Activity-Dependent Potentiation of Large Dense-Core Vesicle Release Modulated by Mitogen-Activated Protein Kinase/Extracellularly
http://www.100md.com
《内分泌学杂志》
Systems Biodynamics National Core Research Center (Y.-S.P., D.-J.J., E.-M.H., S.-K.L., K.-T.K.) Division of Molecular and Life Sciences, Pohang University of Science and Technology, Pohang 790-784, Republic of Korea
Department of Life Science and Food Engineering (B.-S.S.), Handong Global University, Pohang 791-708, Republic of Korea
Abstract
Large dense-core vesicles (LDCVs), containing neuropeptides, hormones, and amines, play a crucial role in the activation of the sympathetic nervous system and synaptic modulation. In some secretory cells, LDCVs show activity-dependent potentiation (ADP), which represents enhancement of subsequent exocytosis, compared with the previous one. Here we report the signaling mechanism involved in ADP of LDCV release. First, ADP of LDCV release, induced by repetitive stimulation of nicotinic acetylcholine receptors (nAChRs), was augmented by increasing calcium influx, showing calcium dependence of ADP. Second, translocation of vesicles was involved in ADP. Electron microscope analysis revealed that nAChR stimulation resulted in LDCV translocation to the plasma membrane and increase of fused LDCVs in response to repetitive stimulation was observed by amperometry. Third, we provide evidence for involvement of MAPK signaling in ADP. MAPK signaling was activated by nAChR-induced calcium influx, and ADP as well as vesicle translocation was suppressed by inhibition of MAPK signaling with MAPK kinase blockers, such as PD 098059 and U0126. Fourth, PD 098059 inhibited nAChR stimulation-induced F-actin disassembly, which has been reported to control vesicle translocation. Taken together, we suggest that ADP of LDCV release is modulated by calcium-dependent activation of MAPK signaling via regulating F-actin disassembly.
Introduction
NEURONS AND NEUROENDOCRINE cells contain large dense-core vesicles (LDCVs) and small synaptic vesicles (SSVs) that mediate the release of neurotransmitters (1, 2). LDCVs and SSVs are responsible for secretion of neuropeptides or hormones and classical neurotransmitters, respectively, and have conserved machinery for vesicle fusion events (3, 4). Despite the fact that LDCVs and SSVs have similar machinery for secretion, the two types of vesicles differ in their morphology, release kinetics, and distribution (5). Hormones packaged in LDCVs undergo slow release with prolonged stimulation, whereas exocytosis of SSVs occurs rapidly in response to a single action potential. Besides, SSVs are locally supplied by endocytosis and refilling of neurotransmitters near the plasma membrane, whereas LDCVs are homogeneously distributed and mainly found at more remote locations (5, 6).
Despite these differences, the process of SSV and LDCV release shows activity dependence (7). At some excitable cells, repetitive stimulation induces activity-dependent potentiation (ADP) of exocytosis that represents enhancement of neurotransmitter release, compared with the prior one, although activity-dependent depression of exocytosis is dominant (8). In the case of SSVs, repetitive bursts of presynaptic action potentials give rise to synaptic enhancement through increasing the release of SSVs (8, 9, 10). Potentiation of SSV release at the synapses is attributed to enlargement of the readily releasable pool (RRP) as well as the increase in vesicle-fusion probability (11). It has been suggested that this short-term synaptic enhancement is associated with establishment of short-term memory (12, 13). Similarly, LDCVs, which play a crucial role in the autonomic nervous system by regulating mood, behavior, and synaptic strength, also show ADP (7, 14, 15). In contrast to the case of SSVs, the mechanism of ADP of LDCV release has been poorly understood.
To elucidate the signaling mechanisms involved in ADP of LDCV release, we used chromaffin cells that contain LDCVs, through which catecholamines are secreted. Chromaffin cells are neuroendocrine and modified sympathetic ganglion cells that have been widely used as a model system for studying exocytosis mechanisms of LDCV release (16). Here we show that potentiation of LDCV release by repetitive stimulation of nicotinic acetylcholine receptors (nAChRs) has calcium dependence. ADP of LDCV release is attributed to enlargement of the RRP pool size and increased fused vesicle number, which is controlled by F-actin disassembly. We also suggest that MAPK signaling induced by nAChR activation is involved in ADP of LDCV release via regulating F-actin disassembly that leads to LDCV translocation.
Materials and Methods
Materials
Fura-2 pentaacetoxymethyl ester (fura-2/AM) and rhodamine phalloidin were from Molecular Probes (Eugene, OR). Acetylcholine, 1,1-dimethyl-4-phenylpiperazinium iodide (DMPP), sucrose, cytochalasin B, 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d] pyrimidine, PD 098059, and U0126 were purchased from Sigma (St. Louis, MO). Jasplakinolide was from Calbiochem (San Diego, CA). Anti-ERK and anti-phospho-ERK polyclonal antibody were from Cell Signaling (Beverly, MA). The enhanced chemiluminescence (Supex) kit was from Neuronex (Pohang, Republic of Korea).
Preparation of bovine chromaffin cells
Chromaffin cells were isolated from bovine adrenal gland medulla by two-step collagenase digestion as previously described (17). For amperometric measurement and calcium imaging, cells were grown on poly-D-lysine-coated glass coverslips at the density of 1 x 106 cells/35-mm dish. The cells were maintained in DMEM/F-12 (Invitrogen, Carlsbad, CA) containing 10% fetal bovine serum (Hyclone Laboratories, Logan, UT) and 1% antibiotics (Invitrogen). Chromaffin cells were incubated in a humidified atmosphere of 5% CO2-95% air at 37 C for 1–3 d before use.
Amperometric measurement
Recordings were performed at room temperature as described previously (18). Chromaffin cells were buffered with amine-free solution containing 137.5 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM D-glucose, and 10 mM HEPES (pH 7.3) titrated by NaOH. Carbon-fiber electrodes were fabricated from 8-μm-diameter carbon fibers (Alfa Aesar, Ward Hill, MA). A carbon-fiber electrode back-filled with 3 M KCl connected to the head stage was attached to a single cell. The amperometric current, generated by oxidation of catecholamines, was measured by using an axopatch 200B amplifier (Axon Instruments Inc., Foster City, CA) and operated in the voltage-clamp mode at a holding potential of + 650 mV. Amperometric signals were low-pass filtered at 1 kHz and sampled at 500 Hz. For data acquisition and analysis, pCLAMP 8 software (Axon Instruments) was used. Peak number and peak height of amperometric current were calculated by using IGOR software (Wave Metrics, Lake Oswego, OR). Solutions were exchanged by a local perfusion system that allows complete exchange of medium bathing the cells within 2 sec.
