM1 knockdown had no effect on the amplitude of voltage-dependent Ca²⁺ currents measured using the same solutions with voltage steps from −60 mV to 0 mV (ss, n = 7 cells; sh, n = 8 cells, not significantly different, ANOVA). A representative example of a record is shown in B, and quantification of peak current data for ss and sh cells is shown in C. D and E, exocytosis measured by capacitance tracking in cells dialyzed with 1.5 μm Ca²⁺ was also unaffected by STIM1 knockdown. A representative example of a capacitance record is shown in D, and quantification of the data for ss and sh cells shown in E (ss, n = 10 cells; sh, n = 8 cells; ns, not significantly different, ANOVA). The rate of exocytosis was determined by linear fitting of the data using Origin software. For the cell shown in D, the linear fit is illustrated and gave a y intercept of 2.05 pF, used to normalize data to cell size, and the slope was 0.76 fF/pF/s. C_m, whole cell capacitance.

STIM1 Protein Interacts with the Sulfonylurea Receptor 1 (SUR1) Subunit of K_ATP Channels

To further investigate the potential role of STIM1 in regulating K_ATP channels, we performed immunoprecipitation studies to determine whether there was a physical interaction between STIM1 and the K_ATP channel subunits SUR1 and Kir6.2. For these studies mouse islet protein extracts or lysates from HEK293T cells transfected with human STIM1(N131Q/N171Q) and FLAG-tagged SUR1 with or without Kir6.2 were used. The Asn-131/171 mutations in STIM1 prevent glycosylation and cell surface expression (33). Cell lysates were subject to immunoprecipitation (IP) using antibodies against SUR1, FLAG, or STIM1 with pulldown using protein A-agarose beads followed by Western immunoblotting (IB). All three lysates showed pulldown of STIM1 by the anti-SUR1 antibody (Fig. 7A). These data suggest an interaction between STIM1 and SUR1 as pulldown occurred in the absence of Kir6.2 in the HEK cell lysates (Fig. 7, C and D). This experiment was reversed with IP by anti-STIM1 and IB using anti-SUR1. Pulldown of SUR1 was observed in all three samples and was enhanced by poly-lysine (50 μg/ml) addition during the IP step (Fig. 7B). Incubation of cell lysates with anti-FLAG antibody is shown to pull down STIM1 in cells expressing FLAG-SUR1 but not in cells that did not express FLAG-SUR1 (Fig. 7C). Similarly, cell lysates treated with protein A-agarose beads alone did not pull down STIM1. These data demonstrate that the pulldown of STIM1 is not due to direct interaction of the FLAG antibody or of the beads with STIM1. Pulldown of STIM1 using anti-FLAG IP was observed in either the presence or absence of Kir6.2 (Figs. 7, A–D), suggesting that STIM1 interacts with SUR1.

FIGURE 7.

