From the Consiglio Nazionale delle Ricerche Institute of Neuroscience, Cellular and Molecular Pharmacology, Department of Medical Pharmacology, University of Milano, 20129 Milano, Italy
Received for publication, September 16, 2002, and in revised form, October 22, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
ATP is released from astrocytes and is involved
in the propagation of calcium waves among them. Neuronal ATP secretion
is quantal and calcium-dependent, but it has been suggested
that ATP release from astrocytes may not be vesicular. Here we report that, besides the described basal ATP release facilitated by exposure to calcium-free medium, astrocytes release purine under conditions of
elevated calcium. The evoked release was not affected by the gap-junction blockers anandamide and flufenamic acid, thus excluding purine efflux through connexin hemichannels. Sucrose-gradient analysis
revealed that a fraction of ATP is stored in secretory granules, where
it is accumulated down an electrochemical proton gradient sensitive to
the v-ATPase inhibitor bafilomycin A1. ATP release
was partially sensitive to tetanus neurotoxin, whereas glutamate
release from the same intoxicated astrocytes was almost completely
impaired. Finally, the activation of metabotropic glutamate receptors,
which strongly evokes glutamate release, was only slightly effective in
promoting purine secretion. These data indicate that astrocytes
concentrate ATP in granules and may release it via a regulated
secretion pathway. They also suggest that ATP-storing vesicles may be
distinct from glutamate-containing vesicles, thus opening up the
possibility that their exocytosis is regulated differently.
Astrocytes propagate long-range calcium signals to neighboring
cells and affect the activity of neurons by evoking calcium transients
(1-6) and modulating neurotransmission (7, 8). It was first believed
that calcium waves are propagated by the diffusion of intracellular
messengers, such as Ca2+ and inositol
1,4,5-trisphosphate, across gap junctions (see Ref. 9, and Refs.
therein). It has more recently become clear that calcium
propagation may occur by means of an extracellular pathway because it also takes place among physically separated astrocytes (3,
10) or when gap-junction-mediated coupling is pharmacologically impaired (11, 12).
Several lines of evidence now suggest that ATP is the major
extracellular messenger for inter-astrocyte calcium-mediated
communication (10, 13, 14). First, ATP is released from astrocytes
during calcium wave propagation (10, 14). Second, the propagation can
be reduced or abolished by purinergic antagonists (10, 11, 13, 15, 16)
or ATP-degrading enzymes (10, 12, 13). Finally, ATP mediates
calcium-based intercellular communications between astrocytes and other
cell elements such as meningeal cells (17), Muller cells (18), and
microglia (19). These findings have highlighted the importance of ATP
in cross-talk among astrocytes and between them and other cell types in
the central nervous system.
The mechanisms of ATP release by glial cells has been actively
investigated (9, 13, 14, 20-22). A recent study has reported a
connexin hemichannel-mediated ATP release from astrocytes (21). Accordingly, in glioma cells, connexin expression potentiates both the
resting and the stimulated ATP release (13). However, the possibility
of a vesicular component of ATP release has never been investigated in
astrocytes. ATP is known to be packaged in vesicles, being co-stored
with acetylcholine in central and peripheral nerves, and with
noradrenaline in the vesicles of sympathetic nerve terminals and in the
related granules from chromaffin cells (23). Furthermore, evidence has
been recently reported of vesicular ATP release from endothelial cells
(24). The existence of typical secretory granules undergoing regulated
secretion in astrocytes has recently been defined (25), and proteins of
the synaptic vesicle fusion machinery have been identified in glial
cells (26-28).
Because these findings could be consistent with vesicular storage and
regulated purine secretion from astrocytes, we investigated the
mechanisms of ATP storage and release from primary cultures of
hippocampal astrocytes.
Materials--
Antibodies against rat
SgII1 were raised in rabbits,
purified by affinity chromatography, and characterized as previously
described (25). The monoclonal antibodies against GFAP came from
Roche Molecular Biochemicals; the polyclonal antibodies against
colony-stimulating factor-1 receptor from Santa Cruz
Biotechnology (Santa Cruz, CA); and the polyclonal antibodies against
synaptobrevin/VAMPII were from Synaptic System GmbH (Gottingen,
Germany). Ribophorin and complex 3 were kindly provided by Prof D. Borgese (Milan, Italy). The secondary antibodies conjugated to
fluorescein isothiocyanate, Texas Red, 10 nm gold particles, and
peroxidase were obtained from Jackson Immunoresearch Laboratories (West
Grove, PA); APV, CNQX, MCPG, t-ACPD, AMPA, and bafilomycin
A1 were from Tocris Neuramin (Bristol, UK); quinacrine
dihydrochloride, bradykinin, glutamate, ATP, PPADS, PMA,
anandamide, apyrase (grade II) and flufenamic acid, BAPTA/AM
were from Sigma. The ATP assay kit came from Molecular Probes Europe
(Leiden, NL) and the lactate dehydrogenase kit from Sigma (Milano, Italy).
Cell Cultures--
Hippocampal mixed-glia cultures from
embryonic rat pups (E18) were obtained using previously described
methods (25). Briefly, after dissection, the hippocampi were
dissociated by treatment with trypsin (0.25% for 10 min at 37 °C)
followed by fragmentation with a fire-polished Pasteur pipette. The
dissociated cells were plated onto glass coverslips at a density of
0.5 × 106 cells/ml, and the cultures were grown in
minimum essential medium (Invitrogen) supplemented with 20%
fetal bovine serum (Euroclone Ltd, UK) and glucose at a final
concentration of 5.5 g/l (glial medium). To obtain a pure astrocyte
monolayer, any microglia cells were harvested by shaking 3-week-old
cultures. The primary hippocampal neuron cultures were prepared from
E18 embryos as previously described (29).
Immunocytochemistry--
The cultures were fixed for 25 min at
room temperature with 4% paraformaldehyde in 0.12 M
phosphate buffer containing 0.12 M sucrose. The fixed cells
were detergent-permeabilized and labeled with primary antibodies
followed by fluorochrome-conjugated secondary antibodies. The
coverslips were mounted in 70% glycerol in phosphate buffer containing
1 mg/ml phenylendiamine. The images were acquired using a BioRad
MRC-1024 confocal microscope equipped with LaserSharp 3.2 software.
Electron microscopy was performed as previously described (25).
Quinacrine staining was performed by incubating living cultures for 30 min at 37 °C with Krebs-Ringer-Hepes (KRH: 125 mM NaCl,
5 mM KCl, 1.2 mM MgSO4, 1.2 mM KH2PO4, 2 mM
CaCl2, 6 mM glucose, 25 mM
Hepes/NaOH, pH 7.4) containing 5 × 10 Subcellular Fractionation and Immunoblotting--
After being
grown on Petri dishes until near confluence, the astrocytes were
scraped, washed, and resuspended 1:4 in homogenization buffer (10 mM Hepes-KOH, pH 7.4, 250 mM sucrose, 1 mM Mg acetate, 0.5 mM phenylmethylsulfonyl
fluoride, 2 µg/ml pepstatin, 10 µg/ml aprotinin). The cells were
homogenized using a cell cracker (European Molecular Biology
Laboratory, Heidelberg, Germany) and centrifuged at 1,000 × g for 10 min to prepare the post-nuclear supernatant. This supernatant was loaded onto a 0.4-1.8 M sucrose gradient and spun
in a 41 SW rotor (Beckman Instruments, Inc., Palo Alto, CA) at 25,000 rpm for 18 h. Fractions (1 ml) were collected and analyzed by
SDS-PAGE followed by Western blotting as previously described (25).
