From the INSERM, U-338 Biologie de la Communication
Cellulaire, 5 rue Blaise Pascal, 67084 Strasbourg Cedex, France and
§ Pulmonary-Critical Care Medicine Branch, NHLBI, National
Institutes of Health, Bethesda, Maryland 20814
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ABSTRACT |
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The ADP-ribosylation factor (ARF) GTP-binding proteins have been implicated in a wide range of vesicle transport and fusion steps along the secretory pathway. In chromaffin cells, ARF6 is specifically associated with the membrane of secretory chromaffin granules. Since ARF6 is an established regulator of phospholipase D (PLD), we have examined the intracellular distribution of ARF6 and PLD activity in resting and stimulated chromaffin cells. We found that stimulation of intact chromaffin cells or direct elevation of cytosolic calcium in permeabilized cells triggered the rapid translocation of ARF6 from secretory granules to the plasma membrane and the concomitant activation of PLD in the plasma membrane. To probe the existence of an ARF6-dependent PLD in chromaffin cells, we measured the PLD activity in purified plasma membranes. PLD could be activated by a nonhydrolyzable analogue of GTP and by recombinant myristoylated ARF6 and inhibited by specific anti-ARF6 antibodies. Furthermore, a synthetic myristoylated peptide corresponding to the N-terminal domain of ARF6 inhibited both PLD activity and catecholamine secretion in calcium-stimulated chromaffin cells. The possibility that ARF6 participates in the exocytotic reaction by controlling a plasma membrane-bound PLD and thereby generating fusogenic lipids at the exocytotic sites is discussed.
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INTRODUCTION |
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ADP-ribosylation factors (ARFs)1 comprise a family of 20-kDa monomeric GTP-binding proteins that were discovered as one of several cofactors required in the cholera toxin-catalyzed ADP-ribosylation of the trimeric Gs proteins (1). Six mammalian family members have been identified which have been classified into three groups according to their size and sequence homology. ARF1, ARF2, and ARF3 form class I, ARF4 and ARF5 form class II, and ARF6 forms class III (1). Members of the ARF family are subjected to myristoylation at the N-terminal glycine residue, a lipid co-translational modification that appears essential for functional activity (2). ARFs are ubiquitous among eukaryotes with an amino acid sequence that is highly conserved across diverse species, suggesting a fundamental role in cellular physiology. Indeed, ARF proteins have been implicated in a wide range of vesicle transport and fusion steps along the secretory pathway (3-5). These include budding, transport, and fusion steps in the Golgi complex, in the endoplasmic reticulum and in the endocytotic and exocytotic pathways.
The recent discovery that some members of the ARF family are effective activators of phospholipase D (PLD) has raised the possibility that a novel signal transduction pathway may regulate intracellular membrane traffic (6, 7). PLD is an enzyme that catalyzes the hydrolysis of phosphatidylcholine to produce membrane-localized phosphatidic acid (PA) and soluble choline (8). In the presence of a primary alcohol, the enzyme can also catalyze a transphosphatidylation reaction that exchanges the polar headgroup of the phospholipid substrate with the given alcohol to form the corresponding phosphatidyl-alcohol (9). This unique and very useful property of PLD has been used to reveal PLD activation following agonist stimulation in many types of cells and tissues (10). Biochemical evidences suggest that multiple PLD isoenzymes with diverse mechanisms of activation occur in mammalian cells (11). An integral membrane-bound PLD that is highly specific for phosphatidylcholine as substrate and is activated by sodium oleate was recently purified (12). In addition, several forms of small G protein-dependent PLDs including ARF-sensitive (6, 7) and RhoA-sensitive (13) isoenzymes have been described. Phosphatidylinositol 4,5-bisphosphate (PIP2), another important activator of PLD, seems to be generally required for the small G protein-dependent PLDs (6, 7) but not for the oleate-dependent PLD (12). Protein kinase C appears also as a major regulator of PLD since phorbol esters are among the most effective stimuli of PLD reported in many cell types (11). To date, several mammalian PLDs have been cloned and sequenced (11, 14).
