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Diabetes 50:1012-1020, 2001
© 2001 by the American Diabetes Association, Inc.

Ca2+-Dependent Exocytosis of L-Glutamate by {alpha}TC6, Clonal Mouse Pancreatic {alpha}-Cells

Hiroshi Yamada1, Masato Otsuka1, Mitsuko Hayashi1, Shuuichi Nakatsuka1, Kazuyuki Hamaguchi2, Akitsugu Yamamoto3,4, and Yoshinori Moriyama1,4

1 Department of Biochemistry, Faculty of Pharmaceutical Sciences, Okayama University, Okayama
2 Faculty of Medicine, Oita Medical University, Oita
3 Department of Physiology, Kansai Medical University, Moriguchi
4 CREST, Japan Science and Technology Corporation, Osaka, Japan


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Pancreatic islet cells express receptors and transporters for L-glutamate and are thus believed to use L-glutamate as an intercellular signaling molecule. However, the mechanism by which L-glutamate appears in the islets is unknown. In the present study, we investigated whether L-glutamate is secreted through exocytosis by {alpha}TC6 cells (clonal mouse pancreatic {alpha}-cells). An appreciable amount of L-glutamate was released from cultured cells after the addition of KCl or A23187 in the presence of Ca2+ and 10 mmol/l glucose in the medium. The KCl-induced glutamate release was significantly reduced when assayed in the absence of Ca2+ or when the cells were pretreated with EGTA-AM. The KCl-induced Ca2+-dependent glutamate release was inhibited ~40% by voltage-gated Ca2+ channel blockers, such as nifedipine at 20 µmol/l. The degree of KCl-induced Ca2+-dependent glutamate release was correlated with an increase in intracellular [Ca2+], as monitored by fura-2 fluorescence. Botulinum neurotoxin type E inhibited 55% of the KCl-induced Ca2+-dependent glutamate release, followed by specific cleavage of 25 kDa synaptosomal-associated protein. Furthermore, bafilomycin A1, a specific inhibitor of vacuolar H+-ATPase, inhibited 40% of the KCl-induced Ca2+-dependent glutamate release. Immunoelectronmicroscopy with antibodies against synaptophysin, a marker for neuronal synaptic vesicles and endocrine synaptic-like microvesicles, revealed a large number of synaptophysin-positive clear vesicles in cells. Digitonin-permeabilized cells took up L-glutamate only in the presence of MgATP, which is sensitive to bafilomycin A1 or 3,5-di-tert-butyl-4-hydroxybenzylidene-malononitrile (a proton conductor) but insensitive to either oligomycin or vanadate. From these results, it was concluded that {alpha}TC6 cells accumulate L-glutamate in the synaptophysin-containing vesicles in an ATP-dependent manner and secrete it through a Ca2+-dependent exocytic mechanism. The Ca2+-dependent glutamate release was also triggered when cells were transferred in the medium containing 1 mmol/l glucose, suggesting that low glucose treatment stimulates the release of glutamate. Our results are consistent with the idea that L-glutamate is secreted by {alpha}-cells through Ca2+-dependent regulated exocytosis.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In the mammalian central nervous system, L-glutamate is the major excitatory neurotransmitter and plays important roles in many neuronal processes, such as fast synaptic transmission and neuronal plasticity (1,2). To use L-glutamate as an intercellular signaling molecule, neuronal cells develop glutamatergic systems comprising glutamate exocytosis (signal output), glutamate receptors (signal input), and glutamate reuptake systems (signal termination). Recent evidence has indicated that peripheral endocrine cells also develop glutamatergic systems: mammalian pinealocytes (endocrine cells for melatonin) use L-glutamate as an inhibitory chemical mediator of melatonin synthesis (3).

The Langerhans’ islet, a pancreatic endocrine miniature organ, is composed of four major types of endocrine cells, i.e., insulin-secreting ß-cells, glucagon-secreting {alpha}-cells, polypeptide-secreting cells, and somatostatin-secreting {delta}-cells. The Langerhans’ islet is another example of the presence of peripheral glutamatergic systems (4). A clonal ß-cell line, MIN6, expresses functional ionotropic glutamate receptors (5). Functional (RS)-{alpha}-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA)-type receptors have been identified in {alpha}- and ß-cells and the polypeptide-secreting cells of islets using reverse-transcriptase–polymerase chain reactions, immunohistochemistry, and electrophysiology (678). The kainate receptor has also been identified in islet {alpha}-cells (7,8). After stimulation of islets with AMPA or kainate, intracellular [Ca2+] increased by way of activation of voltage-gated Ca2+ channels (6,7), resulting in an elevated level of insulin secretion through increased exocytosis of insulin granules in ß-cells (9). In {alpha}-cells, AMPA treatment increased intracellular [Ca2+] and triggered exocytosis of glucagon-containing secretory granules, resulting in increased secretion of glucagon (10). Moreover, the Na+-dependent glutamate transporter sequesters extracellular glutamate in the {alpha}-cell–rich islet mantle (11).