Briefly, after electrophoresis, the proteins were transferred to
nitrocellulose filters which, after being incubated in blocking
buffer (5% milk, 25 mM Tris-HCl, pH 7.5, 150 mM NaCl), were labeled with primary antibodies followed by the appropriate secondary antibodies conjugated to peroxidase diluted
in blocking buffer containing 0.1-0.3% Tween 20. After extensive
washing, the immunodecoration pattern was revealed using an enhanced
chemiluminescence system (SuperSignal from Pierce, Rockford, IL)
following the manufacturer' s protocol.
FURA-2 Videomicroscopy--
The cultured cells were
loaded for 60-90 min at 37 °C with 5 µM FURA-2
pentacetoxy-methylester in KRH, washed in the same solution, and
transferred to the recording chamber of an inverted microscope
(Axiovert 100; Zeiss, Oberkochen, Germany) equipped with a calcium
imaging unit. For the assays, a modified CAM-230 dual-wavelength
microfluorimeter (Jasco, Tokyo, Japan) was used as a light source. The
experiments were performed using an Axon Imaging Workbench 2.2 equipped
with a PCO SuperVGA SensiCam (Axon Instruments, Foster City, CA). The
ratio values in discrete areas of interest were calculated from
sequences of images to obtain temporal analyses. The images were
acquired at 1-3 340/380 ratios/s. The experiments were performed in a
static bath at room temperature (24-25 °C). The increases in
calcium were quantified by measuring the peak and/or area of the response.
Glutamate Measurements--
The biological assay for glutamate
detection was performed as previously described (29). Specifically,
monolayers of astrocyte cultures in 60-mm Petri dishes were kept in 1 ml of KRH in the absence and then in the presence of PMA or t-ACPD for
10-30 min at 37 °C. Neuronal cultures loaded with FURA-2 were then
exposed to the different aliquots of KRH. Immediately before
challenging, the aliquot collected from unstimulated astrocytes was
supplemented with the stimuli. To verify that the biological activity
of the conditioned medium was caused by accumulated glutamate, a subset of recordings were made in the presence of glutamate receptor antagonists, APV and CNQX (29, 30). Although embryonic hippocampal neurons lack t-ACPD receptors coupled to calcium
mobilization,2 the specific
antagonist MCPG was always added to the conditioned medium upon t-ACPD
stimulation. To test the tetanus neurotoxin (TeNT) sensitivity of
glutamate release, the same astrocyte monolayer was challenged with PMA
before and after 20-h incubation with the neurotoxin. Collected
aliquots were frozen and then tested on the same FURA-2-loaded neurons.
The endogenous glutamate concentration in the conditioned medium was
determined by HPLC analysis coupled with fluorimetric detection as
previously described (30).
ATP Measurements--
Off-site ATP bioassay aliquots of KRH (1 ml), conditioned as described above for glutamate detection, were split
into two parts before testing on FURA-2-loaded astrocytes. One part was pretreated with apyrase (30 units/ml) for 15 min before testing. Before
being exposed to ATP sensor cells, each aliquot was supplemented with a
mixture of glutamate antagonists (APV 100 µM, CNQX 20 µM, MCPG 1 mM) and the appropriate stimulus
when conditioned under control conditions. KRH conditioned under
mechanical stimulation was collected from astrocyte monolayers shaken
for 5 min on an orbital shaker (Stuart Scientific, UK). The same
aliquots were tested for lactate dehydrogenase activity following the
manufacturer's protocol.
Bioluminescence Assay--
ATP levels in the superfusates of
pure astrocyte monolayers were measured using a luciferin/luciferase
assay (Molecular Probes, Leiden, NL) and a luminometer (Lumat,
Berthold, LB9501) according to the manufacturer's instructions. Each
sample was run in duplicate. Most of the samples were assayed within
5-10 min of collection; the others were frozen for subsequent ATP
determination. ATP was detected on subcellular fractions by means of
the same assay of equal aliquots of sucrose fractions that were boiled
for 5 min before being frozen.
Biological Assays of ATP Release from Astrocytes--
To study the
mechanisms of ATP release from hippocampal astrocytes, both "on
line" (Fig. 1A) and
"off-site" (Fig. 1D) biological assays were performed.
The first method is based on the finding that microglia co-cultured
with astrocytes may act as ATP reporter cells by selectively responding
to the ATP released from adjacent astrocytes as
[Ca2+]i increases (19). In this
assay, FURA-2-loaded astrocyte-microglia co-cultures (Fig.
1A) were digitally imaged in the presence of glutamate
receptor antagonists (100 µM APV and 20 µM
CNQX) to exclude the possible contributions of released glutamate or
D-serine. A gentle touch of the astrocyte with a glass
pipette (a widely used stimulus for ATP secretion: 10, 12, 13) (Fig. 1,
A and B) generated an increase in
[Ca2+]i in the stimulated cell,
followed by a delayed [Ca2+]i
response in neighboring astrocytes and microglial cells. Despite the
efficient propagation of the calcium signal among astrocytes (19), the
microglia [Ca2+]i responses were
completely blocked or substantially inhibited when mechanically
stimulated in the presence of the nonselective purinergic antagonist
PPADS (50 µM) or the ATP-degrading enzyme apyrase (30 units/ml) (Fig. 1C). No significant changes in mean
astrocyte calcium responses were recorded in the presence of PPADS or
apyrase (percent changes in
The alternative bioassay for the study of ATP secretion was based on
off-site measurements of released purine. Superfusates, conditioned by
differently treated pure astrocyte monolayers, were added to
FURA-2-loaded astrocytes as ATP sensor cells in the presence of
glutamate receptor antagonists (Fig. 1D). Fig. 1E
shows the [Ca2+]i responses
induced by the superfusates collected under static bath conditions
(a) or during mechanical stimulation (b). The
[Ca2+]i responses were completely
prevented when the conditioned medium was treated with apyrase for
10-15 min (Fig. 1F), thus indicating that ATP was the
involved bioactive compound. Analysis of lactate dehydrogenase release
revealed no significant difference between the extracellular media
collected under static bath conditions or during mechanical stimulation
(lactate dehydrogenase activity: control, 4.87 ± 0.69 units/liter;
mechanically stimulated, 6.27 ± 1.2 units/liter, n = 4, p = 0.37; Triton X-100-treated, 247 ± 11.3, n = 4, p < 0.001), thus excluding ATP
leakage caused by shear damage. The lack of cell damage was also
confirmed by the exclusion of Trypan Blue from the mechanically
stimulated astrocytes (data not shown).
