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The Journal of Neuroscience, June 15, 2001, 21(12):4195-4206
Synapsin Controls Both Reserve and Releasable Synaptic Vesicle
Pools during Neuronal Activity and Short-Term Plasticity in
Aplysia
Yann
Humeau1,
Frédéric
Doussau1,
Francesco
Vitiello1,
Paul
Greengard2,
Fabio
Benfenati2, 3, and
Bernard
Poulain1
1 Neurotransmission et Sécrétion
Neuroendocrine, Centre National de la Recherche Scientifique, IFR-37
des Neurosciences, F-67084 Strasbourg Cédex, France,
2 Laboratory of Molecular and Cellular Neuroscience, The
Rockefeller University, New York, New York 10021-6399, and
3 Department of Experimental Medicine, Section of Human
Physiology, University of Genova, 16132 Genova, Italy
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ABSTRACT |
Neurotransmitter release is a highly efficient secretory process
exhibiting resistance to fatigue and plasticity attributable to the
existence of distinct pools of synaptic vesicles (SVs), namely a
readily releasable pool and a reserve pool from which vesicles can be
recruited after activity. Synaptic vesicles in the reserve pool are
thought to be reversibly tethered to the actin-based cytoskeleton by
the synapsins, a family of synaptic vesicle-associated phosphoproteins
that have been shown to play a role in the formation, maintenance, and
regulation of the reserve pool of synaptic vesicles and to operate
during the post-docking step of the release process. In this paper, we
have investigated the physiological effects of manipulating synapsin
levels in identified cholinergic synapses of Aplysia
californica. When endogenous synapsin was neutralized by the
injection of specific anti-synapsin antibodies, the amount of
neurotransmitter released per impulse was unaffected, but marked
changes in the secretory response to high-frequency stimulation were
observed, including the disappearance of post-tetanic potentiation
(PTP) that was substituted by post-tetanic depression (PTD), and
increased rate and extent of synaptic depression. Opposite changes on
post-tetanic potentiation were observed when synapsin levels were
increased by injecting exogenous synapsin I. Our data demonstrate that
the presence of synapsin-dependent reserve vesicles allows the nerve
terminal to release neurotransmitter at rates exceeding the synaptic
vesicle recycling capacity and to dynamically change the efficiency of
release in response to conditioning stimuli (e.g., post-tetanic
potentiation). Moreover, synapsin-dependent regulation of the fusion
competence of synaptic vesicles appears to be crucial for sustaining
neurotransmitter release during short periods at rates faster than the
replenishment kinetics and maintaining synchronization of quanta in
evoked release.
Key words:
Aplysia; synapse; exocytosis; neurotransmitter
release; short-term plasticity; synaptic depression; post-tetanic
potentiation
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INTRODUCTION |
Nerve terminals have the unique
property of sustaining vesicular release of neurotransmitter at high
rates, although only a small fraction of synaptic vesicles (SVs) is
available for immediate release. In addition, nerve terminals are able
to transiently modify synaptic efficacy to give rise to various forms
of short-term plasticity. These are thought to result from changes in
recruitment of reserve vesicles or changes in the trafficking of
vesicles between a releasable pool of cycling SVs and a reserve pool of SVs (Zucker, 1996 , 1999; Brodin et al., 1997; Neher, 1998). The readily releasable pool is thought to correspond to the number of SVs
docked morphologically to the presynaptic membrane that can be released
(Rosenmund and Stevens, 1996; Dobrunz and Stevens, 1997). Morphological
data have demonstrated that SVs in the reserve pool are organized in
clusters in which SVs are reversibly linked to the actin-based
cytoskeleton, possibly by a family of abundant SV-associated
phosphoproteins, the synapsins (Landis et al., 1988; Bernstein and
Bamburg, 1989; Hirokawa et al., 1989; Benfenati et al., 1992a, 1993;
Ceccaldi et al., 1995; Pieribone et al., 1995; Kuromi and Kidokoro,
1998).
