The role of dense core secretory vesicles in the control of cytosolic-free Ca2+ concentrations ([Ca2+]c) in neuronal and neuroendocrine cells is enigmatic. By constructing a vesicle-associated membrane protein 2–synaptobrevin.aequorin chimera, we show that in clonal pancreatic islet β-cells: (a) increases in [Ca2+]c cause a prompt increase in intravesicular-free Ca2+ concentration ([Ca2+]SV), which is mediated by a P-type Ca2+-ATPase distinct from the sarco(endo) plasmic reticulum Ca2+-ATPase, but which may be related to the PMR1/ATP2C1 family of Ca2+ pumps; (b) steady state Ca2+ concentrations are 3–5-fold lower in secretory vesicles than in the endoplasmic reticulum (ER) or Golgi apparatus, suggesting the existence of tightly bound and more rapidly exchanging pools of Ca2+; (c) inositol (1,4,5) trisphosphate has no impact on [Ca2+]SV in intact or permeabilized cells; and (d) ryanodine receptor (RyR) activation with caffeine or 4-chloro-3-ethylphenol in intact cells, or cyclic ADPribose in permeabilized cells, causes a dramatic fall in [Ca2+]SV. Thus, secretory vesicles represent a dynamic Ca2+ store in neuroendocrine cells, whose characteristics are in part distinct from the ER/Golgi apparatus. The presence of RyRs on secretory vesicles suggests that local Ca2+-induced Ca2+ release from vesicles docked at the plasma membrane could participate in triggering exocytosis.

Introduction

In most mammalian cells, the ER (Streb et al., 1984; Rizzuto et al., 1993; Brini et al., 1993) and Golgi complex (Pinton et al., 1998) are believed to represent the major mobilizable intracellular Ca2+ stores (Rutter et al., 1998). Uptake of Ca2+ into these stores is mediated largely by sarco(endo)plasmic reticulum Ca2+–ATPases (SERCAs)* (Moller et al., 1996) and helps to maintain resting cytosolic Ca2+ concentrations ([Ca2+]c) at levels (∼10−7 M) some four orders of magnitude lower than in the extracellular space (∼10−3 M). Release of Ca2+ from the ER and Golgi complex is provoked by hormones and other agonists which generate inositol 1,4,5 trisphosphate (IP3) (Berridge, 1993) to open intracellular receptors (Mikoshiba, 1997). In certain cell types, such as skeletal muscle, these stores are also accessed by “Ca2+-induced Ca2+ release”, via receptors for the insecticide, ryanodine (Lai et al., 1988; Takeshima et al., 1989).

Secretory vesicles of both the dense core (Hellman et al., 1976; Hutton et al., 1983) and small synaptic types (Andrews and Reese, 1986) possess a substantial proportion of the total Ca2+ in cells specialized for peptide secretion. However, despite many studies, the role of secretory vesicles as Ca2+ buffers or stores remains controversial (Pozzan et al., 1994; Rutter et al., 1998; Yoo, 2000).

Mobilization of dense core vesicle Ca2+ by IP3 was originally suggested as a possibility in chromaffin cells (Yoo and Albanesi, 1990). Later data suggested that IP3 receptors may be present on the vesicle membrane in insulin-secreting cells (Blondel et al., 1994), though subsequent experiments showed the antibody used cross-reacted with insulin (Ravazzola et al., 1996). IP3, as well as the receptor (RyR) agonist cyclic ADP ribose (cADPr) (Galione, 1994) have been reported to release Ca2+ from individual acinar cell zymogen granules (Gerasimenko et al., 1996), although contamination of these preparations (e.g., with ER or Golgi apparatus–derived vesicles) is a potential problem (Yule et al., 1997). Finally, evidence for the direct participation of secretory granules in the control of cytoplasmic Ca2+ concentration was recently provided in intact goblet cells (Nguyen et al., 1998).