Regulated ATP Release from Astrocytes--
We used the two
bioassays to obtain insights into the mechanisms that control ATP
release from primary astrocytes on different kinds of stimulation. To
investigate the possible calcium dependence of ATP release, astrocytes
were mechanically stimulated after 45 min treatment with the
intracellular calcium chelator BAPTA/AM (10 µM). Purine
release was largely calcium-dependent, as the response in
ATP reporter cells was significantly attenuated when the medium was
collected from BAPTA-treated astrocytes (off-site bioassay, Fig.
1E, c). BAPTA treatment significantly reduced the calcium response to below baseline levels (Fig. 1F).
Furthermore, ATP release was significantly increased by treatment
with the potent secretagogues PMA (100 nM) (Fig.
1G) or with the glutamate receptor agonists AMPA (100 µM) and t-ACPD (100 µM) (Fig.
1H), which have been previously shown to stimulate glutamate
secretion when simultaneously applied to astrocytes (31). On the basis of a standard dose-response curve of
[Ca2+]i response amplitude to
different concentrations of exogenous ATP, the actual ATP concentration
in the collected medium was estimated to be 130-290 nM
under static bath and 550-700 nM after mechanical
stimulation or secretagogue treatment. Determination of the ATP in the
extracellular medium using the sensitive luciferin-luciferase bioluminescence assay revealed a 4.5 ± 0.4-fold (n = 3) increase in ATP release after PMA stimulation.
We then evaluated whether connexin hemichannels mediate ATP efflux from
primary astrocytes upon stimulation. To test this hypothesis directly,
we measured stimulated ATP release in the presence of the gap-junction
blockers anandamide, which effectively uncouples astrocytes (11) or
flufenamic acid, which has been recently used as connexin hemichannel
blocker (21, 32). As shown in figure
2A, no significant reduction
in [Ca2+]i response was observed
when the astrocytes were mechanically stimulated after 10-30 min
incubation with 100 µM anandamide (on-line bioassay).
Accordingly, [Ca2+]i responses in
ATP reporter cells to the medium conditioned by a PMA-stimulated
astrocyte monolayer were unaffected by the presence of the gap-junction
blocker (off-site bioassay) (Fig. 2B). To ensure that the
doses of anandamide used in this study were appropriate to block
gap-junction communication, FURA-2-loaded astrocytes were mechanically
stimulated in the presence of the purinergic antagonist PPADS and the
ATP-degrading enzyme apyrase with or without anandamide. When the
extracellular pathway was inhibited by the purinergic blockers,
anandamide completely prevented calcium signal propagation, thus
indicating an efficient block of the gap-junction-mediated
communication (wave propagation radius: PPADS, apyrase, without
anandamide: 145.9 ± 11 µm, n = 4; PPADS, apyrase,
with anandamide: 12.1 ± 6.4 µm, n = 5, p < 0.01). Furthermore, as blocking of gap-junctions
may not be indicative of inhibition of connexin hemichannels, we tested
whether anandamide blocks NAD+ influx down a concentration
gradient, which is known to occur through connexin hemichannels (33).
An 86 ± 13% reduction of NAD+ influx was caused by a
15-min preincubation with 100 µM anandamide. These data
indicate that anandamide, which is effective in blocking nucleotide
fluxes through connexin hemichannels, does not significantly impair ATP
release evoked by astrocyte stimulation. Similarly, no significant
reduction in ATP release, monitored as
[Ca2+]i response in microglial
cells (on-line bioassay), was observed when the astrocytes were
mechanically stimulated after 10-30 min incubation with 50 µM flufenamic acid (data not shown).
It has been reported in the literature that exposure of astrocytes to
calcium-free medium facilitates ATP release and that this treatment
promotes opening of connexin hemichannels (15, 21). Treatment of
hippocampal astrocytes with calcium-free medium significantly increased
basal ATP release, as detected by using the sensitive
luciferin-luciferase assay (196 ± 15% increase upon controls). ATP
release was significantly reduced (30 ± 1.8% reduction) in cultures
incubated with the gap-junction inhibitor anandamide.
Subcellular Distribution of ATP in Cultured Astrocytes--
Our
data indicated the existence of a calcium-dependent,
gap-junction blocker-insensitive release of ATP in primary cultures of
hippocampal astrocytes, suggesting a vesicular purine storage. In line
with this hypothesis, labeling of astrocytes with quinacrine fluorescence dye, which is known to stain high levels of ATP bound to
peptides in large granular vesicles (24, 34) revealed the existence of
an ATP-containing population of vesicular organelles, prominently
localized in the perinuclear region of the astrocytes (Fig.
3A). This vesicular staining
was reminiscent of the localization of secretogranin II (SgII), a well
established marker of dense-core vesicles (35) recently detected in
astrocytes (Ref. 25; Fig. 3B; also see Fig. 3D).
To test directly whether ATP is contained in secretory granules, the
astrocytes were analyzed by means of subcellular fractionation on
sucrose equilibrium gradients, and the fractions were probed for ATP
using the luciferin-luciferase bioluminescence assay. This analysis
revealed the presence of a purine peak that completely overlapped SgII
immunoreactivity, as well as a major peak in the lightest fractions
that possibly represents cytosolic ATP (Fig. 3C). These data
indicated that at least a portion of ATP is packaged in the secretory
granules of hippocampal astrocytes.
A number of the proteins of the synaptic vesicle exocytotic machinery,
including synaptobrevin/VAMPII (36), have been identified in astrocytes
(26-28), and there is evidence suggesting the existence of
synaptobrevin/VAMPII-positive vesicles in glial cells (37). The bulk of
synaptobrevin/VAMPII was recovered in the light sucrose gradient
subcellular fractions (fractions 9-13), and only a light trail of the
synaptic vesicle marker overlapped the SgII- and ATP-containing
fractions (fractions 14-18). A similar subcellular distribution of
synaptobrevin/VAMPII versus SgII has been previously reported in neuroendocrine cells, which contain small synaptic-like vesicles and secretory granules (38-42).
The coexistence of two distinct types of vesicles in astrocytes is
further supported by the immunofluorescence staining of cultures for
synaptobrevin/VAMPII and SgII. As shown in Fig. 3D, synaptobrevin/VAMPII immunoreactivity (red) did not
colocalize with the puncta of SgII staining (green)
representing individual secretory granules, but appeared to reflect the
distribution of a smaller vesicle population.
Bafilomycin A1 Strongly Impairs ATP Storage and Reduces
ATP Release--
Studies on chromaffin cells indicate that ATP uptake
in secretory granules requires an electrochemical proton gradient (43, 44), maintained by a v-ATPase that is selectively inhibited by
bafilomycin A1 (45, 46). To get insights into the
mechanisms of ATP storage, astrocytes were treated with bafilomycin
A1 (1-4 µM) for 60 min and then probed for
quinacrine staining. Virtually no fluorescent staining was detected
within treated cells, as shown in Fig. 4
(A-D). A fluorescent granular pattern was
recovered 30-60 min after washing (data not shown), thus suggesting a
reversible action of the drug (47).