Synapsins are highly conserved, SV-associated proteins that are
implicated in the formation of synaptic terminals and the regulation of
neurotransmitter release. Although the vertebrate synapsin family
comprises the products of at least three synapsin genes (synapsin I,
II, and III) (Südhof et al., 1989; De Camilli et al., 1990;
Hosaka and Südhof, 1998; Kao et al., 1998), only a single
synapsin gene, which cannot yet be assigned as being synapsin I-, II-
or III-like, has been found in invertebrates, including
Aplysia (Klagges et al., 1996; Hilfiker et al., 1998; Angers
et al., 1999; Kao et al., 1999). The synapsins have been proposed to
play a major structural role in the assembly and maintenance of the
reserve pool of SVs both during development and in mature neurons, as
well as a dynamic role in controlling SV transitions from the reserve
pool to the releasable pool during neuronal activity (Greengard et al.,
1993; Brodin et al., 1997; Benfenati et al., 1999), possibly by
modulation of the cross-linking of SVs to the actin-based cytoskeleton
of the nerve terminal through synapsin phosphorylation (Benfenati et
al., 1992a, 1993, Ceccaldi et al., 1995; Hosaka et al., 1999). More
recently, a post-docking role of synapsins has been proposed (Rosahl et
al., 1993, 1995; Hilfiker et al., 1998) that may involve a putative
ATPase activity of synapsins (Esser et al., 1998). However, despite
considerable work in the last few years and the generation of synapsin
knock-out mice (Rosahl et al., 1993, 1995; Li et al., 1995; Takei et
al., 1995; Ryan et al., 1996; Terada et al., 1999), the precise role of
synapsins in short-term plasticity remains elusive.
In this paper, we took advantage of well characterized cholinergic
synapses in the buccal ganglion of Aplysia californica expressing post-tetanic potentiation (PTP), facilitation, and synaptic
depression (Mothet et al., 1996; Doussau et al., 1998) to analyze the
role of the synapsins in neurotransmitter release and short-term
plasticity. Our data demonstrate that the ability of synapses to
express PTP and sustain release at a low rate of stimulation is
determined by the size or maintenance, or both, of the
synapsin-dependent reserve pool. Moreover, regulation by synapsin of a
membrane step of the release process allows synchronization of release
to be maintained and minimizes the appearance of high-frequency depression when the release rate exceeds the recycling kinetics.
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MATERIALS AND METHODS |
Material. Peroxidase-conjugated goat anti-rabbit
secondary antibodies were from Bio-Rad (Milano, Italy), and the
chemiluminescence detection system (Renaissance Western Blot
Chemiluminescence Reagent Plus) was from New England Nuclear (Brussels,
Belgium). Synapsin I was purified from bovine forebrain as described
previously (Benfenati et al., 1992b) and stored in 200 mM NaCl, 25 mM Tris-Cl, pH
7.5. Synapsin IIa was expressed in Sf9 cells infected with the
baculovirus expression vector pVLSynIIa encoding for rat synapsin IIa
and purified by affinity chromatography (Siow et al., 1992; Nielander et al., 1997). Nitrocellulose membranes (0.2 µm pore size) were from
Schleicher & Schuell (Milano, Italy). The antibodies used in the
injection experiments were obtained in rabbits by injecting either
bovine synapsin I (antibody G177) or purified rat synapsin IIa
(antibody G423) with Freund's complete adjuvant. Preimmune sera were
used as controls. Antisera were depleted of contaminating anti-keratin
antibodies by passing through a human keratin-Sepharose 4B column as
described previously (Girault et al., 1989). All other chemicals of
analytical grade were obtained from Sigma (Milano, Italy). Immune
rabbit serum recognizing with high-affinity vesicle associated membrane
protein (VAMP)/synaptobrevin in Aplysia (Poulain et
al., 1993) was a generous gift of Dr. C. Montecucco (Padova, Italy).
Tetanus toxin was kindly provided by Dr. P. Boquet (Nice, France).
Immunoblotting. Freshly dissected Aplysia ganglia
or rat cerebral cortices were homogenized with a Teflon-glass
homogenizer in electrophoresis sample buffer (Laemmli, 1970). The
homogenates were boiled for 4 min, and the insoluble material was
removed by centrifugation. Proteins were separated by SDS-PAGE
(Laemmli, 1970) and electrophoretically transferred to nitrocellulose
membranes (Towbin et al., 1979). Immunoblotting was performed as
described previously (Benfenati et al., 1992b). Membranes were blocked
in Tris-buffered saline (50 mM TrisCl, 200 mM NaCl, pH 7.4) containing 5% (w/v) nonfat dry
milk and incubated overnight at 4°C with preimmune, G177, or G423
serum (1:1000 dilution). After washing and incubating with horseradish
peroxidase-conjugated goat anti-rabbit secondary antibodies
(1:3,000-1:10,000 dilution), immunoreactivity was detected using the
chemiluminescence system according to the manufacturer's instructions.
To analyze the specificity of the immunostaining, the primary
antibodies were preadsorbed with a molar excess of purified bovine
synapsin I or a mixture of purified bovine synapsin I and rat
recombinant synapsin IIa for 12 hr at 4°C before immunoblotting.