Interaction of STIM1 and K_ATP. A, IP studies using mouse islet (mIslet) extracts or HEK293T cell extracts expressing STIM1 and FLAG-SUR1 in the presence or absence of Kir6.2. IP was performed using anti-SUR1 antibody followed by IB with anti-STIM1.The SUR1 antibody pulled down STIM1 in all three samples. The input protein (IB only with anti-STIM1) is shown in the right three lanes. The band indicated as HC represents the heavy chain of the IP antibody, detected by the secondary antibody, and is only seen in the three IP samples. Co-immunoprecipitation was observed in two independent experiments using mouse islet or HEK cell proteins performing IP with anti-STIM1 and IB with anti-SUR1. B, IP was performed using anti-STIM1 and IB with anti-SUR1 using the same protein samples as in A, representative of two independent experiments with this protocol. The STIM1 antibody pulled down SUR1 in all three samples, and pulldown was enhanced by adding 50 μg/ml poly-lysine to the samples during IP. The SUR1 antibody exhibited a secondary band of slightly higher molecular weight than the heavy chain band, and subsequent experiments were performed using anti-FLAG to avoid any confounding effects of this band. LC represents the light chain of the IP antibody, and In is the mouse islet input protein. C, control IP studies using HEK293T cells expressing STIM1 in the presence or absence of FLAG-SUR1. IP performed with protein A-agarose beads (Beads) alone with samples expressing STIM1 and FLAG-SUR1 failed to pull down STIM1. IP with the FLAG antibody in cells expressing STIM1 but not FLAG-SUR1 also failed to pull down STIM1. IP using the FLAG antibody did, however, pull down STIM1 in cells expressing both STIM1 and FLAG-SUR1. STIM1 was detected by IB with anti-STIM1 along with nonspecific bands for the heavy-chain (HC) and light-chain (LC) of the FLAG antibody in the two FLAG-IP lanes but not the beads-only lane. IP of STIM1 using the FLAG antibody was observed in six experiments using four different protein samples. D, IP with the FLAG antibody pulled down STIM1 in the presence or absence of Kir6.2 under control conditions, and this pulldown was enhanced by the addition of 50 μg/ml poly-lysine (PolyLys). The image shown is representative of three independent experiments, and the increase in band density with poly-lysine was 27 ± 3% (n = 9, range 17 −44%) relative to control for cells expressing STIM1 and FLAG-SUR1 and 11 ± 1% (n = 6, range 8–16%) for cells expressing STIM1, FLAG-SUR1, and Kir6.2. Densities of STIM1 IB bands were significantly higher for poly-lysine-treated extracts relative to controls for these samples with STIM1 and FLAG-SUR1 (paired t test, p = 0.003) and also for samples with STIM1, FLAG-SUR1, and Kir6.2 (paired t test, p < 0.001). E, the STIM1 band density was 97 ± 2% (n = 6, range 92–102%, not significantly different, paired t test) relative to control for cells expressing STIM1 and FLAG-SUR1 and 94 ± 5% (n = 6, range 83–105%, not significantly different, paired t test) for cells expressing STIM1, FLAG-SUR1, and Kir6.2. F, the FLAG band density was significantly lower with poly-lysine (91 ± 6%, n = 10, range 75–133%, p = 0.04, three independent experiments, paired t test) relative to control for cells expressing STIM1 and FLAG-SUR1 and 83 ± 5% (n = 7, range 74–99%, p = 0.03, two independent experiments, paired t test) for cells expressing STIM1, FLAG-SUR1, and Kir6.2.

Poly-lysine is known to reverse the stimulatory action of phosphatidylinositol 4,5-bisphosphate (PIP₂) on K_ATP channels (34) and enhance sulfonylurea binding to SUR1 (35). PIP₂ also inhibits the binding of syntaxin-1A to SUR1, reversing syntaxin-1A mediated K_ATP channel inhibition (36). These studies led us to investigate whether poly-lysine might influence the interaction of STIM1 with SUR1 either through the effects on PIP₂ or by competing with the lysine-rich C terminus of STIM1 for binding. The interaction of STIM1 and FLAG-SUR1 was enhanced by the addition of 50 μg/ml poly-lysine to the IP reaction (Fig. 7D). This effect was more pronounced in studies with STIM1 and FLAG-SUR1 without Kir6.2 than in studies with Kir6.2 (3 independent experiments, mean band density 1.27 ± 0.03 higher with poly-lysine for samples expressing STIM1 and FLAG-SUR1 and 1.11 ± 0.01 higher for samples with STIM1, FLAG-SUR1, and Kir6.2). This enhancement is similar to the reported effect of poly-lysine to increase sulfonylurea binding to SUR1 by ∼30% (35). To determine whether the effect of poly-lysine was due to enhanced binding of the IB antibody to its target, we performed control Western blots using anti-STIM1 (Fig. 7E) or anti-FLAG (Fig. 7F) in the presence or absence of poly-lysine. Poly-lysine had no significant effect on anti-STIM1 binding and slightly, but significantly, decreased anti-FLAG binding. These data suggest that the enhanced IP signal in the presence of poly-lysine is due to a change in binding of STIM1 and SUR1.

To further investigate the interaction of STIM1 with SUR1, we transfected cells with STIM1 and a plasmid encoding the FLAG-tagged nucleotide binding fold-1 (FLAG-NBF1) region of SUR1. NBF1 has previously been shown to play a role in binding EPAC2 (37) and syntaxin-1A (38, 39). IP studies revealed that STIM1 and FLAG-NBF1 co-immunoprecipitated, suggesting that NBF1 forms at least one interaction site between STIM1 and SUR1 proteins (Fig. 8).

