Role for Plasma Membrane-Related Ca²⁺-ATPase-1 (ATP2C1) in Pancreatic β-Cell Ca²⁺ Homeostasis Revealed by RNA Silencing

Rat β-cell-derived INS1 cells (25) were cultured in RPMI-1640 medium (Gibco BRL) containing 11 mmol/l glucose and 2 mmol/l glutamine, supplemented with 10% (vol/vol) fetal bovine serum (FBS), 100 units/ml penicillin, 100 μg/ml streptomycin, 1 mmol/l sodium pyruvate, and 50 μmol/l β-mercaptoethanol. Mouse β-cell-derived MIN6 cells (26) were cultured in Dulbecco’s modified Eagle’s medium (Sigma) containing 25 mmol/l glucose and 2 mmol/l pyruvate, supplemented with 15% (vol/vol) FBS, 4 mmol/l glutamine, 100 units/ml penicillin, 100 μg/ml streptomycin, and 50 μmol/l β-mercaptoethanol. For [Ca²⁺] measurements with recombinant targeted aequorins (see below), cells were seeded onto 13-mm diameter poly-l-lysine-coated glass coverslips and grown to 50–80% confluency. Cells were then infected with adenoviruses encoding untargeted cytosolic aequorin (Cyt.Aq) (27) or aequorin targeted to either the secretory vesicle matrix by fusion with vesicle-associated membrane protein-2 (VAMP2)/synaptobrevin (VAMP.Aq) (7) or the ER lumen (ER.Aq) (28), at a multiplicity of infection of 30 infectious units per cell. Measurements of aequorin bioluminescence were performed 48 h after infection using a purpose-built photomultiplier system, as described previously (7,29). Rat islets were isolated and cultured as described (30).

Total RNA was extracted from cell lines or rat tissue using TRI reagent (Sigma) according to the manufacturers’ instructions and then reverse-transcribed using M-MLV reverse transcriptase (Promega). PCR amplification was performed with primers designed to amplify a 640-bp fragment of PMR1 corresponding to nucleotides 327–967 of RAT PMR1 mRNA (31): forward, 5′-GTTAGTCATAGGCGAGC-3′; reverse, 5′-GTCCATGCTCTTCTGCAG-3′. The full coding region of mouse PMR1 and a portion of the 3′ untranslated region (nucleotides 179-3016) (31) was amplified from MIN6 cells by PCR: forward primer, 5′-ATGAAGGTTGCACGATTTC-3′; reverse primer, 5′-CCTCTTGAACTTGCCCTTC-3′.

Rat islets (∼150) or clonal β-cells were extracted into radioimmunoprecipitation assay buffer, comprising PBS supplemented with 1% (vol/vol) Nonidet P40, 0.5% (wt/vol) sodium deoxycholate, 0.1% (wt/vol) SDS, 1 μmol/l phenylmethylsulfonyl fluoride (PMSF), 5 μg/ml aprotinin, and 5 μg/ml leupeptin. For the preparation of clonal β-cell membrane fractions, cells were scraped into ice-cold homogenization buffer (0.25 mol/l sucrose, 1 mmol/l EDTA, 20 mmol/l HEPES, 1 μmol/l PMSF, 5 μg/ml aprotinin, and 5 μg/ml leupeptin, pH 7.4) and then homogenized with a Teflon homogenizer on ice. Fractions were collected by differential centrifugation at 4°C as follows: 200g for 10 min, whole cell/nuclear fraction; 3,000g for 10 min, heavy mitochondrial fraction; 16,000g for 10 min, light mitochondrial fraction; 100,000g for 1 h, microsomal fraction; and supernatant from 100,000g spin, cytosolic fraction.

Rabbit polyclonal antiserum was raised to a 17-amino acid peptide [(C)RRAFHGWNEFDISEDE_amide] corresponding to a 100% homologous amino-terminal region of rat and human PMR1/ATP2C1 (16,31). Rabbits (New Zealand white) were immunized with keyhole limpet hemocyanin-conjugated peptide, as previously described (32), and the specificity was confirmed by immunoblot analysis (see below). For antibody purification, non-IgG proteins were precipitated from the antiserum with caprylic acid (33), and the IgG fraction was precipitated using ammonium sulfate.