Acetylcholine release and electrical recordings at
Aplysia synapses. Experiments were performed at
identified inhibitory cholinergic synapses in buccal ganglia of
Aplysia californica (70-120 gm body weight; Marinus Inc.,
Long Beach, CA) (Gardner, 1971; Johannes et al., 1996; Mothet et
al., 1996; Doussau et al., 1998, 2000). Detailed procedures were
described previously (Doussau et al., 1998). During the experiments,
two presynaptic cholinergic interneurons [B4 and B5, according to
Gardner (1971); 80-150 µm in diameter] and one postsynaptic neuron
(B3, B6, or B8; 80-200 µm in diameter) were impaled with two glass
microelectrodes (3 M KCl,
Ag/AgCl2, 2-10 M ). Acetylcholine (ACh)
release from a presynaptic neuron was monitored for the duration of the
experiments (up to 24 hr continuous recording) by evoking an action
potential every 40 sec (0.025 Hz). The ACh release determined at this
low frequency was termed "basal release." To avoid overlap of the
postsynaptic responses originating from the two presynaptic neurons,
the stimulus protocols were alternated every 20 sec. ACh release was
estimated by measuring the amplitude of evoked IPSC using the
conventional two-electrode voltage-clamp technique. The holding
potential of the postsynaptic neuron was maintained 30 mV above
ECl as described (Doussau et al., 1998).
Post-tetanic potentiation. To initiate PTP, four trains of
action potentials (1 sec each at 50 Hz) were produced at 10 sec intervals. Then, the stimulation rate was returned to control conditions. For each episode, the amplitude of PTP was determined as
the maximal IPSC observed during the 5 min after the conditioning stimulus and was normalized with respect to the mean amplitude of the
10 IPSCs preceding the tetanus. In several experiments, when synapsin
was neutralized, the conditioning stimulus induced a depression that
preceded PTP. The amplitude of this post-tetanic depression (PTD) was
determined as the minimal IPSC observed within 40 sec after
conditioning and normalized as described for PTP.
Synaptic depression. In several experiments, synaptic
depression was induced by stimulating presynaptic neurons at 0.5 Hz for
20 min. Plateau levels were calculated as the mean amplitude of 50 IPSCs starting from the 18th min of stimulation at 0.5 Hz and expressed
as a percentage of the mean IPSC amplitude recorded under the basal
rate of stimulation. To induce synaptic depression, we also used 10 and
50 Hz trains of 1 sec duration. The 10 Hz trains could be repeated as a
series of four trains at 2-3 min intervals without induction of PTP,
provided a 15-25 min period of rest (stimulation at 0.025 Hz) was kept
between two series of stimuli. Depression in amplitude of IPSCs
elicited at 20 msec interpulse time intervals was examined by analyzing
the amplitude of IPSCs during 50 Hz postsynaptic responses evoked by
the first of the four conditioning train stimuli used to initiate PTP
(see below). However, the postsynaptic responses evoked by the second, third, and fourth trains of a conditioning protocol were not analyzed because significant PTP or PTD, or both, appears just after the first conditioning stimulus.
Calculation of IPSC amplitude during high-frequency repetitive
stimulation. When the stimulation time interval is shorter than
the time needed for an IPSC to return to baseline, IPSCs overlap
partially. In ~20% of the neuron pairs examined, the IPSC decay was
better fitted by a bi-exponential time course (with  1 ~9-15 msec and 2
~20-50 msec). Therefore, the actual amplitude of IPSC number
n in a train was calculated by deducing the extrapolated residual amplitudes of the five previous IPSCs following the equation: In ~ Ipeak
n   
(Ini*(a*ei* t/ 1 + b*ei*t/2)),
where  t was the time interval between two subsequent IPSCs, 1 and 2 were the
decay times of previous IPSCs, and a and b (with
a + b = 1) were the respective weights of
the two exponentials. 1,
2, a, and b were
calculated in each experimental condition from several IPSCs evoked by
a single action potential. The analysis was limited to the first 20 IPSCs of each 50 Hz train because of the progressive decrease of phasic
release and the concomitant increase of desynchronized release in the
course of the tetanic stimulation. This phenomenon can probably be
attributed to the fact that during tetanic stimulation, the
intraterminal Ca2+ levels approach the
threshold for exocytosis.
Paired-pulse facilitation. To study paired-pulse
facilitation (PPF), two brief supraliminar depolarizing pulses
of 5 msec were applied at 15-8000 msec intervals to the presynaptic
neuron. For each twin response, facilitation F was calculated from the equation F = (I2 I1)/I1
(Mallart and Martin, 1967), where I1 and I2 are the amplitudes of the IPSC
evoked by the conditioning and test stimuli, respectively. The actual
amplitude of I2 was determined as described above
for In in a train. Because of the high
variability of paired-pulse facilitation at the cholinergic synapses
studied, determination of mean paired-pulse facilitation was calculated
from at least 12 paired-pulse evoked responses for the same interpulse
time interval.