Intracellular free calcium concentration ([Ca2+]i) imaging
As described previously (19), calcium imaging experiments were performed with a monochromator-based spectrofluorometric system (DeltaScan Illumination System, DeltaScan Photon Technology International, Inc., Birmingham, NJ). We performed calcium imaging of single cells with fura-2/AM, and the calcium signal was acquired over the whole cell.
Electron microscopy
Bovine chromaffin cells grown on vitrogen collagen matrix (Cohesion, Palo Alto, CA) were stimulated and washed out with Locke’s solution containing 157.4 mM NaCl, 5.6 mM KCl, 2.2 mM CaCl2, 1.2 mM MgCl2, 5.6 mM D-glucose, 5 mM HEPES, and 3.6 mM NaHCO3 (pH 7.4) titrated by NaOH. As described previously (20), these cells were rinsed two times with PBS and fixed with 2% paraformaldehyde and 2% glutaraldehyde in 0.05 M sodium cacodylate buffer (pH 7.4) for 20 min at room temperature. Cells were subsequently postfixed with 0.5% osmium tetroxide in 0.05 M sodium cacodylate buffer (pH 7.4) for 30 min at room temperature. Cells were further dehydrated in graded ethanol solutions and embedded in LR White resin (London Resin Co., Berkshire, UK). The resin was cured at 60 C for 24 h. Silver-gold thin sections were stained with uranyl acetate and lead citrate. The thin sections were examined under JEOL 1200 EX2 transmission electron microscope at 80 kV.
Immunoblotting
Cells treated as indicated were washed with chilled PBS. Proteins were extracted with lysis buffer [10 mM Tris-HCl (pH 7.4), 1 mM EDTA, 0.5 mM EGTA, 10 mM NaCl, 1 mM phenylmethylsulfonyl fluoride, 1 mM Na3VO4, 1 mM dithiothreitol, 2 mM ascorbic acid, and 1% Nonidet P-40]. Proteins were then separated by electrophoresis containing 0.1% sodium dodecyl sulfate and transferred to a nitrocellulose membrane, which was blocked with 5% nonfat dry milk in a solution of 20 mM Tris-HCl (pH 7.4), 140 mM NaCl, and 0.05% Tween 20. ERK and phospho-ERK were detected with anti-ERK and phospho-ERK primary antibodies and horseradish peroxidase-conjugated secondary antibodies (Cell Signaling). Bands were visualized with the ECL detection system (Neuronex).
F-actin disassembly
Staining of F-actin was performed in chromaffin cells as previously described (21) with minor modification. Briefly, cells plated on poly-D-lysine-coated coverslips were fixed in 3.7% formaldehyde solution for 10 min and then permeabilized by 0.1% Triton X-100. After washout with PBS, cells were stained for F-actin with 0.5 U/ml rhodamine-phalloidin for 20 min at room temperature. The slides were observed under Radiance 2100 confocal laser-scanning microscope (Bio-Rad Laboratories, Hemel Hempstead, UK). Each cell examined was classified as having either a continuous or discontinuous cortical rhodamine fluorescent ring.
Statistical analysis
All quantitative data are presented as means ± SEM. Comparison between two groups were analyzed by using Student’s t test, and values of P < 0.05 were considered to be significant.
Results
ADP of LDCV release in chromaffin cells
Amperometry has been used to monitor the release of catecholamines, which is stored in LDCVs (22, 23). Upon acetylcholine stimulation at the splanchnic nerve terminals of the sympathetic nervous system, chromaffin cells release catecholamines stored in LDCVs. In chromaffin cells, application of DMPP, an agonist of nAChRs, induces depolarization and subsequent activation of voltage-gated calcium channels (VGCCs), which leads to calcium-dependent fusion of secretory vesicles. We observed that repetitive stimulation of DMPP resulted in potentiation of the subsequent exocytotic events, compared with the previous ones, i.e. ADP of LDCV release. The extent of ADP was increased until the third stimulation and sustained up to the fifth stimulation (Fig. 1A). ADP was observed even when interval time between the first and second stimulation was 30 min (Fig. 1B). Acetylcholine, an endogenous agonist of nAChRs, also induced ADP (Fig. 1C). Because both the peak number (representing fused vesicles) and the peak height (representing catecholamine contents in a single vesicle) of amperometric currents (24) were increased, ADP might be associated with the increase in the probability of vesicle release as well as amount of catecholamine (Fig. 1C).
The extent of ADP depends on the amount of calcium influx, but ADP is not due to accumulation of calcium
Because it has been well established that intracellular calcium controls vesicle release (7, 25), we tested calcium dependence of ADP via elucidating the relationship between the amount of calcium influx and ADP. Different concentrations of KCl (Fig. 2A) or extracellular calcium ([Ca2+]o) (Fig. 2B) were used to change the amount of calcium influx. Calcium influx upon repetitive stimulation of KCl was similar, showing that repetitive stimulation does not induce calcium accumulation (Fig. 2A). Interestingly, when the amount of calcium influx was larger, potentiation effect was more prominent (Fig. 2, A and B). ADP induced by repetitive stimulation of 50 mM KCl was more prominent than that induced by 30 mM KCl (Fig. 2A). Similarly, repetitive stimulation of DMPP did not cause calcium accumulation (Fig. 2B). Furthermore, repetitive stimulation with high [Ca2+]o augmented the effect of ADP, compared with low [Ca2+]o. In addition, ADP was impaired in cells loaded with bis-(o-aminophenoxy)-ethane-N,N,N',N'-tetraacetic acid/AM, which chelates calcium ions (data not shown). It has been reported that hypertonic sucrose solution depletes the docked and fusion-competent vesicles of the RRP in a calcium-independent manner (26). Therefore, we applied 500 mM hypertonic sucrose as a control and observed that ADP was not induced by repetitive application of 500 mM hypertonic sucrose solution (Fig. 2C). Taken together, these results suggest that ADP of LDCV release shows calcium dependence, although repetitive stimulation does not induce calcium accumulation.