At present, the molecular mechanisms responsible for Ca2+ uptake into secretory vesicles are also a matter of controversy. At high Ca2+ concentrations (>50 μM), uptake into dense core vesicles in permeabilized hypophysial cells (Troadec et al., 1998; Thirion et al., 1999) and into isolated chromaffin cell granules (Krieger-Brauer and Gratzl, 1982) can occur via Na+/Ca2+ exchange, while synaptic vesicles also accumulate Ca2+ via a Ca2+-H+ antiport system (Goncalves et al., 1998). However, the pump/exchange mechanisms active at physiological [Ca2+]c are uncertain.

To examine the pathways by which Ca2+ crosses the limiting membrane of the dense core secretory vesicle of living islet β-cells, we have developed a new approach to monitor the free Ca2+ concentration within the secretory vesicle matrix ([Ca2+]SV) using recombinant targeted aequorin (Rizzuto et al., 1995). Cloned from the jellyfish Aequorea victoria (Inouye et al., 1985), aequorin is a calcium-sensitive bioluminescent protein (Cobbold and Rink, 1987), previously used to measure free Ca2+ concentrations in a variety of subcellular organelles (Rutter et al., 1998). Importantly, aequorin activity is less severely inhibited at low pH values (<6.5; Blinks, 1989) than Ca2+ probes based on green fluorescent protein (GFP) (Miyawaki et al., 1997; Baird et al., 1999; Emmanouilidou et al., 1999). If appropriately targeted, this probe should allow Ca2+ concentrations to be measured in the acidic environment of the secretory granule interior (Orci et al., 1985).

Vesicle-associated membrane protein (VAMP)2/synaptobrevin (Sudhof et al., 1989) is a vesicle-specific SNARE with a single transmembrane-spanning region. Expression of chimaeric cDNA encoding a fusion protein between VAMP2 and aequorin (VAMP.Aq) has therefore allowed the intravesicular-free Ca2+ concentration to be monitored dynamically in live MIN6 β-cells. With this approach, we show that Ca2+ is actively pumped into dense core vesicles when [Ca2+]c increases, and may be released via RyR, but not IP3, receptors. This release may be important at sites of high intracellular Ca2+, including sites of exocytosis at the plasma membrane.

Results

Subcellular targeting of recombinant VAMP.aequorin

Chimeric cDNA encoding hemagglutinin (HA)1-tagged aequorin, fused to VAMP2 (Sudhof et al., 1989), was generated as shown in Fig. 1 A.

Immunocytochemical analysis of MIN6 cells transfected with VAMP.Aq cDNA revealed close overlap with insulin staining (Fig. 1 B). Explored at a higher resolution by immunoelectron microscopy (Fig. 1 C), VAMP.Aq immunoreactivity was highly enriched in 61 of 148 (41.2%; n = 11 cells) vesicles colabeled for insulin (Fig. 1 C). Analyzed by single labeling for VAMP.Aq, staining of the ER, Golgi apparatus, and small synaptic-like microvesicles (Reetz et al., 1991) was very low, while reactivity was also present on the plasma membrane and in endosomes (see the legend to Fig. 1 and Discussion).

Reconstitution and calibration of secretory vesicle and ER-targeted aequorins

Given the high total Ca2+ content of secretory vesicles (Hutton et al., 1983), we used the approach adopted previously to measure Ca2+ in the ER lumen (Montero et al., 1995). Apoaequorin was reconstituted at a low free Ca2+ concentration (Montero et al., 1995), achieved by depleting cells of Ca2+ (Materials and methods). Depletion of vesicle Ca2+ had no marked effect on glucose or K+-stimulated insulin secretion, or on vesicle motility (Pouli et al., 1998b; Tsuboi et al., 2000; unpublished data).

To determine the response of the expressed aequorins to Ca2+ in situ, permeabilized cells were incubated at buffered Ca2+ concentrations in the presence of ionomycin and monensin (Fig. 2, C and E). The sensitivity to Ca2+ (at pH 7.0) of the VAMP.Aq chimaera was similar to that reported previously for mutant (D119A) aequorin (Montero et al., 1995). Intravesicular pH in intact cells was determined using a fusion construct between VAMP2 and a mutated, pH-sensitive GFP (pH.fluorin(e); Miesenbock et al., 1998), and gave a pH value of 6.3 ± 0.02 (n = 85 cells; Fig. 2 A). Confirming that this low intravesicular pH was unlikely to significantly affect VAMP.Aq, near identical calibration data were obtained at pH 5.7 (unpublished data). Therefore, we used the constants obtained in vitro (Montero et al., 1995) to calculate [Ca2+] from the fractional rate of aequorin consumption (F) according to: [Ca2+] = 1.44√10(LOG[F]3.4) (Rutter et al., 1993).