Consistent with the possibility that bafilomycin A1 impairs
ATP storage in secretory organelles, treatment with the v-ATPase inhibitor significantly impaired the calcium-evoked release of the
purine, as indicated by the reduced
[Ca2+]i responses in adjacent
microglia cells (Fig. 4E) (on-line bioassay) (66.3 ± 4.05%
inhibition of microglial/astrocyte calcium response after bafilomycin
A1, n = 19, p < 0.01). No
significant changes in mean astrocyte calcium responses were recorded
in the presence of bafilomycin A1 (see also Ref. 48), thus
ruling out a possible interference of bafilomycin A1 with
the Ca2+ signal that is necessary to trigger ATP release
(percent changes in ATP Release Is Partially Sensitive to Tetanus Toxin--
The
possible existence of two classes of secretory organelles in astrocytes
raised the question as to whether ATP and glutamate (the other main
bioactive compound released by astrocytes) are stored in the same or
different organelles. To clarify this point, the sensitivity of ATP
release to TeNT was evaluated and compared with that of
glutamate in the same intoxicated cells. If glutamate and ATP are
stored in the same organelles, then the secretion of both messengers
should be equally sensitive to the toxin. TeNT (100 nM,
24 h incubation) produced a massive, but not complete, cleavage of
its molecular target synaptobrevin/VAMPII (Fig.
5, A and B).
To test the TeNT sensitivity of glutamate or ATP release, aliquots of
medium conditioned by the same astrocyte monolayer before or after TeNT
treatment were tested in parallel on neurons for glutamate detection
(29, 30) or on astrocytes for ATP detection (Fig. 5C). In
line with previous results (27, 31, 37, 48), TeNT treatment of the
astrocyte monolayers considerably impaired both t-ACPD- and PMA-evoked
glutamate release (Fig. 5D). On the contrary, only a partial
reduction of ATP release was observed in the same conditioned medium
after TeNT intoxication (Fig. 5F). A similar inhibition of
ATP release from stimulated astrocytes was estimated by means of the
on-line biological assay in astrocyte-microglia co-cultures after TeNT
intoxication (32.8 ± 3.8% inhibition of microglial/astrocyte calcium
response after TeNT treatment, n = 15, p < 0.05). No further inhibition of ATP release was
produced by flufenamic acid on TeNT-intoxicated astrocytes (32 ± 1.8%
inhibition in intoxicated astrocytes treated with flufenamic acid,
n = 34).
TeNT treatment did not lead to a general reduction in the physiological
integrity of astrocytes, which did not show any change in the baseline
levels of calcium or sodium, thus indicating the preservation of ionic
homeostasis (percent changes in the 340/380 fluorescence ratio after
TeNT: calcium dye (FURA-2), 103.6 ± 2.95%, n = 8, p < 0.05; sodium dye (SBFI), 102.5 ± 1.98%,
n = 6, p < 0.05, values normalized to
controls). Furthermore, responsiveness to exogenous AMPA was not
significantly altered (percent change in response after TeNT: 127 ± 8.1%, n = 6, p = 0.68, data normalized to controls), thus indicating that receptor trafficking was not significantly affected. The different sensitivity of glutamate and ATP
release to the action of TeNT rules against the accumulation of the two
messengers in the same intracellular exocytotic vesicles. Furthermore,
the partial sensitivity of ATP release to TeNT fits the main storage of
purine in secretory granules, which are known to be only partially
sensitive to clostridial neurotoxins (42).
Release of Glutamate and ATP Can Be Differently
Evoked--
Phorbol esters are potent exocytotic stimuli that enhance
regulated secretion from a variety of cell types including astrocytes. In line with its ability to stimulate the exocytosis of both synaptic vesicles (49) and secretory granules (25, 50), PMA significantly increased astrocyte ATP and glutamate release. Stimulations with PMA or
t-ACPD, without any other glutamate agonists, were similarly effective
in inducing the astrocyte release of glutamate (Fig. 6A), whereas t-ACPD was much
less efficient than PMA in triggering ATP secretion (Fig.
6B). These data were confirmed by the direct evaluation of
ATP and glutamate concentrations in the superfusates (glutamate, by
HPLC: 33.5 ± 5 pM (control), 450 ± 6.8 pM
(after t-ACPD treatment), 559 ± 7 pM (after PMA
treatment); ATP, by luciferin-luciferase assay: 6-8-fold increase
(after PMA stimulation), no increase (after t-ACPD stimulation)). It is
worth noting that t-ACPD did not significantly increase SgII secretion
(data not shown).
Although it is now widely established that ATP is an essential
component of long-range calcium signaling in the nervous system (23)
involved in the propagation of calcium waves (10, 14, 15, 18), the
mechanisms regulating its release from glial cells have not been
clarified yet. It has been suggested that ATP release from astrocytes
may not be vesicular (13, 15, 48, 51) on the basis of three main lines
of evidence: ATP release is tightly linked to connexin expression in C6
glioma cell lines; To investigate ATP storage and release mechanisms, we set up two
bioassays for ATP detection in the culture system. The first bioassay
was based on off-site measurements of ATP in superfusates collected
from differently stimulated astrocytes and represents a modification of
the method previously used by Guthrie et al. (10); the
second is a novel on-line biological assay based on the ability of
microglia co-cultured with astrocytes to respond to the ATP released
from adjacent astrocytes with
[Ca2+]i transients (19). The
absence of gap junctions between astrocytes and microglia (Ref. 52 and
this study) and the absence of calcium-permeable glutamate receptors on
microglial cells,3 allow the
evaluation of the [Ca2+]i
transients mediated by ATP, without considering either the glutamate-
or gap-junction-mediated components. Even more important, this method
also allows ATP detection at the astrocyte-releasing sites, thus
avoiding dilution and degradation. Although the incomplete abrogation
of the microglial calcium response by PPADS and apyrase could not
exclude the minor contribution of other mechanisms, such as the nitric
oxide-G-kinase signaling pathway (53), this assay is particularly well
suited to investigate the mechanisms involved in purine release from
glial cells.
The results of both assays indicated secretagogue- and
calcium-dependent ATP release from astrocytes. The possible ATP
efflux through connexin hemichannels does not account for the release triggered by PMA or mechanical stimulation. On the other hand, as
previously reported (15, 21), we confirm that connexin hemichannels
contribute to basal ATP release facilitated by exposure of the cultures
to calcium-free medium. These data suggest the coexistence, in primary
astrocytes, of mechanisms that mediate ATP secretion via connexin
hemichannels or vesicular organelles depending on environmental
conditions and on the activation of astrocytes.