Protein samples were resolved by SDS-PAGE on 5–7.5% (wt/vol) polyacrylamide gels and transferred onto Immobilon-P transfer membrane (Millipore) following a standard protocol. Membranes were probed with primary antibodies, as stated in the figure legends. Immunostaining was revealed with horseradish peroxidase (HRP)-conjugated secondary antibodies using an enhanced chemiluminescence detection system (Roche Diagnostics).

A small interfering RNA (siRNA) duplex corresponding to nucleotides 337–357 of rat PMR1 cDNA (31) was generated (Dharmacon Research). This region showed no significant homology to any other known gene (analyzed using BLAST) (34). The siRNA duplex consisted of a 21-nucleotide sense strand (5′-GUUAGUCAUAGGCGAGCCUdTdT-3′) and a 21-nucleotide antisense strand (5′-AGGCUCGCCUAUGACUAACdTdT-3′), paired in a manner to have a 19-nucleotide duplex region with a 2-nucleotide dithymidine overhang at each 3′ terminus. A scrambled siRNA (sense strand, 5′-CGUGAUUGCGAGACUCUGAdTdT-3′; antisense strand, 5′-UCAGAGUCUCGCAAUCACGdTdT-3′), which showed no significant homology to any protein known (analyzed using BLAST), was used as a control.

siRNAs were introduced into INS1 or MIN6 cells by lipid-mediated transfection using Oligofectamine (Invitrogen). For each 2-cm² culture dish, lipid-siRNA complexes were formed using 60 pmol siRNA and 3 μl Oligofectamine in 100 μl Optimem1 serum-free medium (Invitrogen). Lipid complexes were then added to cells that were previously seeded in 400 μl antibiotic-free growth medium. After 24 h, the medium was supplemented with 500 μl growth medium. To measure secretory vesicle, ER, or cytosolic Ca²⁺ concentrations in cells transfected with control (scrambled) or PMR1 siRNAs, cells were infected with VAMP.Aq-, ER.Aq-, or Cyt.Aq-encoding adenovirus 24 h after siRNA transfection. Ca²⁺ measurements were carried out 24 h after adenoviral infection.

Where indicated, cells were depleted of Ca²⁺ by incubation with ionomycin (10 μmol/l), monensin (10 μmol/l), and cyclopiazonic acid (10 μmol/l) in modified Krebs-Ringer bicarbonate buffer (KRBB, 140 mmol/l NaCl, 3.5 mmol/l KCl, 0.5 mmol/l NaH₂PO₄, 0.5 mmol/l MgSO₄, 3 mmol/l glucose, 10 mmol/l HEPES, and 2 mmol/l NaHCO₃, pH 7.4), supplemented with 1 mmol/l EGTA, for 10 min at 4°C (7). Aequorin was reconstituted with 5 μmol/l coelenterazine (Cyt.Aq) or 5 μmol/l coelenterazine n (VAMP.Aq and ER.Aq) for 1–2 h at 4°C in KRBB supplemented with 1 mmol/l EGTA.

Intact cells were perifused with KRBB supplemented with additions as stated, at a flow rate of 2 ml/min in a thermostatted chamber (37°C) in close proximity to a photomultiplier tube (ThornEMI) (35). Where indicated, cells were permeabilized with 20 μmol/l digitonin (Fluka) for 1 min at 37°C and subsequently perifused in intracellular buffer (140 mmol/l KCl, 10 mmol/l NaCl, 1 mmol/l KH₂PO₄, 5.5 mmol/l glucose, 2 mmol/l MgSO₄, 1 mmol/l ATP, 2 mmol/l sodium succinate, and 20 mmol/l HEPES, pH 7.05), with additions as stated in the figure legends. At the end of all experiments, cells were lysed in a hypotonic Ca²⁺-rich solution (100 μmol/l digitonin and 10 mmol/l Ca²⁺ in H₂O) to discharge the remaining aequorin pool for calibration of the aequorin signal (7).

MIN6 cells were incubated for 16 h in full growth medium containing 3 mmol/l glucose. Cells were then loaded with 5 μmol/l fluo-3-acetoxymethylester (fluo-3-AM; Sigma) for 40 min at 37°C in KRBB containing 3 mmol/l glucose. [Ca²⁺]_cyt changes in individual cells were imaged after stimulation, as stated in the figure legends, using an Olympus IX-70 inverted microscope (×40 objective lens). Cells were excited at 480 nm at 5-s intervals using a Till photonics monochromator, and emission signals were detected at 515 nm with an Imago SensiCam cooled charge-coupled device camera.