Dynamic measurement of secretory vesicle and ER Ca2+ concentrations

To monitor uptake of Ca2+ into vesicles in living cells, we provoked rapid Ca2+ influx into cells through store-operated channels (Parekh and Penner, 1997). During reintroduction of CaCl2 to cells previously perifused in EGTA, [Ca2+]c approached 500 nM, then fell back to 150–200 nM (Fig. 2 B). By contrast, steady state free [Ca2+] in the secretory vesicle matrix was 30–90 μM (51; 7.5 μM; n = 5 separate cultures; Fig. 2 D), whereas free [Ca2+] in the ER lumen was fivefold higher (249; 12.9 μM; n = 3 preparations; Fig. 2 F).

Pathways of Ca2+ uptake into dense core secretory vesicles

Both the rate and extent of the [Ca2+] increases in the secretory vesicles and the ER were strongly dependent on the presence of added ATP in permeabilized cells (Fig. 3, A and D). The increase in [Ca2+]SV upon reintroduction of Ca2+ ions was completely inhibited by the P-type Ca2+ pump inhibitor, orthovanadatate, at >100 μM (Fig. 3 B), but was insensitive to 10 μM orthovanadatate, a concentration which strongly inhibits the plasma membrane Ca2+-ATPase (Carafoli, 1991). Preincubation with the specific SERCA inhibitor, thapsigargin (Thastrup, 1990), and perifusion with cyclopiazonic acid (CPA) (Mason et al., 1991) had no effect on the changes in [Ca2+]SV (Fig. 3 C and Table I), but markedly (>85%) inhibited ER [Ca2+] increases (Fig. 3 E).

Arguing against vesicular Ca2+ uptake via Ca2+/H+ exchange, monensin, an Na+/H+ exchanger, had no effect on [Ca2+]SV increases in permeabilized cells (Fig. 4 A). Similarly, ionomycin (a Ca2+/H+ exchanger) decreased [Ca2+]SV marginally (∼20%; Fig. 4 B), which is consistent with the inability of this ionophore to bind to and transport Ca2+ against a prevailing H+ gradient, but caused a rapid and complete decline in [Ca2+]SV in the additional presence of monensin (Fig. 4 B). Neither the vacuolar H+-ATPase inhibitor, bafilomycin (Fig. 4, C and D), nor the protonophore carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (FCCP), had any effect on vesicular Ca2+ uptake (Fig. 4 E).

[Ca2+]SV increases were also completely unaffected by raising [Na+] from 0, 2, or 140 mM (Fig. 4 F), and by simultaneous blockade of Na+/Ca2+ exchange and Ca2+/H+ exchange (zero Na+, 300 nM bafilomycin; unpublished data).

Effects of IP3 on [Ca2+]SV and [Ca2+]ER

To achieve large changes in intracellular IP3 levels, we coexpressed the metabotropic glutamate receptor, mGluR5, with VAMP.Aq or ER.Aq. Stimulation of mGluR5-transfected cells with the specific agonist, (S)-3,5-dihydroxyphenylglycine (DHPG) (Thomas et al., 2000) caused a significant decrease in [Ca2+]ER (Fig. 5 E), but an increase in [Ca2+]SV (Fig. 5 A). DHPG had no significant effect on the distribution of vesicles (imaged after expression of VAMP.GFP/pHfluorin (r); Fig. 5 D), indicating that enhanced exocytosis and exposure of the VAMP.Aq to the extracellular medium was unlikely to contribute to the observed increases in [Ca2+]SV (Fig. 5 A, and see Discussion).