FIGURE 8.

STIM1 interacts with NBF-1 of SUR1. HEK293T cells were transfected with STIM1 and FLAG-NBF1 and IP performed using anti-FLAG (A) or anti-STIM1 (B). The data shown are representative of three independent IP experiments using four protein samples from different transfections. A, two protein samples from different transfections were collected and subject to IP using anti-FLAG followed by IB using anti-STIM1. The two left lanes show the results of IP, whereas the two right lanes show the corresponding input protein. The bands marked HC and LC that appear only in the IP lanes represent the heavy and light chains of the IP antibody. B, the same two protein samples as used in A were subjected to IP using anti-STIM1 followed by IB with anti-FLAG. IP lanes are shown on the left side of the panel, whereas the input protein is shown on the right. IP with anti-STIM1 pulled down FLAG-NBF1 and additional bands for the heavy and light chains of the IP antibody can be seen only in the IP lanes.

To determine whether shRNA-STIM1 has an off-target effect on expression of the K_ATP channel subunits SUR1 and inwardly rectifying K⁺ channel 6.2 (Kir6.2), we performed qPCR. SUR1 and Kir6.2 mRNA levels were not different in cells expressing shRNA-scr or shRNA-STIM1. In three independent experiments performed with triplicate samples in each, Kir6.2 expression in shRNA-STIM1 cells ranged from 92 to 117% of shRNA-scr controls, and SUR1 expression was 104–157% of controls. These data indicate that the reduction in inward current amplitude in shRNA-STIM1 cells was not due to decreased expression of K_ATP channel subunits and are consistent with an unexpected regulatory role for STIM1 in the control of K_ATP channel activity.

Discussion

Previous work in MIN6 cells as well as mouse and human primary β-cells demonstrated translocation and accumulation of fluorescently labeled STIM1 to punctae in subplasmalemmal ER after ER Ca²⁺ store depletion with thapsigargin or cyclopiazonic acid, inhibitors of SERCA (sarco(endo)plasmic reticulum calcium ATPase) pumps (18, 19). The role of endogenous STIM1 in β-cell signaling was not established in those studies. Here, we demonstrate that STIM1 regulates SOCE in MIN6 cells. We also discovered that STIM1 interacts with SUR1, a subunit of K_ATP channels, and functions in K_ATP channel activation. Binding of STIM1 to SUR1 was detected in transfected HEK293T cells as well as mouse islets of Langerhans. Our data indicate that STIM1 interacts with multiple proteins in β-cells that contribute to the regulation of β-cell Ca²⁺ signaling dynamics and membrane excitability. These findings suggest that STIM1 is a regulator of multiple signaling events in insulin-secreting cells.

It is known that SOCE in non-β-cells is regulated by direct interaction of STIM1 with the Ca²⁺ channel protein Orai1 and other regulatory proteins (27). Whether endogenously expressed Orai1 and STIM1 functionally interact and form the molecular basis of SOCE in β-cells has not been established. In view of previous reports that store depletion activates a non-selective cation current in β-cells (8), it seems unlikely that Orai1 would be the sole channel-forming subunit as these channels are highly Ca²⁺ selective when activated by STIM1 (40). Non-selective store-operated currents can be formed by an Orai1-TRPC1 complex regulated by STIM1 (41–45). An alternative possibility is that STIM1-Orai1 forms the SOCE mechanism and that the previously reported non-selective current represents the secondary activation of Ca²⁺-sensitive Trp channels. Ca²⁺-activated TRPM4 and TRPM5 channels have been shown to play a role in regulating insulin secretion (46–49). Expression of a Ca²⁺-selective SOCE mechanism distinct from previously reported non-selective cation currents would be consistent with the small currents we observe using whole cell recording.