MIN6 cells were incubated in full growth medium containing 3 mmol/l glucose for 16 h and then incubated in KRBB supplemented with 3 mmol/l glucose for 15 min at 37°C. The medium was removed, and then the cells were stimulated for a further 40 min at 37°C with KRBB plus additions, as stated in the figure legends. Released and total insulin were measured by radioimmunoassay (36).

Free [Ca²⁺] was calculated using the METLIG program (37). Data represent the means ± SE of at least three separate experiments. Statistical analysis was performed using Student’s t test.

To determine whether PMR1 mRNA is present in primary rat islets or clonal β-cells, PCR amplification was performed using primers designed to amplify a 640-bp internal region of rat PMR1 cDNA (31). A fragment of the correct size was amplified from islet, INS1-derived (rat), and MIN6-derived (mouse) cDNA (Fig. 1A). The mRNA encoding mouse PMR1 was subsequently cloned from MIN6 cells (see online appendix [available at http://diabetes.diabetesjournals.org]). In extracts from either rat islets or clonal β-cells, the presence of the 105-kDa PMR1 protein was revealed by immunoblotting with a polyclonal antibody raised to the amino terminus of PMR1 (Fig. 1B). Furthermore, immunoblotting of subcellular fractions separated by differential centrifugation revealed a widespread distribution of PMR1 on both INS1 (Fig. 2A) and MIN6 (Fig. 2B) cell membranes. As predicted, PMR1 immunoreactivity was enriched in microsomal fractions, identified by the presence of the type II ryanodine receptor (RyR II) (Fig. 2A) (8). In addition, a substantial proportion of PMR1 immunoreactivity from either cell type was detected in a heavy mitochondrial fraction that was also rich in dense-core secretory vesicles (see measurements of insulin content in Fig. 2A). However, suggesting that PMR1 was largely confined to ER-, Golgi-, and dense-core vesicle-associated membranes, we were unable to detect any labeling of plasma membrane or mitochondrial structures by immunocytochemical analysis of overexpressed c-myc-PMR1, but there was substantial overlap with markers for the secretory pathway (results not shown).

To investigate the role(s) of PMR1 in β-cells, endogenous PMR1 protein expression was reduced by RNA silencing, and the effect on intracellular Ca²⁺ handling was monitored using recombinant targeted aequorins (38). Transfection of an siRNA duplex corresponding to nucleotides 337–357 of rat PMR1 cDNA (PMR1 siRNA) (31) reduced PMR1 expression in INS1 cells by 65.3 ± 11.9 and 85.7 ± 4.7% (n = 3, P < 0.01) after 24 and 48 h, respectively, consistent with the efficient transfection of the duplex RNA into at least 85% of the cell population. PMR1 levels remained low until 72 h after transfection, when expression increased to 70.3 ± 4.4% (n = 3, P < 0.01) of the control level (Fig. 3A). After PMR1 siRNA transfection, a similar reduction in PMR1 expression was observed in MIN6 cells, indicating that the chosen siRNA was also effective for depleting mouse PMR1 mRNA (Fig. 3A). This silencing effect appeared to be selective because PMR1 siRNA transfection of INS1 cells did not affect SERCA2 or mannose 6-phosphate receptor (m6PR) protein expression (Fig. 3B). Conversely, PMR1 expression was unaffected after transfection with a control (scrambled) siRNA (Fig. 3A).

Previous studies in both yeast (19) and mammalian cells (20) have revealed a role for PMR1 in controlling Ca²⁺ influx. We explored the effect of PMR1 silencing on Ca²⁺ influx in intact INS1 β-cells by monitoring changes in [Ca²⁺]_cyt when Ca²⁺ was reintroduced to Ca²⁺-depleted cells, a condition in which store-operated Ca²⁺ channels are activated (7). The rate of increase of [Ca²⁺]_cyt and the initial [Ca²⁺]_cyt peak were markedly increased in cells treated with PMR1 siRNA (Fig. 4A, Table 1), and [Ca²⁺]_cyt remained significantly higher in PMR1-depleted cells for ∼1 min before returning to control levels.