Similarly, in permeabilized cells, IP3 had no effect on [Ca2+]SV (Fig. 5, B and C), while causing a large decrease in [Ca2+]ER (Fig. 5, F and G).

RyR activation decreases [Ca2+]SV and [Ca2+]ER

The RyR agonists caffeine or 4-chloro-3-ethylphenol (4-CEP) (Zorzato et al., 1993) provoked rapid decreases in [Ca2+]SV and [Ca2+]ER (Fig. 6, A and C) in intact cells. Similarly, cADPr (Galione, 1994) caused clear decreases in both parameters in permeabilized cells (Fig. 6, B and D), and the effects of cADPr were strongly potentiated by palmitoyl CoA (Fig. 6, B and D), a coagonist of RyRs (Chini and Dousa, 1996).

4-CEP mobilizes Ca2+ from an acidic, CPA-insensitive store in fura-2–loaded MIN6 cells

The above results suggested that activation of RyR on vesicles should cause an increase in cytoplasmic [Ca2+]c, even after the depletion of the ER Ca2+ pool. Added to fura-2–loaded MIN6 cells, 4-CEP caused a substantial increase in [Ca2+]c in the absence of external Ca2+ (Fig. 7 A). This [Ca2+]c increase was partly retained after depletion of SERCA-dependent stores, giving a peak [Ca2+]c increase of 30–40% of that in control cells (Fig. 7 B versus A). The [Ca2+]c increase elicited by 4-CEP was also partially retained after treatment of cells with ionomycin (Fig. 7 C), but completely abolished after treatment with ionomycin plus monensin, to deplete acidic Ca2+ stores (Fig. 7 D). Demonstrating that the effects of 4-CEP on [Ca2+]SV were likely mediated by RyR, immunoreactivity to these channels was revealed on vesicle membranes by direct immunoelectron microscopy (Fig. 7 E).

Effects of nutrient secretagogues on [Ca2+]ER and [Ca2+]SV

Exposure of islet β- or MIN6 cells to glucose or other nutrients causes an increase in intracellular-free [ATP] (Kennedy et al., 1999) closure of ATP-sensitive K+ (KATP) channels (Bryan and AguilarBryan, 1997) and Ca2+ entry via voltage-sensitive (L-type) Ca2+ channels (Safayhi et al., 1997).

High (20 mM) glucose, or a combination of nutrient secretagogues (Ashcroft and Ashcroft, 1992), caused a large increase in steady state [Ca2+] both in the ER and in vesicles (Fig. 8, A and C), with no significant change in vesicular distribution (Fig. 8 B), which is consistent with the increases in [Ca2+]c seen under these conditions (Grapengiesser et al., 1988).

Discussion

Aequorin as a reporter of secretory vesicle–free Ca2+ concentration

We show here that aequorin can be targeted to the lumen of dense core secretory vesicles in living cells, with no incorporation into the ER, Golgi, or trans-Golgi network using the cell's own protein-sorting machinery (Fig. 1). However, a proportion of VAMP.Aq was also present on the plasma membrane and endosomes (Fig. 1 C). Since our recordings were made at either high (1.5 mM, intact cells) or low (400 nM, permeabilized cells) external Ca2+ concentrations, the plasma membrane–located photoprotein should be either rapidly exhausted, or inactive, respectively. In any case, it would appear that plasma membrane–targeted aequorin is not reconstituted, since we would have expected to see a “spike” of luminescence upon readdition of high Ca2+ concentrations to intact cells (e.g., Fig. 3 C). None was seen, suggesting that the mistargeted protein may be inactivated by unknown mechanisms during recycling between the plasma membrane and endosomes.

Similarly, endosomal aequorin is likely to be inactive under our conditions, since (a) the total endosomal Ca2+ content is low in neuroendocrine cells (Pezzati et al., 1997) and (b) free [Ca2+] is particularly low (∼3.0 μM) in acidic endosomes (Gerasimenko et al., 1998); active VAMP.Aq was located exclusively in an acidic compartment (Fig. 4 B). Simulation of the contribution of endosomal VAMP.Aq reveals that our measurements may slightly underestimate [Ca2+]SV (by <10%; unpublished data).