To further test the possibility of vesicular storage, we used cell
fractionation analysis. Luciferin-luciferase assays of the ATP content
in subcellular fractions obtained using a sucrose equilibrium gradient
showed that a significant amount is concentrated in the secretory
granule-containing fractions. As previously demonstrated for chromaffin
and neurosecretory granules (43, 54), ATP is accumulated in these
organelles down an electrochemical proton gradient. The existence of
dense core vesicles capable of undergoing regulated release has been
previously reported in hippocampal astrocytes (25). The morphology and
density of the vesicles are similar to those of the secretory granules
found in neuroendocrine cells and, like them, contain the well known
SgII marker of the regulated secretory pathway. Hippocampal astrocytes
release SgII in response to secretagogues in a
calcium-dependent manner. The PMA protein kinase C
activator has been found to be particularly efficient in inducing
regulated SgII release from astrocytes (25). More modest, but still
significant, effects have been observed after treatment with bradykinin
(25), whereas the t-ACPD metabotropic glutamate receptor activator had
hardly any effect.4
Similarly, ATP release was efficiently induced by PMA, but not by
t-ACPD. Together with the results of the cell fractionation experiments, these findings clearly indicate that the dense core vesicles of hippocampal astrocytes co-store ATP with SgII. However, the
existence of different purine pools could not be excluded because ATP
was also detected in lighter fractions that are not immunoreactive to
ER or mitochondrial markers (ribophorin and complex
3),5 and whose identity has
still to be defined.
Having found that astrocytic secretory granules contain ATP, the
question arouse as to whether these organelles also store and release
glutamate, the other main bioactive compound released by astrocytes.
The ability of astrocytes to release glutamate via a
calcium-dependent, vesicular mechanism has been clearly established (31, 37, 48). Our results suggest that glutamate is stored
in vesicles other than the ATP- and SgII-containing large dense-core
granules. The existence of two types of astrocytic secretory organelles
was clearly shown by cell fractionation, immunocytochemistry and
functional assays. Besides the clear lack of colocalization of SgII-
and synaptobrevin/VAMPII-positive organelles revealed by
immunofluorescence, the two markers were clearly differently distributed in the sucrose gradient fractions. The immunoreactivity profile for the two proteins was identical to that obtained in sucrose
gradient fractions of PC12 cells (42) which are characterized by the
presence of typical secretory granules storing SgII, and synaptic-like
microvesicles containing the bulk of synaptobrevin/VAMPII. In line with
the small pool of synaptobrevin/VAMPII in the fractions enriched in
dense-core granules, it has been shown that calcium-evoked catecholamine secretion from PC12 cells is only about 35% inhibited by
TeNT despite the nearly complete cleavage of synaptobrevin/VAMPII (42).
This has been considered a typical feature of secretory granules and,
in the past, led to the hypothesis of the existence of other v-SNARES
(soluble NSF attachment protein receptors on the granule membrane)
(42). We took advantage of the different sensitivity of the two types
of vesicles to TeNT on the assumption that, if glutamate and ATP are
co-stored in the same organelles, the secretion of both messengers
should be equally reduced by TeNT. The evaluation of ATP and glutamate
release from the same intoxicated cultures revealed that the
TeNT-induced cleavage of synaptobrevin/VAMPII leads to the considerable
impairment of glutamate exocytosis but only a partial reduction in ATP
release. This finding is in line with previous reports indicating that
TeNT treatment almost completely abolishes the astrocyte secretion of
the low molecular weight messenger glutamate (31, 37, 55). It also supports the concept that glutamate is stored in a population of light
vesicles in astrocytes which, like neuronal synaptic vesicles, contain
the bulk of synaptobrevin/VAMPII. Furthermore, in addition to
indicating that ATP and glutamate are stored in different organelles,
the only partial sensitivity of ATP release to TeNT supports the
hypothesis that ATP is mainly stored in secretory granules, as already
indicated by subcellular fractionation analysis. The general inability
of clostridial toxins to completely impair purine release explains a
previous observation that calcium waves still propagate among
astrocytes in the presence of botulinum toxin (48). Furthermore, the
storage of ATP in large dense-core granules is also compatible with the
finding that The data reported here provide strong evidence for a vesicular storage
and release of ATP. These findings, together with the previously
reported connexin hemichannel-mediated ATP release, provide support for
the coexistence of different pathways that may be differentially
activated depending on the functional state of the astrocytes. The
possibility that astrocytes may privilege one or the other pathway in
different physiological or pathological conditions has intuitive appeal
in explaining the plasticity of astrocytes in responding to the
enviromental stimuli. Furthermore, the storage of ATP and glutamate in
distinct vesicular organelles opens up the interesting possibility that
their exocytosis is differently regulated. In line with this
hypothesis, we found that the activation of metabotropic glutamate
receptors efficiently induces glutamate exocytosis but not ATP release.
It is well known that in neurons the exocytosis of synaptic and large
dense-core vesicles is differentially regulated, with more potent
stimuli being required to induce large dense-core vesicle fusion (57, 58). A more widespread analysis of the signals that may differentially affect the release of the two main astrocytic extracellular messengers is essential to improve our understanding of the mechanisms of communication among astrocytes and between them and other cell types in
the nervous system.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
7
M quinacrine dihydrochloride. Quinacrine-fluorescent living
astrocytes were examined with a Zeiss microscope equipped with
epifluorescence and photographated using a TMAX 400 (Eastman Kodak
Co.).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
340/380 fluorescence ratio: PPADS,
74.5 ± 7.8, n = 5, p = 0.14;
apyrase, 115.68 ± 21.5, n = 6, p = 0.43, data normalized to controls). These data indicate that ATP is the
extracellular messenger responsible for microglial [Ca2+]i responses. Furthermore, a
significant delay in the residual response was observed in the presence
of the same blockers (a 514 ± 42% increase in the time to peak
response in the presence of PPADS, and 420 ± 35% in the presence of
apyrase, as compared with controls). Similar results were obtained when
the mixed cultures were stimulated with 1 µM bradykinin
(Fig. 1C), which selectively increases
[Ca2+]i in astrocytes (19).
View larger version (37K):
[in a new window]
Fig. 1.
Calcium-dependent ATP secretion.