In yeast pmr1-null mutants, one of the essential components of the high-affinity Ca²⁺ influx system stimulated by store depletion has been identified as a homologue of the α1 pore-forming subunit of voltage-gated Ca²⁺ channels (19). To determine whether voltage-gated Ca²⁺ channels might be involved in the elevation of [Ca²⁺]_cyt observed in PMR1-depleted β-cells, CaCl₂ was reintroduced to Ca²⁺-depleted cells in the presence of the voltage-gated Ca²⁺ channel inhibitor nimodipine. Under these conditions, the elevation of [Ca²⁺]_cyt previously observed in PMR1-depleted cells was completely abolished, and the peak value achieved (∼1.7 μmol/l) was identical in cells treated with either scrambled or PMR1 siRNAs (Fig. 4B, Table 1). Interestingly, in the presence of nimodipine, the initial [Ca²⁺] peak was not followed by an elevated [Ca²⁺] plateau that is normally observed in the absence of the drug (Fig. 4A versus 4B), indicating that this sustained elevation of [Ca²⁺]_cyt is dependent on Ca²⁺ influx through l-type Ca²⁺ channels.

These results indicated that Ca²⁺ entry via voltage-gated Ca²⁺ channels was largely responsible for the enhancement of Ca²⁺ entry in PMR1-depleted β-cells. To determine whether the enhanced increases in [Ca²⁺]_cyt after Ca²⁺ readmission to PMR1-depleted cells may be attributable to alterations in the expression or activity of voltage-gated Ca²⁺ channels or to other factors such as alterations in membrane potential, we monitored increases in [Ca²⁺]_cyt provoked by cell depolarization with 50 mmol/l KCl. Neither the peak nor sustained [Ca²⁺]_cyt values were affected by PMR1 depletion (Fig. 4C), arguing against a role for changes in the activity or total number of voltage-gated Ca²⁺ channels. Correspondingly, PMR1 depletion had no effect on levels of mRNA encoding the l-type Ca²⁺ channel α1 subunit (data not shown).

After the initial peak of [Ca²⁺]_cyt after the reintroduction of CaCl₂ to control or PMR1 siRNA-treated cells, [Ca²⁺]_cyt returned to basal levels after ∼1 min, a process likely to involve extrusion of Ca²⁺ across the plasma membrane and pumping of the ion into intracellular stores. To examine the effect this Ca²⁺ accumulation may have on organelle lumen Ca²⁺ concentrations, we monitored changes in secretory vesicle and ER Ca²⁺ concentration in intact cells. The rate and extent of ER Ca²⁺ uptake were markedly increased (Fig. 4E, Table 1) after PMR1 silencing, presumably reflecting the increase in [Ca²⁺]_cyt observed under these conditions (Fig. 4A), whereas vesicular [Ca²⁺] was unaffected (Fig. 4D, Table 1). Accordingly, the augmented ER [Ca²⁺] observed in PMR1-depleted cells was completely abolished by the presence of nimodipine (Fig. 4F, Table 1).

Because the above measurements in intact cells were complicated by changes in [Ca²⁺]_cyt after PMR1 depletion, we next explored changes in organellar [Ca²⁺] using digitonin-permeabilized cells in which [Ca²⁺]_cyt could be clamped at will. When perifusate [Ca²⁺] was increased from <1 to 400 nmol/l, the initial rate of [Ca²⁺] increase in both the secretory vesicles (3.36 ± 0.42 vs. 2.33 ± 0.14 μmol · l⁻¹ · s⁻¹ for control vs. PMR1⁻ cells, n = 3, P < 0.05) (Fig. 4G) and the ER (3.62 ± 0.48 vs. 2.00 ± 0.28 μmol · l⁻¹ · s⁻¹, n = 3, P < 0.01) (Fig. 4H) was found to be significantly inhibited in PMR1-depleted compared with control cells, whereas steady-state Ca²⁺ concentrations were not significantly different in the two groups.