An unexpected discovery from our experiments was that knockdown of STIM1 using shRNA reduced K_ATP channel activation. We considered the possibility that this represented an “off target” effect of the shRNA. However, reconstitution of K_ATP channel activation in shRNA-STIM1-expressing cells transduced with an adenovirus construct that encoded human STIM1 excludes a nonspecific effect of shRNA-STIM1. This suggests that STIM1 plays a role in regulating K_ATP channel activity. Quantitative PCR assays indicated that shRNA-STIM1 did not affect SUR1 or Kir6.2 expression, therefore, indicating it unlikely that STIM1 participates in K_ATP channel subunit biosynthesis. Nevertheless, our studies raise the prospect that loss of STIM1 expression either directly impacts K_ATP channel gating or indirectly modifies K_ATP channel activation via interaction with K_ATP channel regulatory proteins. A physical interaction between STIM1 and SUR1 mediated through NBF1 is suggested by our immunoprecipitation data, but it is not yet clear whether this is a direct interaction or indirect through an intermediary protein.

At least two mechanisms for direct interaction of STIM1 and SUR1 are possible based on the distinct mechanisms by which STIM1 regulates Orai and TRPC1. Orai1 is activated by interaction of the STIM-Orai activation region (SOAR) of STIM1 with cytosolic N- and C-terminal domains of Orai1 (50). In contrast, STIM1 activates TRPC1 through an electrostatic interaction of a KK amino acid pair in STIM1 with a DD pair in TRPC1 (51). In silico analysis of human SUR1 reveals a region with homology to the extended transmembrane Orai1 N-terminal (ETON) domain of Orai1 that interacts with STIM1 and also four DD pairs, including two pairs within NBF1. NBF1 is known to be important for MgADP activation of K_ATP (52). Additional experiments will be necessary to establish whether either of these sites plays a role in the molecular mechanism for STIM1-regulated K_ATP activity.

A number of STIM1 binding partners have been identified in addition to Orai and TRP channels (53, 54). Whether any of these alternative binding partners might contribute to an indirect effect on K_ATP channel regulation by STIM1 is unknown. One possibility is that actin binds STIM1 (53) and the actin cytoskeleton regulates K_ATP channels in β-cells (55, 56). Leptin is a hormone that activates K_ATP channels (55, 57), an effect mediated by disruption of actin filaments (55). Recent work has shown that Ca²⁺ influx through store-operated channels promotes actin depolymerization (58), raising the possibility that co-localization of STIM1 with K_ATP channels might promote local actin depolymerization and channel activation. Indeed, studies of STIM1 translocation in β-cells indicate aggregation of STIM1 punctae in actin-poor regions of the membrane (18). Thus, the effect of STIM1 on K_ATP might be an indirect effect of localized Ca²⁺ influx through Orai channels causing cytoskeletal actin disruption.

Whether the regulation of K_ATP activity is mediated by STIM1 in the ER, similar to the regulation of Orai1 and SOCE, or by plasma membrane-resident STIM1, similar to the ARC (arachidonic acid-regulated Ca²⁺) channel, is unknown at present. However, our immunoprecipitation studies indicate that a mutated STIM1, which is not glycosylated and inserted in the PM (33), and SUR1, expressed in the absence of Kir6.2, interact. These data suggest that STIM1 and SUR1 interact within the ER but do not exclude interplay of PM STIM1 with SUR1.

It is interesting to note that whereas reconstitution of the expression of STIM1 in MIN6 cells using human STIM1 restored K_ATP activity to above control levels, SOCE was only partially restored. Western blot analysis revealed that our rescue studies increased STIM1 protein levels above that seen in control cells (Fig. 6). A recent report indicates that high STIM1 expression can trap Orai1 intracellularly and prevent its trafficking into the plasma membrane (59). Our observation of partial rescue of SOCE may be attributable to an unfavorable ratio of STIM1 to Orai1 given that Adv-hSTIM1 raised expression of STIM1 protein to 7–12-fold higher than endogenous levels. K_ATP channels also undergo dynamic trafficking to the PM (60–63). Our data indicating that overexpression of STIM1 fully restored K_ATP activity but only partially restored SOCE might indicate that STIM1 does not affect K_ATP translocation in the same way as Orai1, if the trafficking trap mechanism applies in our studies. However, whether STIM1 influences K_ATP trafficking remains to be determined.