ERK signaling activated by calcium influx is involved in ADP
To investigate the signaling mechanisms involved in ADP, we examined factors regulated by calcium influx due to the calcium dependence of ADP. Calcium influx activates ERK signaling in neuronal cells (27, 28). In agreement with previous reports, the level of phospho-ERK was elevated upon stimulation of DMPP in chromaffin cells (Fig. 3A) (29, 30). Removal of extracellular calcium ions blocked phosphorylation of ERK induced by DMPP stimulation (Fig. 3B), confirming that activation of ERK depends on calcium influx. Although prolonged stimulation of nAChRs activates ERK signaling (29, 30), we focused on repetitive stimulation-induced ERK activation and monitored the phospho-ERK level induced by repetitive stimulation with DMPP. ERK signaling, which keeps basal level during the first stimulation, was activated after the first stimulation (Fig. 3C). After the second stimulation, phospho-ERK level was more accumulated and sustained, whereas the third stimulation did not trigger additional enhancement, compared with the second stimulation. ADP, which is sustained after the third stimulation (Fig. 1A), is almost correlated with kinetics of ERK activation evoked by repetitive stimulation. In addition, time course of ADP is also correlated with kinetics of the first stimulation-induced ERK activation. The second stimulation-induced LDCV release was strongly potentiated when interval time was 2–15 min and decreased after 30 min (Fig. 1B). Interestingly, phospho-ERK level after 20 sec DMPP stimulation was increased and sustained after 2 min and decreased after 30 min (Fig. 3D). The extent of ERK activation induced by repetitive stimulation does not exactly coincide with the pattern of activity-dependent manner; however, kinetics of ERK activation after the first stimulation seems to represent the extent of ADP of LDCV release.
To investigate the involvement of ERK signaling pathway in ADP, we examined the effects of PD 098059 and U0126, blockers of MAPK kinase. As shown in Fig. 3C, both inhibitors significantly inhibited ADP without affecting the first stimulation-induced basal secretion (Fig. 3E). Effects of PD 098059 and U0126 were confirmed by monitoring phosphorylation of ERK (Fig. 3B). In addition, the phospho-ERK level was elevated by increasing [Ca2+]o concentration (Fig. 3D), a condition that enhanced ADP (Fig. 2). Taken together, these results suggest that activation of ERK is involved in ADP of LDCV release.
ERK signaling is involved in the activity-induced LDCV translocation
ADP of SSVs is ascribed to activity-dependent increase of vesicle pools located near the plasma membrane. To test whether stimulation of DMPP causes any change of LDCV distribution near the plasma membrane in chromaffin cells, we performed electron microscope analysis. Stimulation with DMPP increased the number of vesicles located near the plasma membrane within 500 nm from the plasma membrane, showing activity-induced change of the LDCV distribution (Fig. 4). Therefore, activation of nAChR evokes calcium influx to trigger vesicle translocation, thereby increasing docked vesicle pools. This enlargement of the docked vesicle pool size (Fig. 4B) correlated with the increase in the amperometric peak number (representing fused vesicle number) (Fig. 1C), suggesting that translocation of vesicles might contribute to ADP. We next examined the effect of ERK signaling on the activity-induced change of LDCV distribution. Preincubation with PD 098059 blocked the DMPP-induced recruitment of LDCVs near the plasma membrane (Fig. 4B), suggesting that ERK signaling is involved in the activity-induced translocation of LDCVs.
ERK signaling modulates F-actin disassembly induced by nAChR activation
Vesicle translocation is regulated by cortical actin network dynamics (31). To investigate whether rearrangement of F-actin is responsible for ADP, we examined the effects of jasplakinolide, an F-actin stabilizing drug. Treatment of cells with jasplakinolide blocked ADP, showing the involvement of F-actin rearrangement in ADP (Fig. 5A). In addition, we tested the effect of cytochalasin B, an actin filament destabilizing agent, which has often been used to inhibit vesicle translocation from RP into RRP and vesicle mobilization from RRP to exocytosis (32, 33, 34). Blocking vesicle translocation and vesicle mobilization by treatment with cytochalasin B also resulted in inhibition of ADP (Fig. 5A). Although both cytochalasin B and jasplakinolide inhibited ADP, cytochalasin B-treated cells showed increased level of catecholamine release from the first stimulation (Fig. 5B). Cytochalasin B-induced basal increase in LDCV release is associated with F-actin disassembly, which mobilizes LDCVs. Taken together, it can be suggested that cortical actin dynamics, which regulate vesicle translocation, is involved in ADP. Next, we thus examined whether ERK signaling is involved in F-actin disassembly. In chromaffin cells, it has been well known that disruption of cortical F-actin induces translocation of vesicles to the plasma membrane, thereby increasing exocytosis (21, 31). F-actin disassembly was increased by DMPP stimulation as shown by images of a fragmented fluorescent ring (Fig. 5, C and D), and treatment with PD 098059 suppressed the DMPP-induced F-actin disassembly (Fig. 5D), suggesting that vesicle translocation induced by F-actin disassembly upon DMPP stimulation can be blocked by inhibition of ERK signaling.
Discussion
In this study, we show that repetitive stimulation of nAChRs induces ADP of LDCV release in chromaffin cells (Fig. 1) and suggest that ADP is caused by vesicle translocation but not calcium accumulation. The amount of calcium influx positively correlates with the extent of ADP, showing calcium dependence of ADP (Fig. 2). We provide evidence that calcium-dependent activation of ERK signaling is involved in ADP. Inhibition of ERK signaling suppressed activity-dependent translocation of LDCVs (Fig. 4) as well as DMPP-induced F-actin disassembly (Fig. 5D). We therefore suggest that ERK activation is involved in ADP via regulating F-actin disassembly, which controls vesicle translocation. To our knowledge, this is the first report to investigate the molecular mechanism involved in ADP of LDCV release by using amperometry, which directly detects exocytosis of catecholamine release.
It has been suggested that ERK signaling modulates nicotinic receptor-mediated catecholamine secretion (29, 30). The above studies demonstrated that treatment of PD098059, a MAPK kinase inhibitor, attenuated nicotine-induced ERK activation and catecholamine release. However, signaling mechanisms how ERK activation mediates catecholamine release were not suggested. Here we characterized ADP of LDCV release and suggested the involvement of ERK signaling in ADP via regulating F-actin disassembly and vesicle translocation, confirmed by electron microscope (Fig. 4) and the increase in amperometric peak number (Fig. 1). In addition, ERK signaling plays a role in general secretion of catecholamine (29, 30), although we focused on its involvement in repetitive stimulation-induced exocytosis. First, blockers of ERK signaling did not inhibit the first stimulation-induced basal secretion but ADP of LDCV release (Fig. 3E). Second, the second stimulation-induced LDCV release was dramatically increased when interval time was 2–15 min and decreased after 30 min (Fig. 1B). Kinetics of ERK activation after the first DMPP simulation (Fig. 3D) is almost correlated with the extent of activity-dependent potentiation of LDCV release, thereby suggesting the involvement of ERK signaling in ADP.