A, schematic representation of the on-line biological assay
for ATP detection. Left panel, co-culture of astrocytes and
microglial cells double stained for the astrocytic marker GFAP
(green) and the microglial marker colony-stimulating
factor-1 receptor (red). Middle and right
panels, pseudocolor images of FURA-2-loaded mixed glial cultures
showing the propagation of a mechanically induced calcium wave from
astrocytes to a microglial cell. Middle panel, peak
[Ca2+]i response in the mechanical
stimulated astrocyte (indicated by a); right
panel: peak [Ca2+]i
rise in a microglial cell present in the field (indicated by
m). Inset, double labeling of the same field with
antibodies against GFAP (green) and colony-stimulating
factor receptor (red). B, temporal plot of
[Ca2+]i changes recorded from a
mechanically stimulated astrocyte (blue trace) and an
adjacent microglia (green trace). Note the delay
of the onset of the [Ca2+]i
response in the microglial cell. C, histograms showing the
mean peak ± S.E. of the
[Ca2+]i response in microglia
normalized to the peaks [Ca2+]i
increases in mechanically or bradykinin-stimulated astrocytes, under
control conditions or in the presence of PPADS or apyrase (mechanical
stimulation = 93.3 ± 9.3; n = 14; mechanical
stimulation + PPDAS: 16 ± 3.4; n = 8; mechanical
stimulation + apyrase: 31.25 ± 6; n = 13;
bradykinin = 90.6 ± 13.8; n = 12; bradykinin + PPADS 15.75 ± 4.46; n = 9). D,
schematic representation of the off-line biological assay for ATP
detection. Medium conditioned by a pure astrocyte monolayer
(left panel, staining for GFAP) induces a
[Ca2+]i response in FURA-2-loaded
astrocytes as ATP sensor cells: pseudocolor images taken before
(middle panel) and 30 s after (right
panel) the addition of the conditioned medium. E,
temporal plot of [Ca2+]i increases
recorded from two distinct FURA-2-loaded astrocytes after the addition
of exogenous ATP and medium conditioned by the same astrocyte monolayer
under static conditions (a), mechanical stimulation
(b), and mechanical stimulation after BAPTA treatment
(c). F, G, and H,
quantitative analysis of ATP levels in medium conditioned by a pure
astrocyte monolayer under control conditions and upon mechanical
stimulation (F), or treatment with PMA (G) or
t-ACPD and AMPA (H). The values represent percent changes
(± S.E.) normalized to the
[Ca2+]i responses induced by the
addition of exogenous 1 µM ATP. Note that, in all three
cases, the conditioned medium pretreated with apyrase (30 units/ml)
failed to evoke a [Ca2+]i
response. F, percent changes in response: 27 ± 2.6 under static conditions, 55.99 ± 2.7 under mechanical
stimulation, 19 ± 1.8 after BAPTA treatment; n = 28 p < 0.01. G, percent changes in
response: 18 ± 2.5 under control conditions, 62 ± 4 after
PMA treatment. H, percent changes in response: 18.2 ± 3 under control conditions; 48 ± 2 after t-ACPD + AMPA.
N.D., not detectable.
View larger version (13K):
[in a new window]
Fig. 2.
The evoked release of ATP from astrocytes is
gap-junction-independent. A, quantitative analysis of
mechanically evoked ATP release from astrocytes in the presence or
absence of anandamide, based on the on-line ATP bioassay. Histograms
show mean values ± S.E. of
[Ca2+]i responses in microglia
normalized to [Ca2+]i increases in
mechanically stimulated astrocytes, with or without the gap-junction
blocker anandamide (control = 93.3 ± 9.3; n = 14; anandamide-treated = 82.35 ± 6.3; n = 14; p > 0.1. B, quantitative analysis of
ATP release upon PMA stimulation with and without anandamide, based on
the off-line ATP bioassay. Values represent percent changes (±S.E.)
normalized to the [Ca2+]i
responses induced by 1 µM exogenous ATP (control
73.1 ± 7; n = 9; anandamide-treated = 89.6 ±12 n = 9; p > 0.1.
View larger version (29K):
[in a new window]
Fig. 3.
Subcellular distribution of ATP in
astrocytes. A, quinacrine staining in living astrocytes.
Note the presence of dot-like structures dispersed in the cytoplasm,
which suggest dye accumulation in vesicular organelles. Bar,
40.65 µm. B, electron micrograph showing dense-core
vesicles immunolabeled for SgII (10-nm gold particles). Note the
presence of unlabeled smaller vesicles in the ultrathin section.
Bar, 0.16 µm. C, plot of ATP content and
profile of SgII and synaptobrevin/VAMPII distribution in the sucrose
gradient fractions. Note the presence of an ATP peak completely
overlapping the SgII-immunoreactive fractions. An example of the
distribution of SgII and synaptobrevin/VAMPII in the gradient fractions
is shown below the graphs. D,
double-immunofluorescence staining for SgII (green) and
synaptobrevin/VAMPII (red). The merged image is shown in the
right panel. Bar, 54.2 µm.
View larger version (66K):
[in a new window]
Fig. 4.
Effect of bafilomycin on ATP storage and
release. A, stores of ATP distributed in vesicular
organelles visualized by quinacrine staining in living astrocytes.
C, after treatment with bafilomycin A1 virtually
fluorescent granular staining was detected. B and
D, differential interference contrast images of the
same fields as A and C, respectively.
E, quantitative analysis of mechanically evoked ATP release
from astrocytes in the presence or absence of bafilomycin
A1, based on the on-line ATP bioassay. Histograms show mean
values ± S.E. of [Ca2+]i
responses in microglia normalized to
[Ca2+]i increases in mechanically
stimulated astrocytes in bafilomycin A1-treated cultures
(33.7 ± 2.69; n = 18; p < 0.01).
Bar, 42.3 µm.
340/380 fluorescence ratio: 112.8 ± 12.7, n = 20, p = 0.55 data normalized to
controls). Moreover, microglial
[Ca2+]i responses to exogenous ATP
were not affected by bafilomycin A1 treatment.
View larger version (36K):
[in a new window]
Fig. 5.
TeNT differently affects ATP and glutamate
release from astrocytes. A, Western blot analysis of control
and TeNT-intoxicated cultured rat astrocytes shows that
synaptobrevin/VAMPII is massively cleaved by the clostridial toxin.
-tubulin staining is shown as a standard. B,
quantitative analysis of synaptobrevin/VAMPII content in control and
intoxicated cultures. Synaptobrevin/VAMPII immunoreactivity is
normalized to
-tubulin staining. C, schematic
representation of glutamate and ATP measurements in medium conditioned
by the same astrocyte monolayer. Aliquots of conditioned medium
collected before and after TeNT intoxication were split into two parts
and tested in parallel on glutamate reporter neurons and ATP reporter
cells. D, quantitative analysis of the glutamate released by
t-ACPD- or PMA-treated astrocyte monolayers before and after 24-h TeNT
intoxication (t-ACPD: 21 ± 2.6% residual
[Ca2+]i response after TeNT,
n = 9; PMA: 23 ± 2.2% residual
[Ca2+]i response after TeNT,
n = 11). E and F, quantitative
analysis of TeNT sensitivity of glutamate (E, 24 ± 3.5 residual [Ca2+]i response after
TeNT, n = 24) and ATP release (F, 65 ± 3% residual [Ca2+]i response
after TeNT, n = 17) from the same intoxicated astrocyte
culture. The histograms in D, E, and F
show percent changes (+S.E.) normalized to the
[Ca2+]i responses induced by
medium conditioned by astrocyte monolayers before TeNT intoxication.
N.D., not detectable.
View larger version (14K):
[in a new window]
Fig. 6.