Before a conclusion can be made regarding the functional operation of the glutamatergic system in the islets, there is an important issue remaining to be solved: How does L-glutamate appear in the extracellular space of the islets? At least two possibilities have been proposed. One is that L-glutamate is secreted by presumptive glutamatergic output neurons innervating the islets. However, no morphological evidence has been obtained as yet regarding this possibility. The other is that L-glutamate is secreted by neighboring islets cells, such as {alpha}-cells, through a paracrine-like chemical transduction mechanism (4). This possibility is more likely because it is known that {alpha}-cells possess phosphate-activated glutaminase (12), a peculiar enzyme for glutamate-releasing nerve endings in the central nervous system (13).

To elucidate the mechanism by which L-glutamate appears in the extracellular space of the islets, we examined whether a clonal {alpha}-cell line, {alpha}TC6, secretes L-glutamate through an exocytic mechanism. The use of clonal cells may exclude the possible participation of other islet cells, including ß-cells. It was found that {alpha}TC6 cells secrete L-glutamate in a Ca2+-dependent manner. Furthermore, we identified synaptophysin-positive vesicles as putative storage organelles for L-glutamate in {alpha}TC6 cells. The present results are fully consistent with the idea that {alpha}-cells store L-glutamate in synaptophysin-containing vesicles and secrete it through a paracrine-like exocytic mechanism.


   RESEARCH DESIGN AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESEARCH DESIGN AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell cultures.
{alpha}TC6 (14), HeLa, and COS cells were maintained in Dulbecco’s modified Eagle medium (DMEM) (Gibco) supplemented with 10% fetal calf serum, 55 µg/ml sodium pyruvate, 4.5 g/l glucose, 0.1 mg/l streptomycin, 100 U/ml penicillin G, and 0.25 mg/l fungizone and incubated at 37°C under 5% CO2. The {alpha}TC6 cells showed morphologic features similar to those in the original report (14) (Fig. 4), and like the cells in the original report, they did not produce detectable levels of insulin and contained 64 ng glucagon per 105 cells (14), suggesting that the cells maintained the original properties. MIN6 cells, a clonal ß-cell line (15), were maintained in DMEM supplemented with 15% fetal calf serum. PC 12 cells were cultured in DMEM containing 5% fetal calf serum, 5% horse serum, and 25 ng/ml nerve growth factor (16). The dispersed cells were washed three times with the above medium then placed in a 35-mm culture dish coated with poly-L-lysine (5.0 x 105 cells per dish) and cultured in the above medium at 37°C under 5% CO2. For experimental procedures, cells were maintained for 5 days, washed with culture medium, further cultured for 1 h, then used for experiments.



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  		FIG. 4. Immunoelectronmicroscopy revealing synaptophysin-containing vesicles. A: Gold particle for synaptophysin is associated with clear vesicles, as indicated by arrows. B: Synaptophysin-containing vesicles are abundant in the peripheral region. C: Gold particle for synaptophysin is also associated with the Golgi apparatus, as indicated by arrows. D: Gold particles are absent in dense-cored vesicles (arrows) but present in clear vesicles (arrowhead). CV, clear vesicles; DV, dense-cored vesicles; G, Golgi apparatus; L, lysosome; M, mitochondrion; N, nucleus; PL, plasma membrane. Bar: A and C, 1 µm; B and D, 200 nm.

 
Assay of glutamate release.
Cultured cells (2.5 x 106 cells per dish) were washed three times with Ringer’s solution comprising 128 mmol/l NaCl, 1.9 mmol/l KCl, 1.2 mmol/l KH2PO4, 2.4 mmol/l CaCl2, 1.3 mmol/l MgSO4, 26 mmol/l NaHCO3, 10 mmol/l glucose, and 10 mmol/l HEPES (pH 7.4), or they were washed with low-concentration Ca2+-Ringer’s solution comprising 128 mmol/l NaCl, 1.9 mmol/l KCl, 1.2 mmol/l KH2PO4, 0.2 mmol/l CaCl2, 1 mmol/l EGTA, 3.8 mmol/l MgSO4, 26 mmol/l NaHCO3, 10 mmol/l glucose, and 10 mmol/l HEPES (pH 7.4). After cells had been incubated in 2 ml of either of the above mediums at 37°C, the release of L-glutamate was stimulated by the addition of 50 mmol/l KCl (17), as described previously. When necessary, various antagonists for voltage-gated Ca2+ channels were included in the incubation medium. Aliquots (10 µl) were taken at time intervals, and the amount of extracellular glutamate was determined by high-performance liquid chromatography (HPLC) with precolumn O-phthalaldehyde derivatization, separation on a reverse-phase Resolve C18 column (3.9 x 150 mm) (Waters), and fluorescence detection (18).