Amperometry is a powerful tool that detects single vesicle fusion in real time and recognizes the release of LDCV, which contain catecholamines. For these reasons, amperometric analysis can be a useful tool to investigate ADP of LDCV release, eliminating other effects, such as endocytosis and involvement of SSVs. Amperometric analysis revealed that the number of fusion events (peak number of amperometric currents), as well as amount of catecholamine release (integration of amperometric current), was increased in an activity-dependent manner (Fig. 1C).
Using an electron microscope, we show that stimulation of nAChRs increases the size of vesicle pools located near the plasma membrane (Fig. 4). Because LDCVs in the vicinity of the plasma membrane can be easily released upon stimulation, they might be referred to as the RRP in chromaffin cells (6, 35). LDCVs, located within 500 nm of the plasma membrane, can be classified as the RRP, which represents fusion-competent vesicles (35). Together with the analysis of amperometric currents (Fig. 1C), we suggest that the vesicle size of the RRP is increased by stimulation of nAChRs.
It has been reported that translocation of SSVs causes potentiation of exocytosis, and this might be the underlying mechanism for long-term potentiation (36). At presynaptic terminals, it has been suggested that calcium accumulation induced by repetitive stimulation plays a prominent role in activity-dependent augmentation of SSV release (10, 12). Our data suggest that ADP of LDCV release is also caused by vesicle translocation. However, repetitive stimulation did not result in calcium accumulation (Fig. 2), suggesting that the mechanism involved in ADP of LDCV release is different from that of SSV release.
In neuronal cells, it has been widely recognized that calcium influx via VGCCs can activate MAPK signaling via transactivation of tyrosine kinases and epithelial growth factor receptor (28). Here we report that MAPK signaling, which is potently activated by calcium influx, affects ADP of LDCV release via regulating vesicle translocation. Inhibition of MAPK signaling blocks the effect of ADP (Fig. 3) and suppresses the activity-induced change of LDCV distribution, which represents vesicle translocation (Fig. 4). It has been shown that disruption of cortical F-actin can evoke vesicle translocation via liberating LDCVs bound to F-actin in chromaffin cells (21, 37, 38). In agreement with the previous studies, we also observed that catecholamine release induced by first DMPP stimulation under treatment of cytochalasin B (a drug for F-actin disassembly) was greater, compared with control (Fig. 5B), suggesting that the increase in LDCV release by first DMPP stimulation under cytochalasin B treatment might be attributed to the increase of vesicle translocation caused by F-actin disassembly. Moreover, our data provide evidence for the involvement of F-actin disassembly in the potentiation of exocytosis. Stimulation of nAChRs induced F-actin disassembly (Fig. 5B), and ADP was inhibited by treatment with jasplakinolide, an inhibitor of F-actin disassembly (Fig. 5A). We also provide evidence that MAPK signaling is involved in regulation of F-actin disassembly, thereby modulating vesicle translocation. However, we cannot eliminate possibilities that MAPK signaling also can be involved in the processes of vesicle docking or fusion events.
MAPK signaling has been suggested to play an important role in the regulation of short-term synaptic plasticity via controlling F-actin disassembly at the synapses. The underlying mechanism for the regulation of synaptic plasticity has been proposed to be modulation of F-actin dynamics at the synapses by activation of MAPK, thereby regulating ADP of SSV release (8, 10, 39, 40). Calcium influx via VGCCs makes actin filament disassembled, which liberates SSVs tethered to F-actin, and finally induces SSV translocation to the plasma membrane (10, 40). In neurons, it has been well demonstrated that MAPK activation by calcium influx leads to phosphorylation of synapsin I, which causes F-actin disassembly and subsequent translocation of SSVs (38, 41). However, synapsin is not expressed in chromaffin cells (42, 43). Therefore, identifying target molecules, downstream of MAPK signaling as well as their involvement in F-actin disassembly requires further investigation.
Chromaffin cells are repetitively stimulated by acetylcholine released from the splanchnic nerve terminals (14). They are the main source for generating high level of catecholamine in the bloodstream when sympathetic nervous system is activated. Stress and fight-or-flight response are typical physiological actions of the sympathetic nervous system and primarily caused by the release of catecholamine from chromaffin cells (44). These catecholamines facilitate immediate physical reactions such as increasing heart rate and breathing, constricting blood vessels, and controlling lipolysis and glycogenolysis (45). Upon activation of the sympathetic nervous system, it is important to maintain a high level of catecholamine in the bloodstream for various physiological actions. Activity-dependent depression of transmitter release, prevalent at most synapses, is attributed to a depletion of the RRP of vesicles (8). Also, repetitive stimulation on secretory cells usually gives rise to desensitization of exocytosis. Interestingly, chromaffin cells show ADP of catecholamine release. We therefore suggest that ADP of LDCV release in chromaffin cells might be an important mechanism to establish a high level of catecholamine in the bloodstream upon repetitive activation of the sympathetic nervous system.
Acknowledgments
We are grateful to Ms. Soo-Jin Kim for electron microscopy instrumentation. We also thank Mr. Kyung-Chul Woo for riding and Byung-Soon Kang (Kyung-Buk Packers Co. Inc., Pohang, Republic of Korea) for providing bovine adrenal glands.
Footnotes
This work was supported by the Brain Neurobiology Research Program (M10412000023-02310) and Systems Biodynamics National Core Research Center sponsored by the Korean Ministry of Science and Technology and Brain Korea 21 Program of the Koran Ministry of Education.
The authors have nothing to declare.
First Published Online November 23, 2005
Abbreviations: ADP, Activity-dependent potentiation; [Ca2+]i, intracellular free calcium concentration; [Ca2+]o, extracellular calcium; DMPP, 1,1-dimethyl-4-phenylpiperazinium iodide; fura-2/AM, fura-2 pentaacetoxymethyl ester; LDCV, large dense-core vesicle; nAChR, nicotinic acetylcholine receptor; RRP, readily releasable pool; SSV, small synaptic vesicle; VGCC, voltage-gated calcium channel.