Glutamate and ATP release from astrocytes can
be differently evoked by t-ACPD. Quantitative analysis of
glutamate (A) and ATP (B) levels in the
extracellular medium conditioned by astrocytes upon PMA or t-ACPD
stimulation. Histograms show percent changes (+S.E.) normalized to the
[Ca2+]i responses induced by the
conditioned medium upon PMA treatment. (t-ACPD/PMA calcium response in
glutamate reporter neurons: 83.4 ± 12 p > 0.1, n = 8; t-ACPD/PMA calcium response in ATP reporter
astrocytes: 26 ± 5.5 p < 0.01, n = 13).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-latrotoxin, a potent exocytotic stimulus, does
not evoke ATP release (15); and calcium waves propagate among
astrocytes in the presence of botulinum toxin (48), a blocker of
vesicle fusion with the plasma membrane. In this study, we report an
evoked, calcium-dependent ATP release from the secretory
granules of primary astrocytes.
-latrotoxin, a potent exocytotic stimulus purified from
spider venom, does not evoke ATP release (15). It has been found that
the stimulation of motor nerve terminals with
-latrotoxin potently
induces the release of acetylcholine-containing vesicles, but is almost
completely ineffective in causing the exocytosis of peptide-containing
large dense-core vesicles (56).
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Prof. E. Fedele (University of Genova) for HPLC glutamate detection, Dr. S. Bruzzone and Prof. A. De Flora (University of Genova) for NAD+ flux measurements, Prof. C. Montecucco (University of Padova) for the gift of tetanus neurotoxin, and U. Schenk for her help in some experiments. We thank Prof. F. Clementi (University of Milano) for helpful discussions and comments.
![]() |
FOOTNOTES |
---|
* This work was supported by grants from the European Commission (QLGR3-CT-2000-01343, Coordinator Eric Scarfone), MURST-PRIN 2000; from Telethon -1042-, from HFSPO RGY0027/2001-B and Azienda Spaziale Italiana (I/R/149/00) (to M. M.) and from the Consiglio Nazionale delle Ricerche (Target Project on Biotechnology) (to P. R.).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.
Present address: Department of Molecular Cell Biology and
Genetics, Max Planck Institute 01307, Dresden, Germany.
§ To whom correspondence should be addressed: CNR Institute of Neuroscience, Cellular, and Molecular Pharmacology, Dept. of Medical Pharmacology, University of Milano, Via Vanvitelli 32, 20129 Milano, Italy. Tel.: 39-02-50317099; Fax: 39-02-7490574; E-mail: C.Verderio@csfic.mi.cnr.it.
Published, JBC Papers in Press, October 31, 2002, DOI 10.1074/jbc.M209454200
2 C. Verderio and M. Matteoli, unpublished data.
3 C. Verderio and M. Matteoli, unpublished data.
4 F. Calegari and P. Rosa, unpublished data.
5 S. Coco, P. Rosa, M. Matteoli, and C. Verderio, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
SgII, secretogranin
II;
PMA, phorbol 12-myristate 13-acetate;
KRH, Krebs-Ringer-Hepes;
APV, 2-amino-5-phosphonovaleric acid;
CNQX, 6-cyano-7-nitroquinoxaline-2,3-dione;
MCPG, -methyl-4-carboxyphenilglycine;
GFAP, glial fibrillar acidic
protein;
t-ACPD, 1-aminocyclopentane-trans-1,3-olicarboxylic acid;
AMPA,
-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid;
PPADS, pyridoxalphosphate-6-azophenyl-2,4-disulfonic acid;
BAPTA/AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N,N-tetraacetic
acid tetrakis (acetoxymethyl ester).
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Nedergaard, M. (1994) Science 263, 1768-1771[Medline] [Order article via Infotrieve] |
2. | Parpura, V., Basarsky, T. A., Liu, F., Jeftinija, K., Jeftinija, S., and Haydon, P. G. (1994) Nature 369, 744-747[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Hassinger, T. D.,
Guthrie, P. B.,
Atkinson, P. B.,
Bennett, M. V. L.,
and Kater, S. B.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
13268-13273 |
4. |
Pasti, L.,
Volterra, A.,
Pozzan, T.,
and Carmignoto, G.
(1997)
J. Neurosci.
17,
7817-7830 |
5. |
Newman, E. A.,
and Zahs, K. R.
(1998)
J. Neurosci.
18,
4022-4028 |
6. |
Parpura, V.,
and Haydon, P. G.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
8629-8634 |
7. | Kang, J., Jiang, L., Goldman, S. A., and Nedergaard, M. (1998) Nat. Neurosci. 1, 683-692[CrossRef][Medline] [Order article via Infotrieve] |
8. | Robitaille, R. (1998) Neuron 21, 847-855[Medline] [Order article via Infotrieve] |
9. | Haydon, P. G. (2001) Nat. Neurosci. 2, 185-193[CrossRef] |
10. |
Guthrie, P. B.,
Knappenberg, J.,
Segal, M.,
Bennett, M. V. L.,
Charles, A. C.,
and Kater, S. B.
(1999)
J. Neurosci.
19,
520-528 |
11. |
Guan, X.,
Cravatt, B. F.,
Ehring, G. R.,
Hall, J. E.,
Boger, D. L.,
Lerner, R. A.,
and Gilula, N.
(1997)
J. Cell Biol.
139,
1785-1792 |
12. |
John, G. R.,
Scemes, E.,
Suadicani, S. O.,
Liu, J. S. H.,
Charles, P. C.,
Lee, S. C.,
Spray, D. C.,
and Brosnan, C. F.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
11613-11618 |
13. |
Cotrina, M. L.,
Lin, J. H.,
Lopez-Garcia, J. C.,
Naus, C. C. G.,
and Nedergaard, M.
(2000)
J. Neurosci.
20,
2835-2844 |
14. | Wang, Z., Haydon, P. G., and Yeung, E. S. (2000) Anal. Chem. 72, 2001-2007[CrossRef][Medline] [Order article via Infotrieve] |
15. |
Cotrina, M. L.,
Lin, J. H.,
Alves-Rodrigues, A.,
Liu, S., Li, J.,
Azmi-Ghadimi, H.,
Naus, C. C.,
and Nerdergard, M.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
15735-15740 |
16. |
Fam, S. R.,
Gallagher, C. J.,
and Salter, M. W.
(2000)
J. Neurosci.
20,
2800-2808 |
17. | Grafstein, B., Liu, S., Cotrina, M. L., Goldman, S. A., and Nedergaard, M. (2000) Ann. Neurol. 47, 18-25[CrossRef][Medline] [Order article via Infotrieve] |
18. |
Newman, E. A.
(2001)
J. Neurosci.
21,
2215-2223 |
19. |
Verderio, C.,
and Matteoli, M.
(2001)
J. Immunol.
166,
6383-6391 |
20. | Queiroz, G., Gebicke-Haerter, P. J., Schobert, A., Starke, K., and Von Kugelgen, I. (1997) Neurosci. 78, 1203-1208[CrossRef] |
21. |
Stout, C. E.,
Costantin, J. L.,
Naus, C. C.,
and Charles, A. C.