Mutated aequorins respond well to Ca2+ concentrations over the range (30–90 μM) which pertains within the secretory vesicle, as well as the rest of the secretory pathway (200–300 μM) (Montero et al., 1997; Pinton et al., 1998), without significant interference from the low pH environment. This feature of aequorin contrasts with GFP-based Ca2+ probes (Miyawaki et al., 1997; Baird et al., 1999; Emmanouilidou et al., 1999), whose fluorescence is strongly reduced at acidic pHs (Miesenbock et al., 1998; Fig. 2 E). Thus, aequorin may represent the most suitable molecularly targetable Ca2+ reporter presently available for the secretory vesicle interior.

Free Ca2+ concentration in the secretory vesicle lumen

Despite possessing a larger total Ca2+ content (Andersson et al., 1982; Hutton et al., 1983; Nicaise et al., 1992), secretory vesicles displayed a significantly lower free [Ca2+] than the ER or Golgi apparatus. Our measured values for [Ca2+]SV (∼50 μM) correspond fairly well to measurements using other techniques in isolated chromaffin granules (Krieger-Brauer and Gratzl, 1982; 24 μM; null point titration), respiratory tract goblet cells (24 μM; calcium orange 5 N fluorescence; Nguyen et al., 1998) and platelet α-granules (12 μM; null point titration; Grinstein et al., 1983). Thus, free Ca2+ represents <0.05% of the total vesicular calcium content of β-cell secretory vesicles (assuming a total Ca2+ concentration of 50–100 mM; Hutton et al., 1983). Secretory vesicles appear to have the highest Ca2+-buffering capacity of all subcellular organelles so far examined, with the percentage of free Ca2+ being much higher both in the cytosol (∼2%) and in the ER (∼10%; Pozzan et al., 1994). Importantly, resting [Ca2+]SV was well below the KM for Ca2+ of proinsulin-processing enzymes (Davidson et al., 1988).

The identity of the Ca2+ binding proteins (or other molecules) responsible for chelating free Ca2+ in these vesicles is unknown. Chromogranins (Yoo and Albanesi, 1990), or the mammalian homologue of Tetrahymena thermophila granule lattice protein 1 (Grlp1) (Chilcoat et al., 1996), are each strong possibilities. In addition, Ca2+ chelation by small molecules, such as ATP (Hutton et al., 1983), may also be involved. Finally, it is likely that in islet β-cells the insulin crystal itself also participates in chelating vesicular Ca2+ (Palmieri et al., 1988; see below).

Uptake of Ca2+ into secretory vesicles

The dense core secretory vesicle pool of islet β-cells has previously been considered relatively inert (Howell et al., 1975; Prentki et al., 1984). We provide evidence here that net uptake of Ca2+ into the dense core secretory vesicle population occurs during activated Ca2+ influx (Figs. 2 and 3), and in response to a receptor agonist (Fig. 5) or to nutrients (Fig. 8). However, and as discussed below (see also Fig. 9), these measurements do not exclude the possibility that discrete vesicle pools may experience different [Ca2+]SV changes.

We considered the possibility that the increases in [Ca2+]SV observed upon challenge of cells with agonists (Fig. 5) or nutrients (Fig. 8) may be due in part to the activation of exocytosis, and thus the exposure of aequorin within the vesicle matrix to the extracellular Ca2+ concentrations (1.5 mM). However, this phenomenon is likely to contribute negligibly to the observed signals, since only a tiny fraction of the total vesicle population (the “primed” pool) in β-cells (5–20/13,000 per min; Rorsman, 1997) undergoes fusion, even during the maximal stimulation of exocytosis. Consistent with this, we could detect no significant net movement of VAMP.GFP fluorescence to the plasma membrane after stimulation with DHPG (Fig. 5 D) or nutrients (Fig. 8 B).