Accepted for publication November 17, 2005.
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Department of Life Science and Food Engineering (B.-S.S.), Handong Global University, Pohang 791-708, Republic of Korea
Abstract
Large dense-core vesicles (LDCVs), containing neuropeptides, hormones, and amines, play a crucial role in the activation of the sympathetic nervous system and synaptic modulation. In some secretory cells, LDCVs show activity-dependent potentiation (ADP), which represents enhancement of subsequent exocytosis, compared with the previous one. Here we report the signaling mechanism involved in ADP of LDCV release. First, ADP of LDCV release, induced by repetitive stimulation of nicotinic acetylcholine receptors (nAChRs), was augmented by increasing calcium influx, showing calcium dependence of ADP. Second, translocation of vesicles was involved in ADP. Electron microscope analysis revealed that nAChR stimulation resulted in LDCV translocation to the plasma membrane and increase of fused LDCVs in response to repetitive stimulation was observed by amperometry. Third, we provide evidence for involvement of MAPK signaling in ADP. MAPK signaling was activated by nAChR-induced calcium influx, and ADP as well as vesicle translocation was suppressed by inhibition of MAPK signaling with MAPK kinase blockers, such as PD 098059 and U0126. Fourth, PD 098059 inhibited nAChR stimulation-induced F-actin disassembly, which has been reported to control vesicle translocation. Taken together, we suggest that ADP of LDCV release is modulated by calcium-dependent activation of MAPK signaling via regulating F-actin disassembly.
Introduction
NEURONS AND NEUROENDOCRINE cells contain large dense-core vesicles (LDCVs) and small synaptic vesicles (SSVs) that mediate the release of neurotransmitters (1, 2). LDCVs and SSVs are responsible for secretion of neuropeptides or hormones and classical neurotransmitters, respectively, and have conserved machinery for vesicle fusion events (3, 4). Despite the fact that LDCVs and SSVs have similar machinery for secretion, the two types of vesicles differ in their morphology, release kinetics, and distribution (5). Hormones packaged in LDCVs undergo slow release with prolonged stimulation, whereas exocytosis of SSVs occurs rapidly in response to a single action potential. Besides, SSVs are locally supplied by endocytosis and refilling of neurotransmitters near the plasma membrane, whereas LDCVs are homogeneously distributed and mainly found at more remote locations (5, 6).
Despite these differences, the process of SSV and LDCV release shows activity dependence (7). At some excitable cells, repetitive stimulation induces activity-dependent potentiation (ADP) of exocytosis that represents enhancement of neurotransmitter release, compared with the prior one, although activity-dependent depression of exocytosis is dominant (8). In the case of SSVs, repetitive bursts of presynaptic action potentials give rise to synaptic enhancement through increasing the release of SSVs (8, 9, 10). Potentiation of SSV release at the synapses is attributed to enlargement of the readily releasable pool (RRP) as well as the increase in vesicle-fusion probability (11). It has been suggested that this short-term synaptic enhancement is associated with establishment of short-term memory (12, 13). Similarly, LDCVs, which play a crucial role in the autonomic nervous system by regulating mood, behavior, and synaptic strength, also show ADP (7, 14, 15). In contrast to the case of SSVs, the mechanism of ADP of LDCV release has been poorly understood.
To elucidate the signaling mechanisms involved in ADP of LDCV release, we used chromaffin cells that contain LDCVs, through which catecholamines are secreted. Chromaffin cells are neuroendocrine and modified sympathetic ganglion cells that have been widely used as a model system for studying exocytosis mechanisms of LDCV release (16). Here we show that potentiation of LDCV release by repetitive stimulation of nicotinic acetylcholine receptors (nAChRs) has calcium dependence. ADP of LDCV release is attributed to enlargement of the RRP pool size and increased fused vesicle number, which is controlled by F-actin disassembly. We also suggest that MAPK signaling induced by nAChR activation is involved in ADP of LDCV release via regulating F-actin disassembly that leads to LDCV translocation.
Materials and Methods
Materials
Fura-2 pentaacetoxymethyl ester (fura-2/AM) and rhodamine phalloidin were from Molecular Probes (Eugene, OR). Acetylcholine, 1,1-dimethyl-4-phenylpiperazinium iodide (DMPP), sucrose, cytochalasin B, 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d] pyrimidine, PD 098059, and U0126 were purchased from Sigma (St. Louis, MO). Jasplakinolide was from Calbiochem (San Diego, CA). Anti-ERK and anti-phospho-ERK polyclonal antibody were from Cell Signaling (Beverly, MA). The enhanced chemiluminescence (Supex) kit was from Neuronex (Pohang, Republic of Korea).
Preparation of bovine chromaffin cells
Chromaffin cells were isolated from bovine adrenal gland medulla by two-step collagenase digestion as previously described (17). For amperometric measurement and calcium imaging, cells were grown on poly-D-lysine-coated glass coverslips at the density of 1 x 106 cells/35-mm dish. The cells were maintained in DMEM/F-12 (Invitrogen, Carlsbad, CA) containing 10% fetal bovine serum (Hyclone Laboratories, Logan, UT) and 1% antibiotics (Invitrogen). Chromaffin cells were incubated in a humidified atmosphere of 5% CO2-95% air at 37 C for 1–3 d before use.
Amperometric measurement
Recordings were performed at room temperature as described previously (18). Chromaffin cells were buffered with amine-free solution containing 137.5 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM D-glucose, and 10 mM HEPES (pH 7.3) titrated by NaOH. Carbon-fiber electrodes were fabricated from 8-μm-diameter carbon fibers (Alfa Aesar, Ward Hill, MA). A carbon-fiber electrode back-filled with 3 M KCl connected to the head stage was attached to a single cell. The amperometric current, generated by oxidation of catecholamines, was measured by using an axopatch 200B amplifier (Axon Instruments Inc., Foster City, CA) and operated in the voltage-clamp mode at a holding potential of + 650 mV. Amperometric signals were low-pass filtered at 1 kHz and sampled at 500 Hz. For data acquisition and analysis, pCLAMP 8 software (Axon Instruments) was used. Peak number and peak height of amperometric current were calculated by using IGOR software (Wave Metrics, Lake Oswego, OR). Solutions were exchanged by a local perfusion system that allows complete exchange of medium bathing the cells within 2 sec.