Studies in neutrophils (15), pancreatic B cells (16), and pheochromocytoma PC12 cells (17) have suggested a role for PLD in exocytosis. In chromaffin cells, however, the occurrence of an agonist-regulated PLD activity remains a controversial issue (18, 19). We recently described a secretory granule-associated ARF6 protein that may represent a key component of the exocytotic pathway in chromaffin cells (20). Since ARF6 is an established regulator of PLD (21), we examine here the PLD activity in resting and stimulated chromaffin cells. We found that stimulation of chromaffin cells triggered the rapid translocation of ARF6 from secretory granules to the plasma membrane and the concomitant activation of PLD in the plasma membrane. Both calcium-evoked PLD activation and calcium-induced catecholamine secretion could be inhibited by a synthetic peptide corresponding to the N-terminal domain of myristoylated ARF6. We propose that ARF6 may participate in the exocytotic reaction by controlling a plasma membrane-bound PLD and thereby contributing to the generation of fusogenic lipids at the exocytotic sites.
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EXPERIMENTAL PROCEDURES |
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Culture of Chromaffin Cells-- Chromaffin cells were isolated from fresh bovine adrenal glands and maintained in primary culture, essentially as described previously (22). Cells were usually cultured as monolayers either on 24 multiple 16-mm Costar plates (Cambridge, MA) at a density of 2.5 × 105 cells/well or on 100-mm Costar plates at a density of 5 × 106 cells/plate. To trigger exocytosis, chromaffin cells were washed twice with Locke's solution (140 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4, 11 mM glucose, 0.56 mM ascorbic acid, and 15 mM Hepes, pH 7.2) and then stimulated 10 min with Locke's solution containing either 10 µM nicotine or 59 mM K+ (made by decreasing NaCl isosmotically). Experiments were carried out at 37 °C on 3-7-day-old cultures.
Permeabilization with Streptolysin O (SLO) and Cell Stimulation-- Cultured chromaffin cells were washed four times with Locke's solution and twice with Ca2+-free Locke's solution (containing 1 mM EGTA). Permeabilization was performed with 15 units/ml SLO (Institut Pasteur, Paris, France) for 2 min in 200 µl/16-mm well or in 5 ml/100-mm plate Ca2+-free KG medium (150 mM K+-glutamate, 10 mM PIPES, pH 7.0, 5 mM nitrilotriacetic acid, 0.5 mM EGTA, 5 mM MgATP, 4.5 mM magnesium acetate, 0.2% bovine serum albumin). Cells were subsequently stimulated for 10 min in the presence of the compound to be tested in KG medium containing 20 µM free Ca2+ and 1 mM free Mg2+ (22).
[3H]Noradrenaline and Endogenous Catecholamine Release from Permeabilized Chromaffin Cells-- Catecholamine stores were labeled by incubating chromaffin cells with [3H]noradrenaline (14.68 Ci/mmol, NEN Life Science Products) for 60 min. Cells were then washed four times, permeabilized with SLO, and stimulated with 20 µM free Ca2+ as described above. [3H]Noradrenaline release after stimulation was determined by measuring the radioactivity present in the incubation medium and in cells after precipitation with 10% (w/v) trichloroacetic acid. Release of [3H]noradrenaline is expressed as a percentage of total radioactivity present in the cells before Ca2+-induced stimulation. Release experiments were performed in triplicate on at least two different cell preparations. In the figures that are representative of a typical experiment, data are given as the mean of triplicate determinations on the same cell preparation ± S.E. Endogenous catecholamine release was estimated by reverse phase high performance liquid chromatography with electrochemical detection as described previously (23).
Subcellular Fractionation of Cultured Chromaffin
Cells--
Cultured chromaffin cells were collected in 0.32 M sucrose, Tris 10 mM, pH 7.4, homogenized, and
then centrifuged at 800 × g for 15 min. After
centrifugation at 20,000 × g for 20 min, the pellet
containing the crude membrane fraction was resuspended in 0.32 M sucrose (10 mM Tris, pH 7.4), layered on a
continuous sucrose density gradient (1-2.2 M sucrose, 10 mM Tris, pH 7.4), and centrifuged for 90 min at
100,000 × g. Twelve 1-ml fractions were collected from
the top to the bottom and analyzed for PLD activity and protein content
by the Bradford procedure. The distribution of dopamine--hydroxylase
(D
H; chromaffin granule marker) and Na+/K+
ATPase (plasma membrane marker) in the fractions of the gradient was
estimated as described previously (24).