Changes in intravesicular Ca2+ concentration detected with vesicle-targeted aequorin seem likely to result largely from the flux of Ca2+ ions across the vesicle membrane. However, two caveats are important: (a) Ca2+ release from/association with binding sites (or a “polyanionic matrix”) (Nguyen et al., 1998) within the vesicle may also occur, especially if the intravesicular concentrations of other ions (e.g., K+) change; (b) the number of Ca2+ ions that are transported is unknown, since this depends on the binding to intraorganellar sites. As discussed above, the present studies demonstrate that the ratio of free to bound Ca2+ within the secretory vesicles is much lower than that in the ER or Golgi lumen. This would appear at first to suggest that changes in intravesicular Ca2+ concentration must involve the flux of a very large number of Ca2+ ions. However, an alternative possibility is that there are two (or more) pools of bound Ca2+ in the secretory vesicle, one (the larger) comprising Ca2+ which is tightly bound (perhaps to a structural component such as the insulin crystal) and the other a more loosely bound and readily exchangeable pool. Depletion of this latter, “labile” pool, may thus lead to a substantial decrease in the free vesicular [Ca2+], while involving the release into the cytosol of relatively few Ca2+ ions.

The present data suggest that Ca2+ accumulation into vesicles is catalyzed at physiological [Ca2+]c chiefly by a P-type Ca2+-ATPase. Previous studies have indicated that transport of Ca2+ into the Golgi apparatus is mediated partly by a SERCA pump, and partly by another, unidentified thapsigargin-insensitive system (Pinton et al., 1998). This second ATP-dependent Ca2+ uptake system may be closely related to ATP2C1 (Hu et al., 2000), the mammalian homologue of the yeast Golgi Ca2+ transport ATPase, PMR1 (Sorin et al., 1997). Supporting this view, mRNA-encoding ATP2C1 is abundant in MIN6 cells, and polyclonal antibodies raised to ATP2C1 reveal a punctate pattern of intracellular staining (unpublished data). Intriguingly, although patients defective in the ATP2C1 gene develop skin lesions (“Hailey-Hailey” disease; Hu et al., 2000), it is unclear whether these individuals also tend to suffer from neuroendocrine or other disorders (e.g., diabetes mellitus).

Secretory vesicles are an IP3-insensitive, but caffeine/cADPr-sensitive Ca2+ store

The present results suggest that IP3 is unlikely to stimulate the release of Ca2+ from dense core secretory vesicles directly (Fig. 5).

By contrast, we now show (Figs. 6 and 7) that functional RyR channels are present on the dense core vesicle membrane of MIN6 cells. Type II RyR mRNA is present in ob/ob mouse islets, mouse β-TC3 cells (Islam et al., 1998), and in rat islets (Holz et al., 1999). RyR II protein has been detected in INS1 β-cells (Gamberucci et al., 1999), and ryanodine binding sites revealed in human islets (Holz et al., 1999), mouse islets and MIN6 cells (Varadi and Rutter, 2001). Evidence for functional RyRs on the ER INS-1 β-cells (Maechler et al., 1999) has also recently been provided. The present data indicate that dense core vesicles may represent a large fraction (∼30%) of the total RyR–accessible Ca2+ pool in MIN6 β-cells (Fig. 7 B). However, the proportion of the total vesicular Ca2+ pool which is mobilizable via RyR is small. Thus, the [Ca2+]c increases observed when the total acidic Ca2+ pool was emptied with CPA, ionomycin, and monensin (Fig. 7 D) were much larger than those apparent when cells were treated with 4-CEP after CPA and ionomycin (Fig. 7 C).

Role of cADPr in β-cells

In the present studies, cADPr caused an apparent release of Ca2+ from both the ER and from the secretory vesicles in permeabilized MIN6 β-cells. Moreover, we have recently observed (Varadi and Rutter, 2001) that photolysis of caged cADPr increases [Ca2+]c in MIN6 cells. These findings were surprising in light of earlier results using ob/ob mouse islets (Islam et al., 1993) or INS-1 β-cells (Rutter et al., 1994), where cADPr was ineffective in permeabilized cells. However, it should be noted that the present studies may provide greater sensitivity to small Ca2+ fluxes by measuring changes in [Ca2+] inside the secretory vesicle or ER.