Intracellular free calcium concentration ([Ca2+]i) imaging
As described previously (19), calcium imaging experiments were performed with a monochromator-based spectrofluorometric system (DeltaScan Illumination System, DeltaScan Photon Technology International, Inc., Birmingham, NJ). We performed calcium imaging of single cells with fura-2/AM, and the calcium signal was acquired over the whole cell.
Electron microscopy
Bovine chromaffin cells grown on vitrogen collagen matrix (Cohesion, Palo Alto, CA) were stimulated and washed out with Locke’s solution containing 157.4 mM NaCl, 5.6 mM KCl, 2.2 mM CaCl2, 1.2 mM MgCl2, 5.6 mM D-glucose, 5 mM HEPES, and 3.6 mM NaHCO3 (pH 7.4) titrated by NaOH. As described previously (20), these cells were rinsed two times with PBS and fixed with 2% paraformaldehyde and 2% glutaraldehyde in 0.05 M sodium cacodylate buffer (pH 7.4) for 20 min at room temperature. Cells were subsequently postfixed with 0.5% osmium tetroxide in 0.05 M sodium cacodylate buffer (pH 7.4) for 30 min at room temperature. Cells were further dehydrated in graded ethanol solutions and embedded in LR White resin (London Resin Co., Berkshire, UK). The resin was cured at 60 C for 24 h. Silver-gold thin sections were stained with uranyl acetate and lead citrate. The thin sections were examined under JEOL 1200 EX2 transmission electron microscope at 80 kV.
Immunoblotting
Cells treated as indicated were washed with chilled PBS. Proteins were extracted with lysis buffer [10 mM Tris-HCl (pH 7.4), 1 mM EDTA, 0.5 mM EGTA, 10 mM NaCl, 1 mM phenylmethylsulfonyl fluoride, 1 mM Na3VO4, 1 mM dithiothreitol, 2 mM ascorbic acid, and 1% Nonidet P-40]. Proteins were then separated by electrophoresis containing 0.1% sodium dodecyl sulfate and transferred to a nitrocellulose membrane, which was blocked with 5% nonfat dry milk in a solution of 20 mM Tris-HCl (pH 7.4), 140 mM NaCl, and 0.05% Tween 20. ERK and phospho-ERK were detected with anti-ERK and phospho-ERK primary antibodies and horseradish peroxidase-conjugated secondary antibodies (Cell Signaling). Bands were visualized with the ECL detection system (Neuronex).
F-actin disassembly
Staining of F-actin was performed in chromaffin cells as previously described (21) with minor modification. Briefly, cells plated on poly-D-lysine-coated coverslips were fixed in 3.7% formaldehyde solution for 10 min and then permeabilized by 0.1% Triton X-100. After washout with PBS, cells were stained for F-actin with 0.5 U/ml rhodamine-phalloidin for 20 min at room temperature. The slides were observed under Radiance 2100 confocal laser-scanning microscope (Bio-Rad Laboratories, Hemel Hempstead, UK). Each cell examined was classified as having either a continuous or discontinuous cortical rhodamine fluorescent ring.
Statistical analysis
All quantitative data are presented as means ± SEM. Comparison between two groups were analyzed by using Student’s t test, and values of P < 0.05 were considered to be significant.
Results
ADP of LDCV release in chromaffin cells
Amperometry has been used to monitor the release of catecholamines, which is stored in LDCVs (22, 23). Upon acetylcholine stimulation at the splanchnic nerve terminals of the sympathetic nervous system, chromaffin cells release catecholamines stored in LDCVs. In chromaffin cells, application of DMPP, an agonist of nAChRs, induces depolarization and subsequent activation of voltage-gated calcium channels (VGCCs), which leads to calcium-dependent fusion of secretory vesicles. We observed that repetitive stimulation of DMPP resulted in potentiation of the subsequent exocytotic events, compared with the previous ones, i.e. ADP of LDCV release. The extent of ADP was increased until the third stimulation and sustained up to the fifth stimulation (Fig. 1A). ADP was observed even when interval time between the first and second stimulation was 30 min (Fig. 1B). Acetylcholine, an endogenous agonist of nAChRs, also induced ADP (Fig. 1C). Because both the peak number (representing fused vesicles) and the peak height (representing catecholamine contents in a single vesicle) of amperometric currents (24) were increased, ADP might be associated with the increase in the probability of vesicle release as well as amount of catecholamine (Fig. 1C).
The extent of ADP depends on the amount of calcium influx, but ADP is not due to accumulation of calcium
Because it has been well established that intracellular calcium controls vesicle release (7, 25), we tested calcium dependence of ADP via elucidating the relationship between the amount of calcium influx and ADP. Different concentrations of KCl (Fig. 2A) or extracellular calcium ([Ca2+]o) (Fig. 2B) were used to change the amount of calcium influx. Calcium influx upon repetitive stimulation of KCl was similar, showing that repetitive stimulation does not induce calcium accumulation (Fig. 2A). Interestingly, when the amount of calcium influx was larger, potentiation effect was more prominent (Fig. 2, A and B). ADP induced by repetitive stimulation of 50 mM KCl was more prominent than that induced by 30 mM KCl (Fig. 2A). Similarly, repetitive stimulation of DMPP did not cause calcium accumulation (Fig. 2B). Furthermore, repetitive stimulation with high [Ca2+]o augmented the effect of ADP, compared with low [Ca2+]o. In addition, ADP was impaired in cells loaded with bis-(o-aminophenoxy)-ethane-N,N,N',N'-tetraacetic acid/AM, which chelates calcium ions (data not shown). It has been reported that hypertonic sucrose solution depletes the docked and fusion-competent vesicles of the RRP in a calcium-independent manner (26). Therefore, we applied 500 mM hypertonic sucrose as a control and observed that ADP was not induced by repetitive application of 500 mM hypertonic sucrose solution (Fig. 2C). Taken together, these results suggest that ADP of LDCV release shows calcium dependence, although repetitive stimulation does not induce calcium accumulation.