Assay for Phospholipase D Activity in Cultured Chromaffin
Cells--
Chromaffin cells were labeled with 1 µCi/ml
[9,10-3H]myristic acid for 24 h at 37 °C. Labeled
cells were then washed, stimulated with nicotine or 59 mM
K+ in the presence of 1% ethanol or permeabilized with
SLO, and then stimulated with 20 µM free Ca2+
in the presence of 1% ethanol. Cells were subsequently collected and
lipids extracted and separated with
CH3OH/CHCl3/0.1 N HCl (1:1:1,
v/v/v) according to the method of Bligh and Dyer (25). The lower
lipid-containing phase was collected, spiked with a mixture of standard
lipids containing L- phosphatidic acid
(1,2-diacyl-sn-glycero-3-phosphate) (PA) and
1-palmitoyl-2-oleoyl-sn-3-phosphoethanol (PEt), dried under
vacuum, and redissolved in 20 µl of
CHCl3/CH3OH (2:1, v/v). Lipids were separated
on one-dimensional TLC 0.25-mm oxalate-coated silica gel plates in a
solvent system composed of
CHCl3/CH3OH/CH3COOH/H2O (75:45:3:1, v/v/v/v). Labeled phospholipids were visualized with tritium imaging plates using the FUJIX BAS1000 Bio-Imaging Analyzer (Fuji, Tokyo, Japan). Standard lipids were stained with iodine vapor.
RF values were 0.31 for PA and 0.67 for PEt.
ARF6-dependent Phospholipase D Activity in Purified
Plasma Membranes--
ARF-dependent phosphatidylcholine
hydrolysis in purified plasma membranes was determined according to the
procedure previously described by Brown et al. (6) in the
presence of MgCl2 and 0.4 M NaCl to favor the
nucleotide exchange on ARF. The reaction was carried out in a final
volume of 60 µl. All assays contained 6.25 µg of plasma membrane
proteins in buffer A (50 mM Na-Hepes, pH 7.5, 3 mM EGTA, 80 mM KCl, 2 mM MgATP, 400 mM NaCl, 4.5 mM MgCl2, 3 mM CaCl2, and 1 mM dithiothreitol).
When indicated, 30 µM GTPS and 1 µM
recombinant myristoylated ARF6 were included. The membranes and the
above constituents in a volume of 34 µl were preincubated for 30 min
at 37 °C. The reaction was subsequently started by the addition of
24 µl of lipid substrate and 1% ethanol. The final concentration of
lipids in the assay were 135 µM PE, 12 µM
PIP2, 8 µM phosphatidylcholine (PC), and 1 µCi of L-
-dipalmitoylphosphatidylcholine (2-palmitoyl-9,10-3H-labeled). Lipids were
previously dried and sonicated in buffer A without MgCl2
and CaCl2. The incubation was performed for 30 min at
37 °C and stopped by the addition of 350 µl of 1 M
HCl, 5 mM EGTA, and 1 ml of
CHCl3/CH3OH/HCL (50:50:0.3, v/v/v). After vortexing and centrifugation (2000 × g for 5 min), 400 µl of the aqueous phase was extracted and analyzed as described
above.
Antibodies, Peptides, and Proteins--
Polyclonal anti-ARF6
antibodies were raised in rabbits against bacterially overexpressed
ARF6 protein. This antibody was a generous gift from Dr. J. B. Helms (Ruprecht-Karls-Universität, Heidelberg, Germany). The
rabbit polyclonal anti-dopamine -hydroxylase (EC 1.14.17.1)
antiserum was prepared in our laboratory, and its specificity has been
demonstrated (26).
Protein Determination, Electrophoresis, and Immunoblotting-- Protein concentration was routinely determined using the Bradford procedure with Bio-Rad dye reagent and bovine serum albumin as standard. One dimensional SDS-polyacrylamide gel electrophoresis was performed on 12% acrylamide gel in Tris-glycine buffer (24). Proteins were transferred to nitrocellulose sheets at a constant current of 120 mV for 1 h. Blots were developed with secondary antibodies coupled to horseradish peroxidase (Amersham, Les Ulis, France), and immunoreactive bands were detected with the ECL system (Amersham).