IPSC rise time. To avoid uncertainty in determining
t0 (IPSC foot) and
tmax (IPSC peak), the time to rise
from 20 to 80% of maximal IPSC amplitude was determined. Mean IPSC
rise times were determined from at least 10 IPSCs recorded under the
same experimental condition.
Extracellular media. Dissected buccal ganglia were
maintained at 22°C using a Peltier-plate system and superfused
continuously (10 ml/hr) with a physiological "control medium"
containing (in mM): NaCl 460, KCl 10, CaCl2 33, MgCl2 50, MgSO4 28, Tris buffer 10, pH 7.5. Note that to
minimize spontaneous neuron firing activity, the physiological medium
used contained high total divalent cations, a condition known to shield
plasma membrane-fixed negative charges and to alter the intramembranous
electric field. When extracellular [CaCl2]
needed to be modified, the respective concentrations of [CaCl2] and [MgCl2]
were calculated according to the following equations:
[CaCl2] (mM) = Q*(83 + [MgS04])/(Q + 1)
and [MgCl2] (mM) = 83 [CaCl2], with Q being the
[Ca2+]/[Mg2+]
ratio (Doussau et al., 1998); in control medium,
[Ca2+]/[Mg2+]
was 0.42.
Intraneuronal injection procedure. Injection electrodes were
pulled from glass tubing without a capillary and contained a silver
wire to allow the electrophysiological monitoring of the impalement.
Samples to be injected were mixed with a vital dye (fast green FCF,
10% v/v; Sigma). The samples were air pressure-injected using a
picopump PV820 (WPI Ltd.) under visual and electrophysiological monitoring. The injected volume was in the range of 1-5% of the cell
body volume. After injection, the injection micropipette was removed.
After intracellular injection, only neurons with membrane potentials of
60 to 45 mV and with no alterations in the action potentials were analyzed.
Data presentation. Because of the intrinsic variability of
different neuronal preparations, the measurements obtained for a neuron
under a given experimental condition and time range were averaged and
normalized against the corresponding mean measurement made during
control conditions (i.e., before the time of injection of antibodies,
proteins, or buffer). Then, the normalized values obtained from
n different neurons were averaged and presented as
means ± SEM. Note that because of the difficulty of applying the
experimental protocols in all experiments following the same time
schedule, data values were pooled for given periods of time as follows:
"+300 min" refers to time range 280-320 min after injection,
"+480 min" to 430-530 min, and "+1100 min" to 900-1300 min.
Statistical significance. The significance of the
differences observed between groups of measurements were determined by
paired or unpaired t tests when a single comparison was
done. When multiple comparisons were done, the significance was tested
by using either one-way or two-way ANOVA analysis depending on the
number of factors considered (treatment alone, time alone, or treatment
plus time); this analysis was followed by the Tukey-Kramer test. To
allow multiple comparisons of the 10 or 50 Hz trains recorded under various experimental conditions, the decay of IPSC amplitude in the
trains was fitted with the equation y = (100 c)*e-t/ + c, where c is the plateau reached, t
the time, and the time constant of depression. The residual
variance was analyzed and significance was determined by the
Fisher-Snedecor test. For all comparisons, when
p > 0.05, the difference observed was denoted as
nonsignificant (n.s.).
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RESULTS |
Specificity of the anti-synapsin antibodies
The ability of the antibodies directed against mammalian synapsins
to recognize the Aplysia synapsin homolog is shown in Figure 1. Antibody G423 (lanes 1 and
3), recognizing the highly conserved domain C of synapsins
(F. Benfenati, unpublished observation), and antibody G177 (lane
5), recognizing synapsins I and, to a lesser extent, synapsin II
in mammals (Cibelli et al., 1996), specifically detect synapsin-like
proteins in acid extracts of Aplysia ganglia. These appear
as a doublet of ~55-60 kDa molecular mass (Fig. 1). These
immunoreactive species were not detectable when the corresponding
preimmune sera were used in the immunoblot assays (lane 2 and data not shown). The immunoreactivity was virtually abolished or
markedly decreased in intensity when the antibodies had been
preadsorbed with a molar excess of purified bovine synapsin I (G177
antibody; lane 6) or a mixture of purified bovine
synapsin I and rat recombinant synapsin IIa (G423 antibody; lane
4).
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Figure 1.