Treatment with botulinum neurotoxin type E.
The intoxication of {alpha}-cells with botulinum neurotoxin type E (BoNT/E) was performed by a procedure similar to that previously described (19,20). {alpha}-Cells (2.5 x 106 cells per dish) were incubated at 37°C for 24 h in a low ionic strength buffer consisting of 5 mmol/l NaCl, 4.8 mmol/l KCl, 2.2 mmol/l CaCl2, 1.2 mmol/l MgSO4, 20 mmol/l HEPES-NaOH, 10 mmol/l glucose, 220 mmol/l sucrose, and 0.5% bovine serum albumin (pH 7.4) in the presence or absence of 10 or 50 nmol/l BoNT/E. Then, the cells were washed with fresh culture medium and incubated for an additional 12 h at 37°C. Finally, KCl-evoked glutamate release was measured as described above.

Measurement of intracellular [Ca2+].
For the analysis of intracellular [Ca2+], an Argus 20/CA ratio imaging system (Hamamatsu Photonics, Shizuoka, Japan) was used. Cells were cultured for 3 days on a thin glass coverslip precoated with poly-L-lysine (0.12 mm thick and 40 mm in diameter, 8.0 x 105 cells per coverslip). After exchanging the old medium for fresh culture medium, the cells were treated with 5 µmol/l Fura 2-acetoxymethylester (AM) (Dojindo, Kumamoto, Japan) for 50 min at 37°C and then washed twice with the same medium. The cells were perfused with the warmed Ringer’s solution or the low-concentration Ca2+-Ringer’s solution. Images were continuously taken at 37°C with a silicon-intensified camera (C2741–08, Hamamatsu Photonics). The velocity of data acquisition for F334 by F380 images was 4 s at a resolution of 256 x 256 pixels per image. A personal computer with appropriate software (U4469, Hamamatsu Photonics) was used to control the optical equipment, then to record and analyze the data. The software enabled subtraction of background fluorescence, pixel-to-pixel division of F334 by F380 images, fitting of the F334-to-F380 ratios to a [Ca2+] calibration curve prepared separately, and digital averaging of the Ca2+ concentration in multiple cells (21).

Immunoblotting.
{alpha}TC6 cells or a membrane fraction of rat brain prepared as described (20) was denatured with SDS sample buffer containing 1% SDS and 10% ß-mercaptoethanol and then electrophoresed on a 12% polyacrylamide gel in the presence of SDS (22). After electrotransfer at 0.3 amperes for 2 h, the nitrocellulose filters were blocked in a buffer consisting of 20 mmol/l Tris-Cl (pH 7.6), 5 mmol/l EDTA, 0.1 mol/l NaCl, and 0.5% bovine serum albumin for 4 h and then probed with 50-µg antibodies in the above buffer. The filters were washed with 20 mmol/l Tris-Cl buffer (pH 7.6) containing 5 mmol/l EDTA, 0.1 mol/l NaCl, and 0.1% Tween 20, treated with peroxidase-labeled anti-rabbit IgG or anti-mouse IgG at a dilution of 1:2000 for 30 min, washed again with the same buffer, and then subjected to enhanced chemiluminescence amplification according to the manufacturer’s manual (Amersham).

Immunoelectronmicroscopy.
The pre-embedding silver enhancement immunogold method described by Mandai et al. (23) was used with a slight modification (24,25). Cells on poly-L-lysine-coated plastic coverslips were fixed in 4% paraformaldehyde in 0.1 mol/l sodium phosphate buffer for 30 min. After the cells were washed three times for 5 min, they were cryo-protected in buffer containing 35% sucrose and 14% glycerol for 15 s, frozen in liquid nitrogen, and thawed at room temperature. The cells were then incubated in buffer containing 0.005% saponin, 10% bovine serum albumin, 10% normal goat serum, and 0.1% cold water fish skin gelatin (Sigma) for 30 min and reacted with anti-synaptophysin antibodies (10 µg/ml) in the above buffer overnight at 4°C. The cells were then washed five times in the same buffer containing 0.005% saponin and 1% bovine serum albumin and incubated with goat anti-rabbit IgG conjugated to colloidal gold (1.4 nm in diameter) (Nanogold; Nanoprobes) in the same buffer containing 0.005% saponin and 1% bovine serum albumin for 2 h. The cells were then washed five times with the buffer for 10 min and fixed with 1% glutaraldehyde for 10 min. After washing, the gold labeling was intensified using a silver enhancement kit (HQ silver; Nanoprobes) for 6 min at room temperature in the dark. After washing in distilled water, the cells were postfixed with 0.5% OsO4 for 90 min at 4°C, washed in distilled water, dehydrated with a graded series of ethanol, and embedded in epoxy resin. Ultra-thin sections were doubly stained with uranyl acetate and lead citrate and observed under a Hitachi H7000 electron microscope. Dr. M. Takahashi (Mitsubishi Life Science Institute, Tokyo) provided mAbBR05 (monoclonal antibodies against 25 kDa synaptosomal-associated protein [SNAP25]), and mAb171b5 (monoclonal antibodies against synaptophysin) (26).