The K_ATP channel is well established as a key regulator of β-cell electrical excitability in response to nutritional status and hormonal inputs (57, 64, 65). Our data indicate that STIM1, through either direct or indirect interaction, regulates the activity of K_ATP channels in addition to regulating SOCE and plays a role in regulating β-cell excitability through these two processes. We propose a working model that incorporates a physiological role for STIM1 in regulating K_ATP channel activity and thereby β-cell membrane potential and Ca²⁺ oscillations. After β-cells are exposed to a high concentration of extracellular glucose, an initial fall in cytosolic Ca²⁺ concentration occurs (termed Phase 0) (66, 67). This phase of the Ca²⁺ response to glucose stimulation precedes β-cell depolarization and is caused by glucose-induced Ca²⁺ uptake into ER Ca²⁺ stores (67). Estimates of the resting [Ca²⁺]_ER in β-cells are 250–500 μm (68–70), a similar range to estimates for the K_D of Ca²⁺ binding to the EF-SAM domain of STIM1 (71, 72). This suggests that store filling induced by glucose elevation might decrease STIM1 activity and potentially play a role in regulating K_ATP. An increase in [Ca²⁺]_ER will induce translocation of STIM1 away from the plasma membrane to inactivate SOCE. A second possibility is that the rise in intracellular ATP will cause displacement of STIM1 from binding to SUR1, analogous to the effects of ATP on syntaxin binding (39). Displacement of STIM1 from SUR1 by ATP would contribute to the inhibition of K_ATP channels after a rise in blood glucose.

Our new observation that STIM1 knockdown inhibits K_ATP activation suggests that STIM1 has a positive effect on channel activity. Therefore, the translocation of STIM1 away from the plasma membrane or displacement of STIM1 from SUR1 might be predicted to reduce K_ATP activity and could contribute to channel inhibition after a rise in blood glucose. The importance of this mechanism relative to inhibition caused by the increase in the ATP/ADP ratio is unknown. However, STIM1-mediated changes in SOCE and K_ATP activity could contribute to the initiation of cell depolarization and [Ca²⁺]_c oscillations. In contrast, release of intracellular stores will cause translocation of STIM1 to PAM regions where it can interact with and activate Orai1 to refill stores and perhaps also interact with and activate K_ATP.

It has been demonstrated that SOCE activates type 8 adenylyl cyclase in both HEK 293 cells and MIN6 cells (73). This effect could lead to elevation of intracellular cAMP that is an important regulator of β-cell electrical activity and secretion mediated by protein kinase A and Epac2 (64, 74). A role for SOCE in regulating [Ca²⁺]_c oscillations in β-cells has been proposed as a consequence of ATP consumption and regulation of K_ATP activity (75). Metabolic oscillations and slow changes in K_ATP activity have been proposed to be important for glucose-induced oscillations (76). It is possible that slow changes in K_ATP activity as a consequence of store depletion and refilling and trafficking of STIM1 might contribute to normal bursting activity in islets.

In summary, we report that STIM1 is a multifunctional regulator of signaling dynamics in β-cells. In addition to defining the role of STIM1 as a regulator of SOCE in β-cells, our data revealed that STIM1 participates in K_ATP channel activation through an interaction with SUR1. This suggests a novel pathway for the control of β-cell excitability and function by interplay between ER Ca²⁺ stores and K_ATP channels.

Experimental Procedures

Cell Culture

MIN6 cells (passages 19–32) and HEK293T cells were cultured at 37 °C with 5% CO₂ in Dulbecco's modified Eagle's medium with 25 mm d-glucose (DMEM-H, Invitrogen) supplemented with 10% fetal bovine serum (Hyclone), 100 units/ml penicillin, and 10 μg/ml streptomycin (Gibco).