Synapsin-like immunoreactivity in
Aplysia nervous tissue. Proteins from extracts of rat
cerebral cortex (5 µg protein, lane 1) and
Aplysia ganglia (30 µg protein, lanes
2-6) were separated by SDS-PAGE on 9% polyacrylamide
gels, transferred to nitrocellulose membranes, and immunoblotted with
the following antibodies diluted 1:1000: lanes 1 and
3, G423; lane 2, preimmune G423;
lane 4, synapsin-preadsorbed G423; lane
5, G177; lane 6, synapsin preadsorbed G177.
Immunoreactivity was revealed using the chemiluminescence detection
system. Preadsorption of the antibodies markedly decreased or virtually
abolished the immunoreactivity in the samples from both rat (data not
shown) and Aplysia nervous systems. Molecular mass
markers are shown on the right in kilodaltons.
rSynIa,b, Rat synapsin Ia/Ib; rSynIIa,
rat synapsin IIa; aSynLI, Aplysia
synapsin-like immunoreactivity; rSynIIb, rat synapsin
IIb.
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Effect of the intraneuronal injection of specific anti-synapsin
antibodies on evoked ACh release
In each hemibuccal ganglion of Aplysia, two identified
presynaptic neurons, B4 and B5, make cholinergic synapses with the same
set of postsynaptic neurons (including B3, B6, and B8 cells) (Gardner,
1971). ACh release evoked by single action potentials elicited in
either B4 or B5 neurons can be monitored by measuring the amplitude of
IPSC recorded in one of these postsynaptic neurons. To probe the role
of synapsins in neurotransmitter release, the specific anti-synapsin
antibodies (either G423 or G177) were pressure injected into either B4
or B5. For all experiments described in this paper, the remaining
neuron was not injected (Fig.
2A) but was submitted
to the same stimulation protocol and thus served as an internal control
of the stability of synaptic efficacy and evoked neuroexocytosis for
the whole duration of the experiment (up to 24 hr).
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Figure 2.
Effects of anti-synapsin antibody injection on ACh
release. Evoked ACh release was monitored at identified synapses in the
buccal ganglion of Aplysia californica. A, Schematic
drawing of the neuronal connections and the recorded neurons.
pre1/pre2, B4/B5 presynaptic neurons;
post, B3, B6, or B8 postsynaptic neuron;
Ab, antibody. B, A representative
experiment is illustrated. The amplitude of IPSCs evoked every 40 sec
was averaged during periods of 30 min (means of 45 measurements ± SEM) before and after the injection (arrow) of the G423
anti-synapsin antibody.  denotes values from the antibody-injected
neuron, and  denotes values from the control, noninjected neuron.
Values were normalized with respect to the mean IPSC amplitudes
recorded before the injection. C, The mean amplitude of
IPSCs evoked from noninjected (n = 16, white
bars), preimmune-injected (n = 4, gray bars), and antibody-injected (G423,
n = 9; G177, n = 3;
black bars) neurons is reported at the indicated times
as a percentage (±SEM) of the mean IPSC amplitude before the
injection. Note that the values reported at +1100 min after injection
were obtained from only three antibody-injection experiments (G423,
n = 2; G177, n = 1) and two
preimmune-injection experiments in which synaptic activity could be
maintained for such a long time. D, Averaged IPSCs
(n = 5 for each condition) recorded
before and +480 min after
the time of injection from control, noninjected (top
panel) and antibody-injected (bottom
panel) neurons. Averaged IPSCs have been scaled to allow
comparison of their time course. E, The mean percentage
changes in the IPSC rise time with respect to the mean value observed
before injection are reported at the indicated times after the
injection for noninjected (n = 10; white
bars), preimmune-injected (n = 4;
gray bars), and antibody-injected (n = 6; black bars) neurons. Means at +1100 min are from
only six noninjected, three preimmune-injected, and three
antibody-injected neurons. C, E, Multiple
comparisons were performed by a two-way ANOVA analysis followed by
Tukey-Kramer test on the relative rise time change considering two
factors: treatment and time. C, n.s, Not
significant. E, Effect of treatment on rise time in
antibody-injected neurons versus preimmune-injected or noninjected:
**p < 0.001; preimmmune-injected versus
noninjected: not significant; effect of time alone on IPSC rise time of
noninjected or preimmune injected neurons: not significant.
F, A representative experiment of a series of four,
performed as described in B, during which anti-VAMP
antibodies were injected. The hatched area denotes bath
application of 2 µM tetanus toxin
(TeNT). G, The mean IPSC rise
times were determined in antibody-injected (black bar)
and noninjected (white bar) neurons 235-245 min after
anti-VAMP antibody injection and 1 hr after tetanus toxin application.
n.s, Not significant.