Glutamate uptake by intracellular organelles in digitonin-permeabilized cells.
{alpha}TC6 cells were rinsed with 1 ml of the buffer comprising 20 mmol/l MOPS-Tris (pH 7.0), 0.3 mol/l sucrose, 2 mmol/l Mg-acetate, and 4 mmol/l KCl. The cells were then permeabilized for 10 min at 37°C in 0.5 ml of the buffer containing 10 µmol/l digitonin (27). The medium was then replaced with fresh buffer containing Tris-ATP at 2 mmol/l in the absence of digitonin. In some experiments, bafilomycin A1 at 1 µmol/l was also included in the medium. Then, glutamate uptake was immediately started by the addition of radioactive glutamate (2.5 µCi, 0.1 mmol/l) at 37°C, as described previously (28,29). After 10 min incubation, uptake was terminated by washing the cells twice with 1 ml of ice-cold 20 mmol/l MOPS-Tris (pH 7.0) containing 0.3 mol/l sucrose. Then, the cells were lysed with 1 ml of 1% SDS, and the radioactivity was counted with a liquid scintillation counter.

Other procedures.
Indirect immunofluorescence microscopy was performed as described previously (24). Content of glucagon and insulin were determined by enzyme-linked immunoassay according to the manufacturer’s manual (Amersham). Protein concentrations were determined with a BioRad Protein Assay Kit with bovine serum albumin as a standard.

Other chemicals.
The L-[2,3-3H]-glutamate (9.25 MBq) was obtained from NEN Life Science Products (Boston, MA). Digitonin was purchased from Wako Chemical (Osaka, Japan). BoNT/E was provided by Dr. S. Kozaki (Osaka Prefecture University). Other chemicals were of the highest grade commercially available.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESEARCH DESIGN AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Ca2+-dependent glutamate release.
As the first step of the study, we examined whether {alpha}TC6 cells release L-glutamate through an exocytic mechanism. In analogy to the glutamate exocytosis by neurons and pinealocytes (17,30,31), the glutamate concentration in the medium of cultured {alpha}TC6 cells was measured by HPLC after stimulation of the cells with KCl. An appreciable amount of L-glutamate (2.52 ± 0.4 nmol per 106 cells, which corresponds to 23% total free glutamate, 50 determinations) has been released by {alpha}TC6 cells at 10 min after the addition of KCl in the presence of Ca2+. The amount of released glutamate was dependent on time (Fig. 1A) and the concentration of KCl (Fig. 1B): the glutamate concentration was saturated at 10 min and with 50 mmol/l KCl, respectively. In the absence of CaCl2, little glutamate was released by the cells with the addition of KCl. When cells were treated with EGTA-AM to remove intracellular free Ca2+, the cells lost the ability to release L-glutamate in the presence of KCl (Fig. 1C). The Ca2+-dependent fraction of KCl-induced glutamate release is designated as the KCl-induced Ca2+-dependent glutamate release in this report. Similarly, A23187, a Ca2+ ionophore, caused rapid glutamate release by the cells only in the presence of Ca2+ (Fig. 1D). These results indicate that L-glutamate is released by cultured cells in a manner dependent on Ca2+. It is possible that entry of Ca2+ from the extracellular space is necessary for the glutamate release. No KCl-induced Ca2+-dependent glutamate release (<0.04 ± 0.01 nmol per 106 cells at 10 min after the addition of KCl) was observed when COS, HeLa, MIN6, and PC 12 cells were assayed.



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  		FIG. 1. Ca2+-dependent glutamate release by {alpha}TC6 cells. A: Time course of the KCl-induced glutamate release. The cells were incubated in the high-concentration Ca2+-Ringer’s solution (•) or low-concentration Ca2+-Ringer’s solution ({circ}) containing 10 mmol/l glucose, and then KCl at 50 mmol/l was applied. After the times indicated, the medium was carefully taken, and the glutamate concentration was determined. B: The KCl-dose dependence of the Ca2+-dependent glutamate release was measured at 10 min after the addition of the indicated concentrations of KCl. C: {alpha}TC6 cells (2.5 x 106 cells) were incubated in the absence (control) or presence of EGTA-AM at 50 µmol/l for 30 min and then washed with Ringer’s solution. Then, the KCl-induced Ca2+-dependent glutamate release was measured as previously described. D: The time course of A23187-evoked glutamate release was measured in the high-concentration Ca2+-Ringer’s solution (•) or low-concentration Ca2+-Ringer’s solution ({circ}). At time 0, A23187 at 5 µmol/l was added to the medium. After incubation for the times indicated, the glutamate concentration in the medium was measured. A23187 was dissolved in DMSO. The solvent itself does not affect the glutamate release at all. All the results in the figure are means ± SE (four independent experiments).