Could changes in intracellular cADPr concentration play a role in the regulation of insulin secretion as proposed by Takasawa et al. (1993)? Supporting this view, mice homozygous for inactivation of the CD38 (ADPribose cyclase) gene display impaired glucose tolerance (Kato et al., 1999). However, no change in the islet cADPr content was observed by others in response to acutely elevated glucose concentrations in vitro (Scruel et al., 1998), suggesting further studies are necessary.

Conclusions

Dense core secretory vesicles rapidly take up Ca2+ ions via an ATP-dependent non-SERCA Ca2+ pump (see Scheme, Fig. 9). Ca2+ accumulated into vesicles may later be released via RyRs once these vesicles reach a high local Ca2+ concentration beneath the plasma membrane (Neher, 1998) or elsewhere in the cell. This release may further contribute to the high local [Ca2+]c in the vicinity of the submembrane vesicles (Emmanouilidou et al., 1999), and may be important to trigger exocytosis (Scheenen et al., 1998).

Materials And Methods

Materials

Cell culture reagents were obtained from GIBCO BRL or Sigma-Aldrich and molecular biologicals from Roche Diagnostics. Fura-2 and cADPr were from Sigma-Aldrich and IP3 was from Molecular Probes.

Construction of VAMP.Aq cDNA

VAMP2 cDNA was amplified by PCR using specific primers (forward: 5′-AAG CTT ACC ATG TCG GCT ACC GCT GCC ACC-3′; reverse: 5′-AAG CTT AGT GCT GAA GTA AAC GAT GAT-3′, HindIII sites are underlined). This fragment was then inserted upstream of the HindIII-EcoRI fragment encoding HA1-tagged D119-A aequorin (Kendall et al., 1992) in plasmid pBS+. Chimeric VAMP.Aq cDNA was shuttled as a NotI-XhoI fragment into pcDNAI (Invitrogen; Fig. 1 A).

Construction of adenoviral VAMP.Aq and ER.Aq

ER.Aq (Montero et al., 1995) and VAMP.Aq cDNAs were transferred into plasmid pShuttleCMV (He et al., 1998) as Kpn1-Xho1 and Xba1-HindIII fragments, respectively. The resultant constructs were recombined with vector pAdEasy-1, and viral particles were generated as described (Ainscow and Rutter, 2001).

Cell culture, transient transfection, and adenoviral infection

MIN6 β-cells (Miyazaki et al., 1990; passages #20 and #30) were cultured in DME, with additions as given previously (Ainscow and Rutter, 2001). Cells cultured on 13 mm poly-l-lysine–coated coverslips were transfected with VAMP.Aq or ER.Aq plasmid DNA (1 μg) using Lipofectamine (Promega) in Optimem I serum-free medium (GIBCO BRL), or infected with adenoviruses as described (Ainscow and Rutter, 2001). The total quantity of reconstituted aequorin was higher in cells transfected with ER.Aq than VAMP.Aq cDNAs due to the inclusion in the former of intronic regulatory elements. Thus, typical integrated counts/culture (Fig. 2) after the expression of VAMP.Aq were 3 × 105 and 1.5 × 106 for conventional and adenoviral expression, respectively, and 2 × 106 and 2 × 107 for ER.Aq.

[Ca2+] measurement with expressed aequorins

Cells were Ca2+ depleted by incubation with ionomycin (10 μM), the SERCA inhibitor CPA (10 μM; Mason et al., 1991; Pinton et al., 1998), and monensin (10 μM) in modified Krebs-Ringer bicarbonate buffer (KRB; 140 mM NaCl, 3.5 mM KCl, 0.5 mM NaH2PO4, 0.5 mM MgSO4, 3 mM glucose, 10 mM Hepes, 2 mM NaHCO3, pH 7.4) supplemented with 1 mM EGTA for 5 min at 4°C. Aequorin was reconstituted in 0.1 mM EGTA, 5 μM coelenterazine (cytosolic aequorin) or coelenterazine n (Shimomura et al., 1993; VAMP.Aq, ER.Aq) for 1–2 h at 4°C.