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When either the B4 or the B5 presynaptic neuron was microinjected with
the G423 or G177 antibodies or preimmune serum, no significant change
in the amplitude of IPSC was recorded for the following 500 min as
compared with IPSC evoked from the control, noninjected neuron (Fig.
2B,C). These results indicate that
at low rates of stimulation, the amount of ACh released per impulse remains unmodified after injection of anti-synapsin antibodies. This is
consistent with observations made in the lamprey reticulospinal synapses (Pieribone et al., 1995). In several experiments, recording of
IPSCs could be made with a high signal-to-noise ratio that allowed
accurate analysis of IPSC kinetics. The rise time of the IPSC evoked by
the stimulation of the antibody-injected neuron was significantly
slowed down (Fig. 2D,E) in either
G423-injected (n = 4) or G177-injected
(n = 2) neurons, in the absence of significant changes
in IPSC decay time. In four of the six experiments analyzed, significant slowdown of the IPSC rise time was detectable as early as
160-180 min after injection (slow by 1.22; p < 0.05;
data not shown). This effect lasted for the rest of the recording, up
to 1100 min after the injection (Fig. 2E). Closely
similar results were observed after the injection of a conserved
synapsin peptide into the squid giant presynaptic terminal (Hilfiker et
al., 1998).
The above-mentioned changes in IPSC kinetics cannot be attributed to an
alteration of recording conditions during these very long experiments
because the IPSCs evoked by stimulation of the control, noninjected
neuron and recorded from the same postsynaptic neuron (Fig.
2A) were not significantly modified (Fig.
2D,E). In addition, the slowdown of
the IPSC rise time was specific for the presence of anti-synapsin
antibodies because the IPSCs evoked by stimulation of
preimmune-injected neurons were not significantly slower than those
obtained from control, noninjected neurons (Fig. 2E,
gray bars). Hence, the slowdown of IPSC rise time observed after synapsin neutralization indicates that synapsin may be involved in some post-docking event of exocytosis, as suggested previously (Rosahl et al., 1993, 1995; Hilfiker et al., 1998).
Because synapsins are abundant SV-associated proteins, the possibility
that the IPSC slowdown was the result of steric hindrance of exocytosis
caused by extensive coating of SVs by the antibody cannot be excluded.
To test this possibility, we examined the consequences of the injection
of antibodies directed against VAMP/synaptobrevin, an integral SV
membrane protein present on SVs with a number of copies similar to or
higher than those of synapsin (Walch-Solimena et al., 1995; Taubenblatt
et al., 1999), which participates in the formation of the fusion core
complex in exocytosis (for review, see Robinson and Martin, 1998;
Benfenati et al., 1999) and the cleavage of which by tetanus
toxin blocks neurotransmitter release (Schiavo et al., 1992). Figure 2
shows that when anti-VAMP antibodies were injected into presynaptic
neurons, they did not significantly affect the evoked ACh release (Fig.
2F) and IPSC rise time (Fig. 2G) for >5
hr (the first significant effects of anti-synapsin injection appeared
after 3 hr). This is consistent with previous observations obtained in
PC12 cells and Aplysia synapses (Elferink et al., 1993;
Poulain et al., 1993). Although no release parameter was modified after
anti-VAMP injection, the ability of anti-VAMP antibodies to prevent the
blocking action of tetanus toxin in all the experiments that were
performed (Fig. 2F) indicates that they did indeed
reach their SV target, as published previously (Poulain et al., 1993).
These results show that a procedure aimed at coating SVs with
antibodies does not necessarily alter ACh release kinetics and that the
effects observed after injection of anti-synapsin antibodies are likely
attributable to synapsin neutralization or interference with some
post-docking activity of synapsin.
Effects of intraneuronal injection of anti-synapsin antibodies or
of synapsin I on post-tetanic potentiation
PTP is a form of short-term synaptic plasticity that is
characterized by a transient increase in synaptic strength in response to a presynaptic tetanic conditioning stimulus (for review, see Zucker,
1996, 1999). Although postsynaptic mechanisms have been reported to
play some role in PTP at certain Aplysia synapses (Bao et
al., 1997), PTP is mostly caused by an enhanced neurotransmitter release initiated by an accumulation of
Ca2+ during tetanic stimulation (Kretz et
al., 1982; Swandulla et al., 1991; Kamiya and Zucker, 1994; Bao et al.,
1997) and has been proposed to involve the recruitment of SVs from the
reserve pool, possibly under the control of synapsins (Greengard et
al., 1993; Rosahl et al., 1995; Zucker, 1996, 1999). Hence, we
investigated the possible implication of synapsin in PTP.