 
We then examined the effects of voltage-gated Ca2+ channel blockers on the KCl-induced Ca2+-dependent glutamate release because the requirement of Ca2+ suggests the participation of Ca2+ channel(s) in this process. Table 1 indicates that the cadmium ion, a nonspecific inhibitor of voltage-gated Ca2+ channels (32), inhibited the KCl-induced Ca2+-dependent glutamate release; the concentration for 50% inhibition was 100 µmol/l. Furthermore, it was found that at 20 µmol/l each, the L-type Ca2+ channel blockers nifedipine, nitrendipine, and diltiazem inhibited 40, 60, and 64% of the KCl-induced Ca2+-dependent glutamate release, respectively (Table 1). On the other hand, the KCl-induced Ca2+-dependent glutamate release was not affected by {omega}-conotoxin GIVA (an N-type Ca2+ channel blocker) (33), {omega}-conotoxin MVIIC (a P/Q-type Ca2+ channel blocker) (34), or {omega}-agatoxin IVA (a P-type Ca2+ channel blocker) (35,36) (Table 1). These results suggest that L-type Ca2+ channels are at least partly involved in the KCl-induced Ca2+-dependent glutamate release by {alpha}TC6 cells.


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  		TABLE 1 The effects of Ca2+-channel antagonists on the KCl-induced Ca2+-dependent glutamate release and intracellular [Ca2+] in {alpha}TC6 cells

 
Intracellular [Ca2+] change.
As mentioned above, Ca2+ entry through voltage-gated Ca2+ channels may facilitate the KCl-induced Ca2+-dependent glutamate release. To confirm that intracellular [Ca2+] actually increases after KCl treatment, intracellular [Ca2+] was measured in fura-2–loaded cells. Under the experimental conditions used, almost all cells were labeled with Fura-2-AM. The intracellular [Ca2+] level in the resting cells was 98 ± 12 nmol/l (300 determinations) (Table 1). The addition of KCl at 50 mmol/l increased the intracellular [Ca2+] level to 548 ± 73 nmol/l in the presence of Ca2+ in the medium. In contrast, essentially no increase in intracellular [Ca2+] was observed in the absence of Ca2+ (Table 1) or when the cells were pretreated with EGTA-AM at 50 µmol/l (104 ± 9 nmol/l, 62 determinations). Furthermore, nifedipine, nitrendipine, and diltiazem inhibited 71, 86, and 90% of the KCl-evoked increase in intracellular [Ca2+], respectively (Table 1). {omega}-Conotoxin GIVA, {omega}-conotoxin MVIIC, and {omega}-agatoxin IVA (antagonists for N- or P/Q-type channels) did not affect the KCl-evoked [Ca2+] increase in these cells (Table 1). Thus, degrees of the KCl- and Ca2+-dependent intracellular [Ca2+] change and glutamate release were well correlated to each other. Taking all the results together, it was concluded that the KCl-induced Ca2+-dependent glutamate release is triggered by the entry of Ca2+ through voltage-gated Ca2+ channels in cultured {alpha}TC6 cells.

Evidence for exocytosis of glutamate.
We further characterized KCl-induced Ca2+-dependent glutamate release by {alpha}TC6 cells with regard to the effects of temperature. As shown in Fig. 2, KCl-induced Ca2+-dependent glutamate release was affected by the temperature: change was not observed at 4°C, but an increase in glutamate release appeared gradually with increasing temperature and was maximum at 37°C. Furthermore, once glutamate had been secreted, for a time the {alpha}TC6 cells lost the ability to release Ca2+-dependent glutamate in the presence of KCl. Although a first stimulation of glutamate release with KCl decreased the cell glutamate content by only 20%, a second stimulation applied within 1 h was ineffective. Its efficiency was gradually restored with incubation and had recovered completely after 12 h, suggesting that charged and discharged processes are involved in the KCl-induced Ca2+-dependent glutamate release. These properties are similar to those of exocytosis of glutamate by synaptic vesicles and synaptic-like microvesicles (17,30,31), supporting the idea that glutamate is secreted through exocytosis.



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  		FIG. 2. The effect of incubation temperature on KCl-induced Ca2+-dependent glutamate release. {alpha}TC6 cells (2.5 x 106 cells) were incubated at the indicated temperatures for 15 min, and then the KCl-induced Ca2+-dependent glutamate release by {alpha}TC6 cells was measured at 10 min. The results are presented as means ± SE (four independent experiments).