ERK signaling activated by calcium influx is involved in ADP
To investigate the signaling mechanisms involved in ADP, we examined factors regulated by calcium influx due to the calcium dependence of ADP. Calcium influx activates ERK signaling in neuronal cells (27, 28). In agreement with previous reports, the level of phospho-ERK was elevated upon stimulation of DMPP in chromaffin cells (Fig. 3A) (29, 30). Removal of extracellular calcium ions blocked phosphorylation of ERK induced by DMPP stimulation (Fig. 3B), confirming that activation of ERK depends on calcium influx. Although prolonged stimulation of nAChRs activates ERK signaling (29, 30), we focused on repetitive stimulation-induced ERK activation and monitored the phospho-ERK level induced by repetitive stimulation with DMPP. ERK signaling, which keeps basal level during the first stimulation, was activated after the first stimulation (Fig. 3C). After the second stimulation, phospho-ERK level was more accumulated and sustained, whereas the third stimulation did not trigger additional enhancement, compared with the second stimulation. ADP, which is sustained after the third stimulation (Fig. 1A), is almost correlated with kinetics of ERK activation evoked by repetitive stimulation. In addition, time course of ADP is also correlated with kinetics of the first stimulation-induced ERK activation. The second stimulation-induced LDCV release was strongly potentiated when interval time was 2–15 min and decreased after 30 min (Fig. 1B). Interestingly, phospho-ERK level after 20 sec DMPP stimulation was increased and sustained after 2 min and decreased after 30 min (Fig. 3D). The extent of ERK activation induced by repetitive stimulation does not exactly coincide with the pattern of activity-dependent manner; however, kinetics of ERK activation after the first stimulation seems to represent the extent of ADP of LDCV release.
To investigate the involvement of ERK signaling pathway in ADP, we examined the effects of PD 098059 and U0126, blockers of MAPK kinase. As shown in Fig. 3C, both inhibitors significantly inhibited ADP without affecting the first stimulation-induced basal secretion (Fig. 3E). Effects of PD 098059 and U0126 were confirmed by monitoring phosphorylation of ERK (Fig. 3B). In addition, the phospho-ERK level was elevated by increasing [Ca2+]o concentration (Fig. 3D), a condition that enhanced ADP (Fig. 2). Taken together, these results suggest that activation of ERK is involved in ADP of LDCV release.
ERK signaling is involved in the activity-induced LDCV translocation
ADP of SSVs is ascribed to activity-dependent increase of vesicle pools located near the plasma membrane. To test whether stimulation of DMPP causes any change of LDCV distribution near the plasma membrane in chromaffin cells, we performed electron microscope analysis. Stimulation with DMPP increased the number of vesicles located near the plasma membrane within 500 nm from the plasma membrane, showing activity-induced change of the LDCV distribution (Fig. 4). Therefore, activation of nAChR evokes calcium influx to trigger vesicle translocation, thereby increasing docked vesicle pools. This enlargement of the docked vesicle pool size (Fig. 4B) correlated with the increase in the amperometric peak number (representing fused vesicle number) (Fig. 1C), suggesting that translocation of vesicles might contribute to ADP. We next examined the effect of ERK signaling on the activity-induced change of LDCV distribution. Preincubation with PD 098059 blocked the DMPP-induced recruitment of LDCVs near the plasma membrane (Fig. 4B), suggesting that ERK signaling is involved in the activity-induced translocation of LDCVs.
ERK signaling modulates F-actin disassembly induced by nAChR activation
Vesicle translocation is regulated by cortical actin network dynamics (31). To investigate whether rearrangement of F-actin is responsible for ADP, we examined the effects of jasplakinolide, an F-actin stabilizing drug. Treatment of cells with jasplakinolide blocked ADP, showing the involvement of F-actin rearrangement in ADP (Fig. 5A). In addition, we tested the effect of cytochalasin B, an actin filament destabilizing agent, which has often been used to inhibit vesicle translocation from RP into RRP and vesicle mobilization from RRP to exocytosis (32, 33, 34). Blocking vesicle translocation and vesicle mobilization by treatment with cytochalasin B also resulted in inhibition of ADP (Fig. 5A). Although both cytochalasin B and jasplakinolide inhibited ADP, cytochalasin B-treated cells showed increased level of catecholamine release from the first stimulation (Fig. 5B). Cytochalasin B-induced basal increase in LDCV release is associated with F-actin disassembly, which mobilizes LDCVs. Taken together, it can be suggested that cortical actin dynamics, which regulate vesicle translocation, is involved in ADP. Next, we thus examined whether ERK signaling is involved in F-actin disassembly. In chromaffin cells, it has been well known that disruption of cortical F-actin induces translocation of vesicles to the plasma membrane, thereby increasing exocytosis (21, 31). F-actin disassembly was increased by DMPP stimulation as shown by images of a fragmented fluorescent ring (Fig. 5, C and D), and treatment with PD 098059 suppressed the DMPP-induced F-actin disassembly (Fig. 5D), suggesting that vesicle translocation induced by F-actin disassembly upon DMPP stimulation can be blocked by inhibition of ERK signaling.
Discussion
In this study, we show that repetitive stimulation of nAChRs induces ADP of LDCV release in chromaffin cells (Fig. 1) and suggest that ADP is caused by vesicle translocation but not calcium accumulation. The amount of calcium influx positively correlates with the extent of ADP, showing calcium dependence of ADP (Fig. 2). We provide evidence that calcium-dependent activation of ERK signaling is involved in ADP. Inhibition of ERK signaling suppressed activity-dependent translocation of LDCVs (Fig. 4) as well as DMPP-induced F-actin disassembly (Fig. 5D). We therefore suggest that ERK activation is involved in ADP via regulating F-actin disassembly, which controls vesicle translocation. To our knowledge, this is the first report to investigate the molecular mechanism involved in ADP of LDCV release by using amperometry, which directly detects exocytosis of catecholamine release.
It has been suggested that ERK signaling modulates nicotinic receptor-mediated catecholamine secretion (29, 30). The above studies demonstrated that treatment of PD098059, a MAPK kinase inhibitor, attenuated nicotine-induced ERK activation and catecholamine release. However, signaling mechanisms how ERK activation mediates catecholamine release were not suggested. Here we characterized ADP of LDCV release and suggested the involvement of ERK signaling in ADP via regulating F-actin disassembly and vesicle translocation, confirmed by electron microscope (Fig. 4) and the increase in amperometric peak number (Fig. 1). In addition, ERK signaling plays a role in general secretion of catecholamine (29, 30), although we focused on its involvement in repetitive stimulation-induced exocytosis. First, blockers of ERK signaling did not inhibit the first stimulation-induced basal secretion but ADP of LDCV release (Fig. 3E). Second, the second stimulation-induced LDCV release was dramatically increased when interval time was 2–15 min and decreased after 30 min (Fig. 1B). Kinetics of ERK activation after the first DMPP simulation (Fig. 3D) is almost correlated with the extent of activity-dependent potentiation of LDCV release, thereby suggesting the involvement of ERK signaling in ADP.