At the identified cholinergic synapses of the buccal ganglion, PTP is
reproducibly initiated by application of four trains of impulses of 1 sec at 50 Hz. A typical PTP is illustrated in Figure
3A, top panel. When
PTP was elicited under control conditions, evoked ACh release increased
with respect to basal values for a mean duration of 35 ± 2 min,
reaching a peak increase of 48 ± 3% (means ± SEM; 41 PTPs
initiated in 26 neurons). In most experiments performed under control
conditions, PTP was found to reach its maximal amplitude within 40 sec
after the fourth conditioning stimulus was applied (data not
shown).
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Figure 3.
Post-tetanic potentiation (PTP) is
depressed after injection of anti-synapsin antibodies.
A, Four conditioning tetanic stimulations (1 sec at 50 Hz, each) were applied to initiate PTP. The expression of PTP was
monitored by plotting the IPSC amplitude, evoked at 0.025 Hz, before
and after the conditioning stimuli (arrow) were applied.
The three PTPs illustrated were recorded at the indicated times, before
(top panel) and after (middle and
bottom panels) G423 antibody injection. Note the
appearance of post-tetanic depression (PTD) after
antibody injection. B, Representative experiment in
which the amplitudes of PTP (circles) and PTD
(triangles) were followed as a function of time after
the injection (arrow) in G423-injected (,  ) and
noninjected (, ) neurons. PTP and PTD amplitudes were determined
as described in Materials and Methods. C, The amplitudes
of PTP (top panel) and PTD (bottom
panel) induced in noninjected (white
bars), preimmune-injected (gray bars),
and antibody-injected (black bars) neurons are shown as
means ± SEM. PTP amplitudes were normalized against the mean PTP
amplitudes recorded in the same neurons before the injection. Average
PTD amplitudes are expressed as percentage of the mean basal IPSC
amplitude determined immediately before the conditioning stimulus.
Number of experiments at +300 or +480 min: anti-synapsin antibodies,
n = 9 (G423, n = 6; G177,
n = 3); preimmune, n = 4;
noninjected, n = 13. Number of experiments at +1100
min: anti-synapsin antibodies, n = 3 (G423,
n = 2; G177, n = 1); preimmune,
n = 2; noninjected, n = 5. Comparison between the different groups in C (top
graph) was tested by using two-way ANOVA followed by the
Tukey-Kramer test. PTP reduction in anti-synapsin-injected neurons was
p = 0.0045; PTP elicited in antibody-injected
versus preimmune-injected or noninjected neurons was
p = 2 × 106 and 1.7 × 106, respectively. All
other comparisons: n.s.
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Figure 3B shows the comparative time course of PTP elicited
from a presynaptic neuron injected with the G423 antibody and from the
control, noninjected neuron in the same preparation. After injection
(seven experiments), PTP was strongly depressed or abolished within 500 min of the antibody injection (Fig. 3C, top
graph). In the three experiments performed using G177 antibodies, a similar disappearance of PTP was observed, but with slower time course, possibly caused by a lower titer or a lower affinity of the
antibody. PTP recorded from noninjected neurons in the same preparations was not significantly changed (Fig. 3C,
top graph, open bars), albeit a slight decrease
could be observed at very long recording times, possibly because of
aging of the preparations (Fig. 3C, +1100). No
significant changes in PTP amplitude could be observed after injection
of preimmune serum as compared with noninjected neurons (Fig.
3C). The abolition of PTP observed in anti-synapsin-injected
neurons was not associated with any significant change in basal ACh
release, except for the effect on IPSC kinetics (Fig. 3C,
top graph, compare with 2C,E). These
observations suggest that the mobilization of a synapsin-dependent
reserve pool of SVs is necessary for neurons to express PTP.
When conditioning protocols were induced at a time when PTP was
abolished, a transient decrease in the amplitude of the postsynaptic response was observed (Fig. 3A-C). We termed it
PTD. The occurrence of PTD was not restricted to a particular synapse
of the network, because we observed this same phenomenon by recording
IPSCs in B3, B6, or B8 postsynaptic neurons in the same hemiganglion
(Fig. 4A). One of the
three experiments is illustrated in Figure 4B. It is
possible that when the synapsin-dependent reserve pool of SVs is
depleted after action of the antibodies, the readily releasable pool
becomes exhausted during the conditioning stimuli and cannot be
replenished at a sufficient rate, leading to transient PTD. Such an
effect should be emphasized when increasing the duration of
conditioning stimuli or minimized when the release probability is
reduced. Indeed, the magnitude and time course of PTD depended on the
number of 50 Hz trains applied to initiate PTP (Fig. 4C), and when PTP protocols were induced in low extracellular
Ca2+ (Ca/Mg = 0.14) to reduce release
probability, no PTD was initiated unless the number or duration of the
50 Hz trains was increased (data not shown).
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Figure 4.