 
The sensitivity to BoNT/E constitutes evidence of Ca2+-dependent regulated exocytosis because this neurotoxin is a protease that is specific to SNAP25, splitting the SNAP25 and resulting in inhibition of exocytosis via secretory granules, synaptic vesicles, and synaptic-like microvesicles (17,19,373839). BoNT/E cleaved SNAP25, yielding a low-molecular fragment that migrated faster on a polyacrylamide gel during electrophoresis (Fig. 3A, asterisk). In contrast, BoNT/E did not affect vesicle-associated membrane protein 2, syntaxin-1, synaptotagmin, synaptophysin, N-ethylmaleimide-sensitive fusion protein, ß-SNAP, or V-ATPase subunit A or E (data not shown). Under the same assay conditions, BoNT/E inhibited the KCl-induced Ca2+-dependent glutamate release (Fig. 3B). The inhibitory potency of BoNT/E was essentially the same as that observed in the exocytosis of various neurotransmitters (17,19,373839). The BoNT/E treatment did not affect the KCl-induced [Ca2+] increase in these cells (550 ± 31 nmol/l, 120 determinations). These results indicated that SNAP25 is involved in the KCl-induced Ca2+-dependent release of glutamate by cultured cells.



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  		FIG. 3. BoNT/E-evoked cleavage of SNAP25 and inhibition of KCl- and Ca2+-dependent glutamate release. A: Cells were treated with BoNT/E at the indicated concentrations, washed with phosphate-buffered saline containing 0.01% DNase and 10 µg/ml each of leupeptin and pepstatin A, and then dissolved in the sample buffer containing 10% SDS and ß-mercaptoethanol. After the dissociation of proteins, samples were applied to a 12.5% polyacrylamide gel containing SDS then electrophoresed and analyzed by immunoblotting with antibodies against SNAP25. B: After the same BoNT/E treatment described above, the KCl-induced Ca2+-dependent release at 10 min was measured as described in RESEARCH DESIGN AND METHODS. The results are presented as means ± SE (four independent experiments); 100% corresponds to 2.48 ± 0.39 nmol per 106 cells.

 
It is known that bafilomycin A1, a specific inhibitor of V-ATPase in vivo and in vitro (404142), effectively inhibits the exocytosis of L-glutamate (43,44) because the compound dissipates an electrochemical proton gradient necessary for glutamate uptake into vesicles (45,46). As expected, bafilomycin A1 at 1 µmol/l inhibited 40% of the KCl-induced Ca2+-dependent glutamate release without affecting the KCl-evoked [Ca2+] increase in these cells (553 ± 21 nmol/l, 100 determinations). The results suggest that an electrochemical proton gradient for vesicular glutamate uptake is necessary at least in part for the KCl- and Ca2+-dependent glutamate release by {alpha}TC6 cells. From all the results shown above, it was concluded that glutamate is released by cultured {alpha}TC6 cells through a soluble NSF attachment protein protein-dependent exocytic mechanism.

Mechanism of storage of L-glutamate.
Before exocytosis, L-glutamate must be accumulated in vesicular structures, such as synaptic vesicles or synaptic-like microvesicles (3,47,48). We attempted to identify the presumptive secretory vesicles for L-glutamate in {alpha}TC6 cells using immunoelectronmicroscopy with antibodies against synaptophysin because synaptophysin-containing vesicles are responsible for the storage of L-glutamate in neurons and pinealocytes (28,29,45,46). As shown in Fig. 4A, immunogold for synaptophysin is primarily associated with clear vesicles with diameters of 50–200 nm. The synaptophysin-positive vesicles are distributed throughout the cells and are especially abundant in peripheral regions (Fig. 4B). A lower amount of labeling was also observed in the region of the Golgi apparatus (Fig. 4C, arrows). Only the background level of labeling was observed in other areas, including the nucleus, mitochondria, lysosomes (Figs. 4A–C), and dense-cored vesicles (Fig. 4D). These results suggest that synaptophysin-containing vesicles are distinct from these organelles.

As in the case of glutamate uptake by synaptic vesicles and synaptic-like microvesicles (28,29,45,46), the glutamate transporter energetically coupled with V-ATPase may be responsible for the accumulation of L-glutamate in synaptophysin-positive vesicles. As shown in Fig. 5, radiolabeled L-glutamate was taken up by digitonin-permeabilized cells in a manner dependent on the presence of ATP. The omission of Mg2+ drastically reduced the ATP-dependent glutamate uptake. Consistent with glutamate exocytosis by {alpha}TC6 cells, bafilomycin A1 at 1 µmol/l inhibited ATP-dependent glutamate uptake (Fig. 5). Furthermore,3,5-di-tert-butyl-4-hydroxybenzylidenemalononitrile(SF6847), a proton conductor that dissipates an electrochemical proton gradient, also inhibited the ATP-dependent glutamate uptake, whereas neither oligomycin at 5 µmol/l (an inhibitor for mitochondrial ATPase) nor vanadate at 1 mmol/l (an inhibitor for P-type ATPase) affected the uptake (Fig. 5). These results constitute evidence that the glutamate transporter energetically coupled with V-ATPase is responsible for the storage of L-glutamate in vesicles in {alpha}TC6 cells.