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RESULTS |
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Activation of Phospholipase D in Response to Chromaffin Cell Stimulation-- To measure PLD activity, we made use of the specific transphosphatidylation reaction in which the phospholipid headgroup is exchanged for ethanol, producing PEt at the expense of PA. Therefore, cultured chromaffin cells were labeled with [3H]myristic acid. Preliminary time-course experiments indicated that maximal incorporation of the radioactivity in total lipids occurred after 20 h of incubation. [3H]Myristic acid was predominantly incorporated into phosphatidylcholine (data not shown), which represents the major PLD substrate (10). Stimulation of [3H]myristic acid labeled chromaffin cells with nicotine or with a depolarizing concentration of potassium triggered the formation of [3H]PA and [3H]PEt in the presence of 1% ethanol (Table I). Both secretagogues elicited a similar increase in PA and PEt levels, suggesting that an agonist-stimulated phospholipase D activity was present in chromaffin cells.
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Subcellular Localization of the Phospholipase D Activity in
Stimulated Chromaffin Cells--
To determine the intracellular
localization of the PLD activity in stimulated chromaffin cells, we
analyzed the phospholipid content of subcellular fractions collected
from a sucrose density gradient layered with a crude membrane
preparation. Chromaffin cells labeled with
[3H]myristic acid were permeabilized with SLO and
subsequently exposed to 1% ethanol and GTPS in the presence or
absence of 20 µM free Ca2+. Cells were then
collected and processed for subcellular fractionation. As expected, the
[3H]PEt levels detected in the fractions obtained from
resting cells remained negligible (Fig.
2A). Interestingly,
radioactive [3H]PEt was essentially collected in
fractions 2 and 3 in gradients prepared from stimulated cells (Fig.
2A). These fractions contain plasma membranes as estimated
by the Na+/K+-ATPase activity. It is noteworthy
that fractions 11 and 12, enriched in chromaffin granules revealed by
the peak of D
H, contain very little radioactive
[3H]PEt in both resting and stimulated cells. In other
words, PLD activity in chromaffin cells is not associated with
secretory granules. However, a rise in cytosolic calcium triggers the
activation of a G protein-regulated PLD activity, essentially in the
plasma membrane of chromaffin cells.
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Translocation of ARF6 from Secretory Chromaffin Granules to the
Plasma Membrane in Stimulated Chromaffin Cells--
We previously
reported that ARF6 is specifically associated with secretory granule
membranes in chromaffin cells, most likely through an interaction with
the subunit of a granule-bound trimeric G protein (20). Stimulation
of chromaffin cells with nicotine or direct elevation of cytosolic
Ca2+ in permeabilized cells triggered the rapid
dissociation of ARF6 from secretory granules (20). To identify the
target membrane to which ARF6 translocates in stimulated cells, we
compared here the distribution of ARF6 in membrane fractions collected
from resting or nicotine-stimulated chromaffin cells. Fig.
2B illustrates an immunoreplica analysis, using an anti-ARF6
antibody, of fractions collected from a sucrose density gradient
layered with chromaffin cell crude membranes. The parallel distribution
of ARF6 immunoreactivity and D
H activity (Fig. 2, compare
A and B) confirms the specific association of
ARF6 with chromaffin granule membranes in resting cells, in agreement
with our previous observations (20). Stimulation of chromaffin cells
with 10 µM nicotine for 5 min completely modified the
distribution of ARF6 in the sucrose gradient since the highest immunosignal for ARF6 was then detected in fraction 3 containing the
plasma membrane (Fig. 2B). Similar results were obtained in gradients prepared from SLO-permeabilized cells stimulated by a rise in
cytosolic calcium (Fig. 4). These findings strongly suggest that ARF6
translocates from secretory granules to the plasma membrane upon
chromaffin cell stimulation.
Presence of an ARF6-regulated Phospholipase D Activity in Chromaffin Cell Plasma Membranes-- The recent identification of PLD as a possible effector of ARF proteins (6, 7) led us to investigate whether PLD might be activated by an ARF6-dependent pathway in chromaffin cell plasma membranes.