Amperometry is a powerful tool that detects single vesicle fusion in real time and recognizes the release of LDCV, which contain catecholamines. For these reasons, amperometric analysis can be a useful tool to investigate ADP of LDCV release, eliminating other effects, such as endocytosis and involvement of SSVs. Amperometric analysis revealed that the number of fusion events (peak number of amperometric currents), as well as amount of catecholamine release (integration of amperometric current), was increased in an activity-dependent manner (Fig. 1C).
Using an electron microscope, we show that stimulation of nAChRs increases the size of vesicle pools located near the plasma membrane (Fig. 4). Because LDCVs in the vicinity of the plasma membrane can be easily released upon stimulation, they might be referred to as the RRP in chromaffin cells (6, 35). LDCVs, located within 500 nm of the plasma membrane, can be classified as the RRP, which represents fusion-competent vesicles (35). Together with the analysis of amperometric currents (Fig. 1C), we suggest that the vesicle size of the RRP is increased by stimulation of nAChRs.
It has been reported that translocation of SSVs causes potentiation of exocytosis, and this might be the underlying mechanism for long-term potentiation (36). At presynaptic terminals, it has been suggested that calcium accumulation induced by repetitive stimulation plays a prominent role in activity-dependent augmentation of SSV release (10, 12). Our data suggest that ADP of LDCV release is also caused by vesicle translocation. However, repetitive stimulation did not result in calcium accumulation (Fig. 2), suggesting that the mechanism involved in ADP of LDCV release is different from that of SSV release.
In neuronal cells, it has been widely recognized that calcium influx via VGCCs can activate MAPK signaling via transactivation of tyrosine kinases and epithelial growth factor receptor (28). Here we report that MAPK signaling, which is potently activated by calcium influx, affects ADP of LDCV release via regulating vesicle translocation. Inhibition of MAPK signaling blocks the effect of ADP (Fig. 3) and suppresses the activity-induced change of LDCV distribution, which represents vesicle translocation (Fig. 4). It has been shown that disruption of cortical F-actin can evoke vesicle translocation via liberating LDCVs bound to F-actin in chromaffin cells (21, 37, 38). In agreement with the previous studies, we also observed that catecholamine release induced by first DMPP stimulation under treatment of cytochalasin B (a drug for F-actin disassembly) was greater, compared with control (Fig. 5B), suggesting that the increase in LDCV release by first DMPP stimulation under cytochalasin B treatment might be attributed to the increase of vesicle translocation caused by F-actin disassembly. Moreover, our data provide evidence for the involvement of F-actin disassembly in the potentiation of exocytosis. Stimulation of nAChRs induced F-actin disassembly (Fig. 5B), and ADP was inhibited by treatment with jasplakinolide, an inhibitor of F-actin disassembly (Fig. 5A). We also provide evidence that MAPK signaling is involved in regulation of F-actin disassembly, thereby modulating vesicle translocation. However, we cannot eliminate possibilities that MAPK signaling also can be involved in the processes of vesicle docking or fusion events.
MAPK signaling has been suggested to play an important role in the regulation of short-term synaptic plasticity via controlling F-actin disassembly at the synapses. The underlying mechanism for the regulation of synaptic plasticity has been proposed to be modulation of F-actin dynamics at the synapses by activation of MAPK, thereby regulating ADP of SSV release (8, 10, 39, 40). Calcium influx via VGCCs makes actin filament disassembled, which liberates SSVs tethered to F-actin, and finally induces SSV translocation to the plasma membrane (10, 40). In neurons, it has been well demonstrated that MAPK activation by calcium influx leads to phosphorylation of synapsin I, which causes F-actin disassembly and subsequent translocation of SSVs (38, 41). However, synapsin is not expressed in chromaffin cells (42, 43). Therefore, identifying target molecules, downstream of MAPK signaling as well as their involvement in F-actin disassembly requires further investigation.
Chromaffin cells are repetitively stimulated by acetylcholine released from the splanchnic nerve terminals (14). They are the main source for generating high level of catecholamine in the bloodstream when sympathetic nervous system is activated. Stress and fight-or-flight response are typical physiological actions of the sympathetic nervous system and primarily caused by the release of catecholamine from chromaffin cells (44). These catecholamines facilitate immediate physical reactions such as increasing heart rate and breathing, constricting blood vessels, and controlling lipolysis and glycogenolysis (45). Upon activation of the sympathetic nervous system, it is important to maintain a high level of catecholamine in the bloodstream for various physiological actions. Activity-dependent depression of transmitter release, prevalent at most synapses, is attributed to a depletion of the RRP of vesicles (8). Also, repetitive stimulation on secretory cells usually gives rise to desensitization of exocytosis. Interestingly, chromaffin cells show ADP of catecholamine release. We therefore suggest that ADP of LDCV release in chromaffin cells might be an important mechanism to establish a high level of catecholamine in the bloodstream upon repetitive activation of the sympathetic nervous system.
Acknowledgments
We are grateful to Ms. Soo-Jin Kim for electron microscopy instrumentation. We also thank Mr. Kyung-Chul Woo for riding and Byung-Soon Kang (Kyung-Buk Packers Co. Inc., Pohang, Republic of Korea) for providing bovine adrenal glands.
Footnotes
This work was supported by the Brain Neurobiology Research Program (M10412000023-02310) and Systems Biodynamics National Core Research Center sponsored by the Korean Ministry of Science and Technology and Brain Korea 21 Program of the Koran Ministry of Education.
The authors have nothing to declare.
First Published Online November 23, 2005
Abbreviations: ADP, Activity-dependent potentiation; [Ca2+]i, intracellular free calcium concentration; [Ca2+]o, extracellular calcium; DMPP, 1,1-dimethyl-4-phenylpiperazinium iodide; fura-2/AM, fura-2 pentaacetoxymethyl ester; LDCV, large dense-core vesicle; nAChR, nicotinic acetylcholine receptor; RRP, readily releasable pool; SSV, small synaptic vesicle; VGCC, voltage-gated calcium channel.
Accepted for publication November 17, 2005.
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