We next investigated the effect of PMR1 silencing on insulin secretion stimulated by glucose. MIN6 cells were used for these experiments because of their greater responsiveness to nutrient secretagogues compared with INS1 cells (39). The stimulation of insulin secretion in response to 30.0 mmol/l (vs. 3.0 mmol/l) glucose was significantly increased by PMR1 depletion (4.3 ± 0.8 vs. 6.3 ± 0.8-fold stimulation for control vs. PMR1⁻ cells, n = 4, P < 0.05) (Fig. 5A). In contrast, basal insulin secretion (3.0 mmol/l glucose) or secretion stimulated by a submaximal glucose concentration (11.0 mmol/l) were unaffected by PMR1 silencing (Fig. 5A).

From the data shown in Fig. 4, we inferred that the enhancement of glucose-induced insulin secretion observed in PMR1-depleted cells may be attributable to increased Ca²⁺ influx. However, measurements with Cyt.Aq revealed that after stimulation with 30.0 mmol/l glucose, average [Ca²⁺]_cyt was unaffected by PMR1 silencing (data not shown). Therefore, we explored the possibility that PMR1 depletion enhances glucose-induced insulin secretion by modifying spatial or temporal aspects of the evoked [Ca²⁺] increases. In the additional presence of tetraethylammonium (TEA), an inhibitor of Ca²⁺- and voltage-activated K⁺ channels (40) that amplifies glucose-induced [Ca²⁺] oscillations in single MIN6 cells (39), there was no significant difference in the average [Ca²⁺]_cyt between control and PMR1-depleted cells: 43.81 ± 6.37 vs. 50.19 ± 6.72% change for control vs. PMR1⁻ cells, n = 10 cells (Fig. 5C), corresponding to a [Ca²⁺]_cyt of ∼330 nmol/l (basal values ∼200 nmol/l), as measured in single cells using fluo-3-AM and in cell populations using Cyt.Aq (data not shown). In contrast, PMR1 depletion appeared to transform the [Ca²⁺] changes into slow oscillations, with rapid oscillations superimposed at the peak but not at the nadir (Fig. 5B). The amplitude of these slow oscillations was not affected by PMR1 silencing (126.8 ± 9.8 vs. 133.23 ± 9.1% change for control vs. PMR1⁻ cells, n = 10 cells) (Fig. 5D). However, oscillation duration (29.61 ± 1.4 vs. 71.28 ± 10.7 s for control vs. PMR1⁻ cells, n = 10 cells, P < 0.001) (Fig. 5E) and decay rate on return to nadir (4.33 ± 0.27 vs. 7.17 ± 1.01 s, control vs. PMR1⁻ cells, n = 10 cells, P < 0.05) (Fig. 5F) were significantly increased. Correspondingly, insulin secretion was significantly enhanced in PMR1-depleted cells when stimulated under the same conditions (11 mmol/l glucose + 10 mmol/l TEA; 5.9 ± 1.5 vs. 10.9 ± 2.5-fold stimulation for control vs. PMR1⁻ cells, n = 4, P < 0.05) (Fig. 5A).

We show here that PMR1/ATP2C1 is expressed in both pancreatic islets and clonal β-cells derived from both rat and mouse (Fig. 1). In contrast to previous studies that have shown a predominantly Golgi localization of endogenous PMR1 in yeast (14) and overexpressed PMR1 in mammalian cell lines (20,41), the present study reveals the presence of endogenous PMR1 in two distinct fractions containing either ER (RyR II) or secretory vesicle (insulin) markers (Fig. 2). In agreement with this distribution, we demonstrate that inactivation of the pump with selective siRNAs has marked effects on Ca²⁺ handling by both the ER and secretory vesicles in these cells. Importantly, these findings demonstrate that PMR1 plays a significant role in the accumulation of Ca²⁺ by the ER (Fig. 4H) and may underlie the thapsigargin-independent uptake of Ca²⁺ into this compartment (7,42,43), which represents 15–20% of the total Ca²⁺ uptake capacity of this organelle. We also show here that PMR1 contributes ∼20% to the Ca²⁺ uptake capacity of dense-core secretory vesicles in β-cells (Fig. 4G), although the identity of the remaining Ca²⁺-ATPase activity (7) remains to be established. However, the limited effect of PMR1 depletion on organelle Ca²⁺ sequestration appears to be too small to impact on [Ca²⁺]_cyt in intact cells (Fig. 4B–C), where alternative mechanisms, including the plasma membrane Ca²⁺-ATPase and Na⁺/Ca²⁺ exchange, act in concert with Ca²⁺ sequestration via SERCA pumps to clear Ca²⁺ from the cytosol (44). Indeed, PMR1 depletion did not significantly affect organelle steady-state [Ca²⁺] (Fig. 4G–H), consistent with a relatively minor role for this pump in overall Ca²⁺ clearance.