To probe the existence of an ARF6-dependent PLD in chromaffin cells, we first examined the effect of recombinant ARF6 on PLD activity associated with purified plasma membranes. Plasma membranes recovered from a sucrose density gradient were assayed for PLD activity using lipid vesicles labeled with [3H]dipalmitoyl phosphatidylcholine. As illustrated in Fig. 3, the presence of 30 µM GTP
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Correlation between Catecholamine Secretion and ARF6-Phospholipase D Activation in Chromaffin Cells-- Fig. 5 illustrates the time course and calcium dose-response curve for secretion and PLD activation in SLO-permeabilized chromaffin cells. Secretion was estimated by measuring the release of endogenous catecholamines, and PLD activity was detected by measuring the formation of labeled PEt and PA. We found a strong similarity between the calcium sensitivity for PLD activation and the calcium concentration required for the exocytotic reaction in permeabilized cells (Fig. 5A). Furthermore, time-course experiments revealed that catecholamine secretion was always accompanied by the formation of PEt (Fig. 5B), in agreement with the hypothesis that PLD activation represents an important event in the pathway of regulated exocytosis. We previously reported that translocation of ARF6 was maximal within 1 min of stimulation (20). By comparison, maximal PLD activation was observed after 3 min of stimulation (Fig. 5B), an observation that correlates well with the idea that PLD stimulation requires the translocation of ARF6 to the plasma membrane.
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DISCUSSION |
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Dramatic advances have recently been made in our understanding of
the protein machinery responsible for the formation, targeting, and
fusion of vesicles along the secretory pathway. Most of the emerging
models emphasize the convergence in protein molecules and mechanisms
underlying the multiple steps of intracellular vesicular transport (30,
31). Since ARF proteins belong to the molecules that have been
implicated as ubiquitous regulators in membrane traffic (3-5), we
recently investigated the possible function of ARF in calcium-regulated
exocytosis in chromaffin cells (20). We found that ARF6 is specifically
associated with the membrane of secretory chromaffin granules, most
likely through an interaction with subunits of a trimeric G
protein. Interestingly, nicotine-induced stimulation of intact cells or
direct elevation of cytosolic calcium in permeabilized cells triggered
the rapid dissociation of ARF6 from secretory granules (20). Although ARF proteins are generally believed to cycle on and off the membrane in
a manner that is tightly coupled to the binding and hydrolysis of GTP,
we could not detect ARF6 in the cytosol in subcellular fractionation
experiments (20) or among the cytosolic proteins released through the
pores created in the plasma membrane of SLO-permeabilized cells.2 Thus, we postulated
that ARF6 translocated from chromaffin granules to an unknown
membrane-bound compartment upon cell stimulation. Interestingly, the
subcellular distribution of human ARF proteins has recently been
examined in detail in Chinese hamster ovary cells, and ARF6 was the
only isoform that could not be detected in the cytosol (32). It is also
noteworthy that both wild-type and mutant forms of ARF6 were
exclusively localized in membrane compartments when overexpressed in
fibroblasts (33). Thus, ARF6 seems to behave quite distinctly from
other ARFs, at least regarding its membrane binding activity and
intracellular localization. Based on subcellular fractionation
techniques and immunological detection, we report here that agonist
stimulation triggers the translocation of ARF6 from secretory granules
to the plasma membrane in chromaffin cells. Moreover, we found a close
correlation between the presence of ARF6 in the plasma membrane and the
activation of a GTP-dependent PLD activity in the plasma
membrane, suggesting that PLD may be a possible effector for ARF6 in
the exocytotic pathway.
The participation of ARF in exocytosis in endocrine and neuroendocrine
cells has been previously postulated (34-36). To probe the importance
of ARF6 in agonist-stimulated PLD activity and exocytotic response in
chromaffin cells, we used here synthetic N-terminal ARF peptides
described to block ARF activities in various cellular processes (37,
38). We found that myrARF6-(2-13), a peptide corresponding in sequence
to the myristoylated N-terminal domain of ARF6, specifically inhibited
calcium-evoked catecholamine release and PLD activation in stimulated
chromaffin cells. By comparison, the non-myristoylated ARF6-(2-13)
peptide had little effect, an observation that may be related to the
presumed importance of the myristoyl group in the binding of ARF6 to
membranes (2, 33). Dose-response experiments indicated that
myrARF6-(2-13) was able to block almost completely the calcium-evoked
secretory response in permeabilized chromaffin cells (90%
inhibition). Thus, ARF6 activation of PLD may represent a key event in
the exocytotic pathway in neuroendocrine cells.
Interestingly, we found that AlF4,
which activates specifically heterotrimeric G proteins (29), inhibited
the calcium-induced PLD activity in permeabilized chromaffin cells.