Cytoplasmic Ca²⁺ Imaging

[Ca²⁺]_c was measured using Fura 2 as previously described (77). MIN6 cells were grown on glass coverslips, and the cells were incubated for 15 min at 37 °C with 1 μm Fura 2 acetoxymethyl ester (Invitrogen), 0.02% (v/v) Pluronic acid in Krebs-Ringer buffer (KRBH). KRBH contained 119 mm NaCl, 25 mm NaHCO₃, 4.7 mm KCl, 1.2 mm MgSO₄, 2.5 mm CaCl₂, 1.2 mm KH₂PO₄, 10 mm HEPES-NaOH (pH 7.4), and 2 mm d-glucose. All solutions were made using ultrapure water. After loading, cells were transferred onto the temperature-controlled stage of an inverted microscope (Nikon 2000u, Nikon Ti or Olympus IX81) and visualized with a 40× superfluor oil immersion objective. Cells were superfused continuously at 37 °C with KRBH. Fura 2 excitation wavelengths were 340 and 380 nm, and the fluorescence emission was detected at 510 nm. The ratio of the emission intensities of Fura 2 excited at 340 and 380 nm (R340/380) were recorded using MetaFluor software (Molecular Devices) for image acquisition and analysis (78).

[Ca²⁺]_c measurement in some studies was performed using a FlexStation 3 plate reader in 96-well plate format using solutions and wavelengths for recording as described above. Test solutions were injected into individual wells containing MIN6 cells at 60–80% confluence. Experimental parameters and data acquisition were controlled using SoftMax Pro 5.4.6 software (Molecular Devices).

Electrophysiology

Whole cell recordings of membrane currents were made at room temperature (21–24 °C) using an EPC-9 amplifier controlled by PatchMaster software (HEKA Electronik, Lambrecht/Pfalz, Germany). Electrodes were pulled from patch clamp glass capillaries (catalog no. G85150T-4, Warner Instruments, Hamden, CT) using a Sutter P97 puller. MIN6 cells were bathed in KRBH. Isosmotic substitution of NaCl (NMDG-Cl or KCl) was used in some experiments, or CaCl₂ was omitted (Ca⁺-free, 10 μm EGTA) or substituted with BaCl₂ (in this Ba²⁺ solution, MgSO₄ was replaced with MgCl₂). For some experiments a TEA bath solution comprised 140 mm TEA-Cl, 10 mm CaCl₂, 1.2 mm MgCl₂, and 10 mm HEPES (pH 7.4). Glucose was added to these solutions as indicated. Patch pipettes were filled with a base solution containing 90 mm Cs₂SO₄, 10 mm NaCl, 1 mm MgCl₂, 5 mm HEPES, and 0.2 mm EGTA, pH adjusted to 7.4 with CsOH. ATP, ADP, and GTP were added to the pipette solution as indicated. Cells were held at −60 mV for current recording, and voltage ramps were applied to measure reversal potentials. PatchMaster software compensated junction potentials and was used to generate voltage ramps at 1 V/s, the software was also used to measure cell capacitance to normalize current amplitudes to cell size. Whole cell capacitance measurements of Ca²⁺-dependent exocytosis were made using the LockIn module of PatchMaster with an applied sine wave of 10 mV amplitude at 1 kHz and a pipette solution containing (in mm) 125 cesium glutamate, 10 NaCl, 10 KCl, 1 MgCl₂, 10 EGTA, 11 CaCl₂, 3 MgATP, 0.2 LiGTP, and 5 HEPES (pH 7.4). Extracellular test solutions were applied by pressure ejection from glass micro-capillaries using a PicoSpritzer III (Parker Instrumentation). Experiments with fluorescence imaging used a PTI DeltaRam monochromator light source controlled with MetaFluor software. For FRET experiments, emission images at 485 and 535 nm were captured simultaneously using a DualView beam splitter and MetaFluor software.