(2002)
J. Biol. Chem.
277,
10482-10488 |
22. |
Arcuino, G.,
Lin, J. H. C.,
Takano, T.,
Liu, C.,
Jiang, L.,
Gao, Q.,
Kang, J.,
and Nedergaard, M.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
9840-9845 |
23. | Zimmermann, H. (1994) Trends Neurosci. 17, 420-426[CrossRef][Medline] [Order article via Infotrieve] |
24. | Bodin, P., and Burnstock, G. (2001) J. Cardiovasc. Pharmacol. 38, 900-908[CrossRef][Medline] [Order article via Infotrieve] |
25. |
Calegari, F.,
Coco, S.,
Taverna, E.,
Bassetti, M.,
Verderio, C.,
Corradi, N.,
Matteoli, M.,
and Rosa, P.
(1999)
J. Biol. Chem.
274,
22539-22547 |
26. | Parpura, V., Fang, Y., Basarsky, T., Jahn, R., and Haydon, P. G. (1995) FEBS Lett. 377, 489-492[CrossRef][Medline] [Order article via Infotrieve] |
27. | Jeftinija, S. D., Jeftinija, K. V., and Stefanovic, G. (1997) Brain Res. 750, 41-47[CrossRef][Medline] [Order article via Infotrieve] |
28. | Maienschein, M., Marxen, M., Volknandt, W., and Zimmermann, H. (1999) Glia 26, 233-244[CrossRef][Medline] [Order article via Infotrieve] |
29. | Verderio, C., Coco, S., Fumagalli, G., and Matteoli, M. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 6449-6453[Abstract] |
30. | Verderio, C., Bruzzone, S., Zocchi, E., Fedele, E., Schenk, U., De, Flora, A., and Matteoli, M. (2001) J. Neurochem. 78, 646-657[CrossRef][Medline] [Order article via Infotrieve] |
31. | Bezzi, P., Carmignoto, G., Pasti, L., Vesce, S., Rossi, D., Rizzini, B. L., Pozzan, T., and Volterra, A. (1998) Nature 391, 281-285[CrossRef][Medline] [Order article via Infotrieve] |
32. |
Zhang, Y.,
McBride, D. W., Jr.,
and Hamill, O. P.
(1998)
J. Physiol. (Lond.)
508,
763-776 |
33. |
Bruzzone, S,
Guida, L.,
Zocchi, E.,
Franco, L.,
and De Flora, A.
(2001)
FASEB J.
15,
10-12 |
34. | Belai, A., and Burnstock, G. (2000) Neuroreport 11, 5-8[Medline] [Order article via Infotrieve] |
35. | Rosa, P., Hille, A., Lee, R. W., Zanini, A., De, Camilli, P., and Huttner, W. B. (1985) J. Cell Biol. 101, 1999-2011[Abstract] |
36. | Schiavo, G., Benfenati, F., Poulain, B., Rossetto, O., Polverino de Laureto, P., DasGupta, B. R., and Montecucco, C. (1992) Nature 359, 832-835[CrossRef][Medline] [Order article via Infotrieve] |
37. |
Pasti, L.,
Zonta, M.,
Pozzan, T.,
Vicini, S.,
and Carmignoto, G.
(2001)
J. Neurosci.
21,
477-484 |
38. | Navone, F., Di, Gioia, G., Jahn, R., Browning, P., and De Camilli, P. (1986) J. Cell Biol. 103, 2511-2527[Abstract] |
39. | Wiedenmann, B., Rehm, H., Knierim, M., and Becker, C. M. (1988) FEBS Lett. 240, 71-77[CrossRef][Medline] [Order article via Infotrieve] |
40. | Cameron, P. L., Sudhof, T. C., Jahn, R., and De Camilli, P. (1991) J. Cell Biol. 115, 151-164[Abstract] |
41. | Linstedt, A. D., and Kelly, R. B. (1991) Neuron 7, 309-317[Medline] [Order article via Infotrieve] |
42. | Chilcote, T. J., Galli, T., Mundigl, O., Edelmann, L., McPherson, P. S., Takei, K., and De Camilli, P. (1995) J. Cell Biol. 129, 219-231[Abstract] |
43. | Aberer, W., Kostron, H., Huber, E., and Winkler, H. (1978) Biochem. J. 172, 353-360[Medline] [Order article via Infotrieve] |
44. |
Gualix, J.,
Abal, M.,
Pintor, J.,
Garcia-Carmona, F.,
and Miras-Portugal, M. T.
(1996)
J. Biol. Chem.
271,
1957-1965 |
45. | Bowman, E. J., Siebers, A., and Altendorf, K. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 7972-7976[Abstract] |
46. | Hanada, H., Moriyama, Y., Maeda, M., and Futai, M. (1990) Biochem. Biophys. Res. Commun. 170, 873-878[Medline] [Order article via Infotrieve] |
47. |
Yoshimori, T.,
Yakamoto, A.,
Moryama, Y.,
Futai, M.,
and Tashiro, Y.
(1991)
J. Biol. Chem.
266,
17707-17712 |
48. |
Araque, A., Li, N.,
Doyle, R. T.,
and Haydon, P. G.
(2000)
J. Neurosci.
20,
666-673 |
49. |
Waters, J.,
and Smith, S. J.
(2000)
J. Neurosci.
20,
7863-7870 |
50. | Gillis, K. D., Mossner, R., and Neher, E. (1996) Neuron 16, 1209-1220[Medline] [Order article via Infotrieve] |
51. | Fields, R. D., and Stevens, B. (2000) Trends Neurosci. 23, 625-633[CrossRef][Medline] [Order article via Infotrieve] |
52. | Venance, L., Cordier, J., Monge, M., Zalc, B., Glowinski, J., and Giaumie, C. (1995) Eur. J. Neurosci. 7, 451-461[Medline] [Order article via Infotrieve] |
53. |
Willmott, N. J.,
Wong, K.,
and Strong, A. J.
(2000)
J. Neurosci.
20,
1767-1779 |
54. |
Moriyama, Y.,
and Futai, M.
(1990)
J. Biol. Chem.
265,
9165-9169 |
55. | Bezzi, P., Domercq, M., Brambilla, L., Galli, R., Schols, D., De, Clercq, E., Vescovi, A., Bagetta, G., Kollias, G., Meldolesi, J., and Volterra, A. (2001) Nat. Neurosci. 4, 676-678[CrossRef][Medline] [Order article via Infotrieve] |
56. | Matteoli, M., Haiman, C., Torri-Tarelli, F., Polak, J. M., Ceccarelli, B., and De Camilli, P. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 7366-7370[Abstract] |
57. | Lundberg, J. M., and Hokfelt, T. (1986) Prog. Brain Res. 68, 241-262[Medline] [Order article via Infotrieve] |
58. | Nicholls, D. G. (1998) Prog. Brain Res. 116, 15-22[Medline] [Order article via Infotrieve] |