PMR1 silencing in β-cells had a dramatic effect on the increases in [Ca²⁺]_cyt observed after the reintroduction of CaCl₂ to previously Ca²⁺-depleted cells, and this is likely to be caused, at least in part, by an activation of Ca²⁺ influx. Correspondingly, the enhanced [Ca²⁺] increases observed in PMR1-depleted cells were eliminated by the blockade of l-type Ca²⁺ channels with nimodipine. These observations are reminiscent of the effect of pmr1 deletion in S. cerevisiae (19). Previous findings in islets have suggested that depletion of thapsigargin-sensitive Ca²⁺ stores potentiates voltage-dependent Ca²⁺ influx, in part by stimulating a depolarizing current carried by store-operated Ca²⁺ channels (45). In addition, K⁺-induced Ca²⁺ influx was stimulated in SERCA3^−/− mouse islets (46). By contrast, in the present study we found that PMR1 depletion had no effect on K⁺-induced [Ca²⁺]_cyt increases, suggesting that indirect mechanisms, possibly involving changes in plasma membrane potential, are responsible for the effects of PMR1 depletion on Ca²⁺ influx via voltage-gated Ca²⁺ channels. In contrast to the effects of thapsigargin (45), it seems unlikely that increased Ca²⁺ influx via store-operated Ca²⁺ channels underlies the plasma membrane depolarization in the absence of PMR1. Thus, no differences were observed between control and PMR1-deficient cells in terms of the [Ca²⁺]_cyt increases provoked by the readdition of CaCl₂ in the presence of nimodipine (Fig. 4B). Instead, activation of a store-dependent current, possibly the depolarizing nonselective cation current primarily carried by Na⁺ that was previously described by Dukes and colleagues (47–49), might explain the enhanced influx of Ca²⁺ through voltage-gated channels after depletion of PMR1. Detailed electrophysiological studies will be required both to characterize the effect(s) of PMR1 ablation on these and other polarizing or depolarizing currents and to explore in detail the mechanisms by which depletion of a PMR1-dependent Ca²⁺ store may affect these currents.

Suppression of PMR1 caused a significant enhancement of glucose-stimulated, but not basal, insulin secretion (Fig. 5A). This result is similar to the effects of thapsigargin, which augments glucose-induced, but not basal, insulin secretion from isolated islets (50). The effects of PMR1 suppression seen here were not associated with an increase in average [Ca²⁺]_cyt but a modification of [Ca²⁺] oscillation shape. Thus, PMR1 depletion transformed rapid transients into broader [Ca²⁺] increases, on which rapid nonbaseline increases were superimposed (Fig. 5B). The fact that [Ca²⁺]_cyt remains elevated for longer periods in PMR1-depleted cells during the slow oscillations may underlie the enhanced secretion, although other mechanisms may also be involved. Importantly, if the principal role of PMR1 were to mediate Ca²⁺ sequestration from the cytosol, then after PMR1 depletion one would expect to observe an increase in oscillation amplitude caused by decreased Ca²⁺ buffering, a response similar to that observed in SERCA3^−/− mouse islets (46). In contrast, PMR1 depletion had no effect on oscillation amplitude, but instead it prolonged oscillation duration, supporting our proposal (see above) that the primary role of PMR1 is not to sequester cytosolic Ca²⁺ per se. Instead, the most important roles of this Ca²⁺-ATPase appear to contribute to Ca²⁺ uptake by secretory vesicles and to modulate, albeit by an indirect mechanism, the activity of plasma membrane l-type Ca²⁺ channels.

This study was supported by a Biotechnology and Biological Sciences Research Council Studentship (to K.J.M.) and grants to G.A.R from the Wellcome Trust (Program Grant 067081/Z/02/Z), the Human Frontiers Science Program, the Medical Research Council (U.K.), and Diabetes U.K. G.A.R. is a Wellcome Trust Research Leave Fellow.

We thank Rebecca Rowe for the preparation of pancreatic islets and Professor Anant Parekh (University of Oxford) for useful discussion.