Quantitative Real-time PCR

Total RNA was isolated and purified from two to four replicate plates of each cell type using an RNeasy Mini kit (Qiagen) according to the manufacturer's instructions. RNA (1 μg) was reverse-transcribed to cDNA using oligo(dT₁₅) primers according to the manufacturer's instructions (Promega). Primer sequences for S18 were 5′-GCCATCACTGCCATTAAGGG and 5′-CCAGTCTGGGATCTTGTACTG and for STIM1 were 5′-GAAGCAAAATGCAGAGAGGC and 5′-CATCATCCAGGGAAGAGCTG. Primers for Kir6.2 were 5′-GTGTTCACCACGCTGGTGGAC and 5′-CAGGTCACCGTGGGCGAAGG. Primers for SUR1 were 5′-GGACAGAAGATCGGGATCTGC and 5′-GAAGGAGGACTTCCCACTGCC. Each primer was used at a concentration of 500 nm in a 25-μl reaction volume containing Brilliant II SYBR Green qPCR master mix (Agilent Technologies). Quantitative real-time PCR was completed in a Stratagene MX3000P real-time PCR thermal cycler under the following conditions: initial hold at 95 °C for 10 min followed by 40 cycles of 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s. Fluorescence intensity data were collected during each cycle at 72 °C. Amplicon dissociation was initiated immediately after the final PCR extension, with a 30-s hold at 72 °C followed by increasing the temperature to 95 °C with 0.25 °C steps. One cDNA was produced from each RNA sample, and each sample was tested in duplicate by qPCR. C_q values were averaged for duplicates, and relative quantification calculations were completed as described by Pfaffl (79).

Inhibition of STIM1 Expression Using shRNA

Cells in which STIM1 expression was transiently or stably deleted were used. For transient suppression of STIM1, MIN6 cells were transfected with a plasmid vector that expressed small interfering RNA (siRNA) targeted to STIM1. MIN6 cells stably expressing short hairpin RNA targeted to STIM1 (shRNA-STIM1) or shRNA containing a scrambled sequence that targeted no known genes (shRNA-scr) were produced by transduction with lentiviral particles (Santa Cruz Biotechnology). Stably transduced cells were maintained in DMEM-H medium containing 2 μg/ml puromycin. In some experiments, STIM1 expression was reconstituted by transducing cells with an adenoviral vector to express the human isoform of STIM1 (AdV-shRNA-hSTIM1) (ViraQuest Inc.).

Western Immunoblotting of Endogenous MIN6 Protein

Western blotting was used to supplement qPCR to assess the effectiveness of STIM1 knockdown and restoration in cells stably expressing shRNA-STIM1. Whole-cell protein lysates were collected from two replicate plates of MIN6 shRNA-STIM1, shRNA-STIM1/AdV-hSTIM1, shRNA-scr, and control, wild-type cells. Lysis buffer contained 69 mm Tris-HCl, 1% SDS, 10% glycerol, and 0.04% protease inhibitor mixture (Roche Applied Science) (pH 6.8). Lysates were immediately frozen and stored at −80 °C until use. Samples were prepared for SDS-PAGE by adding β-mercaptoethanol (10% v/v) to 50 μg of protein and then heated to 95 °C. SDS-PAGE was performed on a 4–20% gradient gel (Bio-Rad) at 130 V. Proteins were transferred to PVDF membranes (Immobilon-P, Millipore) and blocked in Tris-buffered saline with Tween 20 (TBST, 20 mm Tris-Base, 136 mm NaCl, 0.1% Tween 20, pH 7.4) buffer containing 5% nonfat dry milk for 45 min. Membranes were incubated overnight with gentle rocking at 4 °C in anti-STIM1 antibody (1:250; BD Biosciences catalog no. 610954, lot #5199779) or anti-β-actin antibody (1:1000; Sigma catalog no. A5316, lot # 119K4765) in nonfat dry milk. The blots were incubated for 1 h at room temperature in horseradish peroxidase (HRP)-conjugated anti-mouse IgG secondary antibody (detailed below) diluted 1:1000 plus Streptactin-HRP (Bio-Rad) diluted 1:6600. Blots were developed using the SuperSignal West Pico Chemiluminescence Substrate kit (Thermo Scientific) for 5 min, and protein bands were detected using a Bio-Rad Gel Documentation system with QuantityOne software. Band intensities were quantified to determine protein knockdown efficiency using Quantity One software (Bio-Rad Laboratories).