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  		FIG. 5. ATP-dependent uptake of radiolabeled L-glutamate by digitonin-permeabilized cells. Glutamate uptake by permeabilized cells at 10 min was carried out as described in RESEARCH DESIGN AND METHODS, in the presence or absence of the listed compounds. In some experiments, Mg-acetate (–Mg2+) or ATP (–ATP) was omitted. The results are the means ± SE of four independent experiments.

 
The effect of glucose on L-glutamate release.
In the final part of the study, we investigated whether {alpha}TC6 cells secrete glutamate by physiological stimulation. At first, to define the sensitivity of {alpha}TC6 cells to glucose, the cultured cells were further incubated for 2 h in the medium containing either 10 or 1 mmol/l glucose, then glucagon concentrations in the medium were determined according to the procedure previously described (49). In the presence of 1 mmol/l glucose, glucagon (13.3 ± 0.7 ng per 105 cells, n = 4) was released by {alpha}TC6 cells. However, in the presence of 10 mmol/l glucose, glucagon release was reduced to 40 ± 4.4% (n = 4) of that found in the presence of 1 mmol/l glucose. These results were consistent with previous observations (49). When the glucose concentration was lowered as described above, intracellular [Ca2+] increased to 573 ± 64 nmol/l (n = 11), and an appreciable amount of glutamate (1.70 ± 0.22 nmol per 106 cells at 10 min, n = 5) was released from the cells. We observed that <0.38 ± 0.02 nmol glutamate per 106 cells (n = 4) was released when cells were incubated in the presence of 10 mmol/l glucose. When the low glucose treatment was conducted in the absence of Ca2+, neither an increase in intracellular [Ca2+] nor an increase in glutamate release was observed. These results indicate that low glucose treatment stimulates secretion of glutamate as well as glucagon by {alpha}TC6 cells.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Recent biological and biochemical cell studies have indicated that endocrine cells may secrete classical neurotransmitters through an exocytic mechanism. Endocrine cells contain synaptophysin-containing vesicles in which classical neurotransmitters are accumulated (3,50,51). Synaptophysin-containing vesicles contain V-ATPase and vesicular transporters specific to the neurotransmitters and accumulate the corresponding neurotransmitters in a manner dependent on MgATP. For instance, synaptophysin-containing vesicles in pancreatic ß-cells accumulate {gamma}-aminobutyrate (GABA) through a vesicular GABA transporter (52). GABA that has accumulated in the vesicles is exocytized through a Ca2+-dependent regulated secretion pathway (53,54). Upon secretion, GABA may bind to the GABAA receptors in neighboring {alpha}-cells and inhibit glucagon secretion through decreased exocytosis of glucagon-containing secretory granules (55); however, there is also evidence against a role of GABA in the inhibition of glucagon secretion (56). We hypothesized that {alpha}-cells accumulate L-glutamate through a mechanism similar to that observed in ß-cells and secrete the L-glutamate through regulated exocytosis. In the present study, we obtained compelling evidence that {alpha}TC6 cells accumulate L-glutamate in synaptophysin-containing vesicles through a glutamate transporter and secrete it through Ca2+-dependent exocytosis.

Most regulated exocytosis is triggered by an increase in Ca2+ through voltage-gated Ca2+ channels after depolarization. In the first line of evidence, we showed that {alpha}TC6 cells secrete L-glutamate in a manner dependent on Ca2+. Both the KCl-induced Ca2+-dependent glutamate release and the increase in intracellular [Ca2+] were sensitive to antagonists for voltage-gated Ca2+ channels, especially L-type Ca2+ channels, but not N-type or P/Q-type channels, suggesting that L-type Ca2+ channels are involved in KCl-induced Ca2+-dependent glutamate release. The presence of L-type Ca2+ channels in {alpha}TC6 cells was also evident by indirect immunofluorescence microscopy with antibodies against the L-type Ca2+ channel subunit (M.H. and Y.M., unpublished observations).

The second line of evidence is the dependence on temperature and the requirement of an appropriate duration of response after the second stimulation. These properties may reflect complex membrane dynamics, including the charging and discharging of neurotransmitters. Similar phenomena were observed for Ca2+-dependent exocytosis found in various endocrine cells and neuronal cells (17,19,30,31). The relatively slow rate of glutamate release (Fig. 1A) is also similar to that of GABA release by ß-cells and glutamate release by rat pinealocytes (17,19,53,54).