This observation correlates well with our previous findings that
AlF4
can prevent the calcium-induced
uncoupling of ARF6-G
on the secretory granule membrane (20) and
strongly reduce the calcium-evoked exocytotic activity in stimulated
cells (22). The regulation of ARF activities in the Golgi complex (39)
and in the endocytotic pathway (40) by trimeric G proteins has already
been reported. In chromaffin cells, activation of the secretory
granule-associated Go inhibits the ATP-dependent priming
step of exocytosis (22, 23). This suggests that activated
Go blocks the exocytotic machinery when the
o subunit is dissociated from
. Our data support a model in which exocytosis requires the inactivation of the
granule-bound Go, leading to the reassociation of
o with
. G
interacting with
G
o is then unable to retain ARF6, which translocates to the plasma membrane and activates PLD. We recently identified another
putative effector of Go in the exocytotic pathway, namely the monomeric GTP-binding protein Rho, which seems to regulate the
peripheral actin network (41). Interestingly, several reports describe
a reciprocal regulatory relationship between actin reorganization and
PLD activity (42-44). Thus, an attractive speculation is that the
granule-bound Go plays a double control of the plasma
membrane-associated PLD in the exocytotic pathway: through
and
ARF6, which directly activate the enzyme and through
o,
and Rho, which may modulate PLD by a specific cytoskeletal
reorganization.
PLD hydrolyzes PC to generate PA and choline. PA seems to be important for the exocytotic reaction since primary alcohols, which divert the production of PA to phosphatidylalcohol, inhibited calcium-evoked catecholamine secretion in permeabilized cells. How does PA relate to our current understanding of the protein machinery responsible for regulated exocytosis? In principle, PA can be rapidly converted into diacylglycerol, and one of the functions of the PLD pathway might be, therefore, the provision of diacylglycerol with the consequent activation of protein kinase C. PA is also a known stimulator of phosphatidylinositol 4-phosphate 5-kinase (45), an enzyme that has been implicated in the priming of exocytosis in chromaffin and PC12 cells (46, 47). Perhaps the most exciting speculation relates, however, to the changes in the lipid bilayer properties created by the activation of PLD. The conversion of PC to PA in the plasma membrane replaces a nonfusogenic phospholipid with a fusogenic one (48), a potentially positive effect for the exocytotic reaction. The local elevation of PA may also favor interactions between membranes and annexins (49). Annexin II is a Ca2+-dependent phospholipid-binding protein that forms cross-links between secretory granules and plasma membranes in stimulated chromaffin cells (50). Furthermore, the translocation of annexin II from the cytosol to the subplasmalemmal region in stimulated cells seems to be an essential event for catecholamine secretion (51). In view of a recent report describing that PLD lowers the calcium concentration required for annexin-induced liposome aggregation by increasing the PA composition (52), it is tempting to postulate that the activation of PLD and the production of PA may facilitate annexin II-mediated membrane-membrane apposition in the early stages of the exocytotic pathway in neuroendocrine cells. Since the role played by the lipid bilayer is almost completely lacking in the recent models for calcium-regulated secretion, it will be quite interesting to prove or refute any of these possibilities.
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ACKNOWLEDGEMENTS |
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We gratefully acknowledge Dr. Joel Moss for the generous gift of the bacterial expression system for myrARF6 and Dr. J. Bernd Helms for kindly providing us with anti-ARF6 antibodies. We thank Danièle Thiersé for culturing chromaffin cells, Gérard Nullans for the synthesis of peptides, and Dr. Nancy Grant for revising the manuscript.
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FOOTNOTES |
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* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ To whom correspondence should be addressed. Tel.: 33-3-88-45-67-13; Fax: 33-3-88-60-08-06; E-mail: bader{at}neurochem.u-strasbg.fr.
1
The abbreviations used are: ARF,
ADP-ribosylation factor; PLD, phospholipase D; PA, phosphatidic acid;
PIP2, phosphatidylinositol 4,5-bisphosphate; GTPS,
guanosine 5
-3-O-(thio)triphosphate; SLO, streptolysin O;
PIPES, 1,4-piperazinediethanesulfonic acid; D
H,
dopamine-
-hydroxylase; PEt,
1-palmitoyl-2-oleoyl-sn-3-phosphoethanol; PC,
phosphatidylcholine; PMA, phorbol 12-myristate 13-acetate.
2 M.-C. Galas and M.-F. Bader, unpublished data.
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REFERENCES |
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