Co-immunoprecipitation and Western Blotting of HEK293T Cell Lysates

HEK293T cells were transfected with plasmids (1 μg/10 cm plate) for human STIM1(N131Q/N171Q), FLAG-SUR1, FLAG-NBF1, and Kir6.2 using Lipofectamine 2000 according to manufacturer's instructions. After 2 days cells were lysed with 0.5 ml ice-cold radioimmune precipitation assay buffer (50 mm Tris-Base, 150 mm NaCl, 1% IGEPAL, 0.5% sodium deoxycholate, 0.1% SDS (pH 8.0)) with 1 mm dithiothreitol, 25 μm pepstatin A, 0.2 μm soybean trypsin inhibitor, and 1 mm Perfabloc SC added fresh before use. The lysate was collected and centrifuged at 13,000 rpm (Eppendorf 5415D microcentrifuge) for 10 min at 4 °C. The supernatant was transferred into a clean tube, and protein concentration was determined using a Bio-Rad DC protein assay kit. Protein samples were stored at −80 °C until use.

For immunoprecipitation, 100 μg of protein lysate was incubated for 16 h at 4 °C with antibodies (1–5 μg) as indicated. Then the antigen-antibody complex was immobilized on protein A-agarose beads (20 μl of 50% beads aqueous slurry, Thermo Scientific) for 3 h at 4 °C. The beads were washed 5 times with 250 μl of CHAPS lysis buffer (50 mm Tris-Base, 150 mm NaCl, 1 mm EDTA, 1% CHAPS (pH 8.0)) containing 10% β-mercaptoethanol and pelleted by centrifugation at 13,000 rpm for 30 s at 4 °C. The beads were resuspended with 20 μl of 2× SDS sample buffer (final concentrations: 25 mm Tris, 192 mm glycine, 1% SDS (pH 8.3)), and 20 μl of radioimmune precipitation lysis buffer. The samples were then centrifuged for 60 s at 13000 rpm, and the supernatant was used for Western blotting. Primary antibodies used were anti-SUR1 (Abcam catalog no. S289-16, Lot # GR-103710-18), anti-FLAG, clone M2 (Sigma catalog no. F1804, lot # SLBN5629V), and anti-STIM1 (as described for the Western blots above). Antibody specificity was assessed in HEK293T cells by comparing Western blots of control cells with cells transfected with the appropriate plasmid to induce protein expression.

The immunoprecipitated protein samples (∼30 μg of protein) and molecular weight markers (Precision Plus All Blue Protein Standard, Bio-Rad catalog no. 161–0373) were then resolved by SDS-PAGE (4–20% gradient protean GTX mini gels, Bio-Rad) and wet-transferred to Immobilon-P PVDF membranes (Millipore) using a buffer containing 25 mm Tris-base, 192 mm glycine, and 20% methanol (pH 8.3). The blots were washed once with TBST and incubated for 45 min at room temperature with blocking buffer (TBST with 5% blotting grade nonfat skim milk (Bio-Rad); membranes were incubated with specific antibodies (1:1,000 dilution in blocking buffer) for 16 h at 4 °C followed by 3 washes with TBST and incubation with HRP-coupled anti-mouse secondary antibody (Abcam catalog no. ab6728, lot # GR152005-0; 1:10,000 dilution) for 1 h at room temperature. After 3 washes with TBST, membranes were incubated with SuperSignal West Pico Chemiluminescence Substrate kit (Thermo Scientific) for 5 min, and protein bands were detected using a Bio-Rad Gel Documentation system with QuantityOne software. For quantification of band density, background-corrected measurements were made on images with no saturated pixels from three different exposure times to test for linearity.

Statistical Analysis

Student's t test or ANOVA was used for intergroup comparisons (p < 0.05 was considered statistically significant) as detailed in the figure legends. Data traces are plotted as the mean ± S.E. Box and whisker plots show mean (solid circle), median (dashed line), 25 and 75% values (box), and minimum/maximum values (whiskers).

Author Contributions

C. A. L., R. F. K., H. A. N., J. N., and M. W. R. conducted the experiments, analyzed the results, and edited the manuscript. C. A. L. conducted the patch clamp electrophysiology experiments. C. A. L., R. F. K., and M. W. R. conducted the calcium imaging experiments. R. F. K. performed qPCR. H. A. N. and J. N. performed immunoprecipitation and Western blotting. M. W. R. conceived the idea for the project and wrote the paper with C. A. L.