The third line of evidence is sensitivity to BoNT/E and bafilomycin A1. It is well established that BoNT/E is a protease specific to SNAP25. Once activated in sensitive cells, BoNT/E cleaves SNAP25 into low molecular forms, causing inhibition of exocytosis of synaptic vesicles and secretory granules (17,19,373839). The sensitivity to BoNT/E is thus considered one of the criteria for regulated exocytosis. Bafilomycin A1 is a relatively hydrophobic macrolide antibiotic that specifically inhibits V-ATPase–linked secondary transport after the dissipation of an electrochemical proton gradient in the vesicles (45,46). It can deplete L-glutamate in secretory vesicles, resulting in a reduced level of exocytosis by neurons and astrocytes (43,44). Because bafilomycin A1 inhibits the glutamate release from {alpha}TC6 cells, glutamate may accumulate in vesicular structures before being released by {alpha}TC6 cells in the same manner it is released in other secretory vesicles.

We then looked for the organelles responsible for the storage of glutamate. Vesicles containing synaptophysin are candidates for such an organelle because two kinds of synaptophysin-containing organelles, synaptic vesicles in glutamatergic neurons and synaptic-like microvesicles in pinealocytes, are known to be responsible for the storage and exocytosis of glutamate (28,29,45,46). Immunoelectronmicroscopy revealed synaptophysin-containing vesicles in {alpha}TC6 cells (Fig. 4). Synaptophysin-containing vesicles are different from dense-cored vesicles because the latter are devoid of both synaptophysin (Fig. 4D) and glucagon, as revealed with indirect immunofluorescence microscopy (data not shown). Furthermore, we showed that {alpha}TC6 cells accumulated L-glutamate in an ATP-dependent manner sensitive to bafilomycin A1 and proton conductors (Fig. 5). This strongly suggests that a vesicular glutamate transporter energized by V-ATPase is responsible for the accumulation of glutamate. These results suggest that the synaptophysin-containing vesicles are counterparts to endocrine synaptic-like microvesicles and that they are responsible for storage and secretion of L-glutamate in {alpha}TC6 cells.

After stimulation, the glutamate-containing vesicles may secrete internal L-glutamate through Ca2+-dependent exocytic processes. The released glutamate may bind to the glutamate receptor in neighboring islet cells, after which the receptor transmits glutamate signals (4). These mechanisms are essentially similar to the accumulation and release of L-glutamate in glutamatergic synaptic terminals (48) and endocrine pinealocytes (3). It is noteworthy that low glucose treatment stimulates the release of glutamate as well as glucagon from {alpha}TC6 cells. Furthermore, exogenous glutamate stimulates the release of glucagon from rat pancreatic islets (10). These results suggest a regulatory role of L-glutamate in the endocrine function of {alpha}TC6 cells.

In conclusion, {alpha}TC6 cells possess machinery for glutamate signal output. {alpha}TC6 cells may constitute a suitable experimental system for the peripheral glutamatergic mechanism. Our present study may provide an insight into the origin of L-glutamate in the extracellular space of islets, as stated above, and it raises the following questions: 1) Does the regulated exocytosis of L-glutamate occur in {alpha}-cells in islets? and 2) If so, what is the in vivo stimulation that triggers glutamate exocytosis? Further studies are necessary to answer these questions and to determine the entire features of the glutamatergic system in islets.


   ACKNOWLEDGMENTS
 
This work was supported in part by grants from the Japanese Ministry of Education, Science and Culture; CREST; the Japan Science and Technology Corporation; and the Japan Securities Scholarship Foundation. M.H. was supported by the Hayashi Memorial Foundation for Female Natural Scientists. H.Y. is a fellow at the Venture Business Laboratory of Okayama University.

We thank Prof. M. Takahashi of Mitsubishi Life Science Institute for the monoclonal antibodies against synaptophysin and Prof. S. Kozaki of the University of Osaka Prefecture for supplying the botulinum neurotoxin type E. We are also grateful to M. Kinoshita for her excellent technical assistance.


   FOOTNOTES
 
Address correspondence and reprint requests to Dr. Y. Moriyama, Department of Biochemistry, Faculty of Pharmaceutical Sciences, Okayama University, Okayama 700-8530, Japan. E-mail: moriyama{at}pheasant.pharm.okayama-u.ac.jp.

H.Y., M.O., and M.H. contributed equally to this work. H.Y.’s current address is the Department of Biochemistry, Faculty of Medicine, Okayama University, Okayama, Japan.

Received for publication 12 June 2000 and accepted in revised form 19 January 2001.

AM, acetoxymethylester; AMPA, (RS)-{alpha}-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid; BoNT/E, botulinum neurotoxin type E; DMEM, Dulbecco’s modified Eagle medium; GABA, {gamma}-aminobutyrate; HPLC, high-performance liquid chromatography; SF6847, 3,5-di-tert-butyl-4-hydroxybenzylidenemalononitrile; SNAP25, 25 kDa synaptosomal-associated protein.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
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