Intact cells were perifused (2 mL/min−1) with KRB plus additions as stated (37°C) close to a photomultiplier tube (ThornEMI), and emitted light was collected (one data point) with a photon-counting board (ThornEMI; Cobbold and Rink, 1987). Where indicated, cells were permeabilized with digitonin (20 μM, 1 min at 22°C) and subsequently perifused in intracellular buffer (IB) (140 mM KCl, 10 mM NaCl, 1 mM KH2PO4, 5.5 mM glucose, 2 mM MgSO4, 1 mM ATP, 2 mM Na+ succinate, 20 mM Hepes, pH 7.05) at 37°C.

[Ca2+] imaging with fura-2

Cells were loaded with 5 μM fura-2-AM (Sigma-Aldrich) for 40 min (37°C) in KRB plus 0.05% Pluronic F-127 (BASF). [Ca2+]c was monitored in single cells at 37°C as described (Ainscow et al., 2000), using a Leica DM-IRBE inverted microscope (40× objective), and a Hamamatsu C4742-995 charge-coupled device camera driven by OpenLab (Improvision) software.

Imaging acridine orange fluorescence

Permeabilized cells were loaded with 3 μM acridine orange (Tsuboi et al., 2000) before imaging (37°C) on a Leica DM-IRBETM inverted optics epiflurosecence microscope, at 493 ± 10 excitation, 530 nm emission (filters were from Chroma Techology).

Immunocytochemistry

48 h after transfection or infection, cells were fixed in 3.7% (vol/vol) paraformaldehyde and probed with antibodies essentially as described (Pouli et al., 1998a).

Immunoelectron microscopy

Gelatin-embedded, aldehyde-fixed cells were frozen in liquid nitrogen and ultrathin cryosections were obtained with a Reichert-Jung Ultracut E. Immunogold localization was performed as described (Confalonieri et al., 2000). Sections were immunostained either with rabbit anti-HA antibody (Sigma-Aldrich), or with a sheep anti-RyR antibody (Molecular Probes) followed by 10-nm protein A-gold. For double labeling, anti-HA–labeled sections were incubated with 1% glutaraldehyde in 0.1 M Na2PO4, pH 7.0, followed by incubation with a guinea pig antiinsulin antibody (Dako) and 15-nm protein A-gold. Sections were examined with a ZEISS EM 902 electron microscope. No labeling was detected in control sections (unpublished data).

Confocal imaging

Cells were imaged at 37°C on a Leica SP2 confocal spectrophotometer, (488 nm excitation) using a 100× oil immersion objective.

Other methods and statistics

Free [Ca2+] and [Mg2+] were calculated using “METLIG” (Rutter and Denton, 1988). Data are presented as the mean ± SEM for the number of observations given, and statistical significance calculated using Student's t test.

Acknowledgments

We thank Dr. Mark Jepson and Alan Leard (Bristol MRC Cell Imaging Facility) for their assistance with cell imaging, Dr. Nadia Todesco for help with the construction of VAMP.Aq cDNA, Maria Cristina Gagliani for help in electron microscopy, and Dr. Elek Molnar for the provision of mGluR5 cDNA and DHPG.

Supported by grants to G.A. Rutter from the Medical Research Council (UK), Human Frontiers Science Program, the Wellcome Trust, the Biotechnology and Biological Research Council, DiabetesUK, and the European Union; and to R. Rizzuto from Italian “Telethon” (Projects 850 and E0942), the Italian University and Health Ministries, the Italian Space Agency, The Italian National Research Council, The Armenise Harvard Foundation, and the Italian Association for Cancer Research.

*

Abbreviations used in this paper: cADPr, cyclic ADP ribose; CPA, cyclopiazonic acid; DHPG, dihydroxyphenylglycine; GFP, green fluorescent protein; HA, hemagglutinin; IB, intracellular buffer; IP3, inositol 1,4,5 trisphosphate; KRB, Krebs-Ringer bicarbonate buffer; RyR, ryanodine receptor; 4-CEP, 4-chloro-3-ethylphenol; SERCA, sarco(endo)plasmic reticulum Ca2+–ATPase; VAMP, vesicle-associated membrane protein.

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