Amplitude of PTP and PTD depends on the
release conditions. A, Schematic representation of the
buccal ganglion of Aplysia californica.
B4 and B5 denote presynaptic cholinergic
neurons. B3, B6, and B8
are their postsynaptic targets. B, A representative
experiment of a series of three is shown. Conditioning stimuli were
applied as described in Figure 3 at G423 antibody-injected
(B4, arrow) and noninjected
(B5) neurons. The ensuing PTDs () or PTPs () were
recorded at the indicated times after the injection by impaling
successively B3, B6, and B8 neurons. C, Effects of the
strength of the conditioning stimulus on PTP and PTD amplitudes
recorded from G423-injected () and noninjected () neurons at the
indicated times after the injection. The following conditioning stimuli
were applied: 1 train (1 sec at 50 Hz) (S), 4 trains (SS), 10 trains (SSS).
D, The effect of anti-synapsin antibody injection on PTP
amplitude depends on the release probability. Relative PTP amplitudes
were determined as a percentage of the mean PTP amplitude recorded
before the injection as described in Figure 3C. PTP
amplitudes were determined from antibody-injected (G423,
n = 2; G177, n = 1;
filled bars) and noninjected (n = 3;
open bars) neurons 460-515 min after antibodies were
injected. Then, a low Ca2+-containing medium (Ca/Mg
ratio = 0.14) was substituted for the high Ca2+
medium (Ca/Mg ratio = 0.42), and the relative PTP amplitudes were
determined (i.e., 520-550 min after antibody injection).
*p < 0.05, **p < 0.01;
Student's unpaired t test.
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As shown in Figures 3A and 4C, PTD overlaps with
PTP. This raises the question of whether the induction of PTD
contributes to the inhibition of PTP. First, the duration of PTD was
much shorter than that of PTP (Figs. 3A, 4C).
Second, after PTP had been depressed by injecting G423 or G177
antibodies under normal Ca2+ conditions
(Fig. 4D, compare the left open and
filled bars), a decrease in extracellular
Ca2+, applied to minimize PTD, induced
only small recovery of PTP as compared with the PTP recorded in
noninjected neurons under the same Ca2+
condition (Fig. 4D, filled bars). These
data confirm that synapsin neutralization directly affects PTP mechanisms.
To further demonstrate the involvement of synapsin in the expression of
PTP, we investigated the effects of an increase in the intraneuronal
synapsin pool by injecting purified mammalian synapsin I into the
presynaptic neurons. Purified dephosphorylated synapsin I (Fig.
5, Syn) or the corresponding
buffer was injected into B4 or B5 neurons. Assuming a homogenous
distribution of the protein in the cytosol, the final intrasomatic
concentration of synapsin was estimated to range between 0.5 and 1 µM (15 µM in the
injection micropipette). In contrast to the antibody injection experiments, no change in either IPSC amplitude or kinetics could be
detected up to 8 hr after the intracellular application of either
synapsin or synapsin buffer as compared with the noninjected neurons
(Fig. 5A-C). However, a marked and progressive
enhancement in PTP amplitude was observed when ACh release was evoked
from the synapsin-injected neurons (Fig.
5D-F). The change in PTP amplitude started soon after injection of synapsin (Fig. 5E, compare
with 3B).
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Figure 5.
Synapsin injection increases PTP without
changing basal ACh release. A, Effects of synapsin I
injection on basal release. A representative experiment of a series of
six is illustrated. Dephosphorylated synapsin I was injected at an
estimated final concentration in the soma of 0.5-1 µM.
The experiments and data presentation are as described in Figure 2. and denote IPSC amplitude from synapsin I-injected and noninjected
neurons, respectively. B, C, The mean
amplitude of IPSCs and the mean IPSC rise time evoked from
synapsin-injected (black bars; n = 6) and buffer-injected (gray bars;
n = 6) neurons after the injection are reported as
a percentage of the IPSC amplitude and rise time determined before
injection. D-F, Effects of synapsin I
injection on PTP. D, Typical PTP recorded 300 min before
(top panel) and 320 min after (bottom
panel) synapsin injection. Same presentation as in
Figure 3A. E, PTP amplitude was followed as a function
of time after the injection (arrow) of synapsin.
Presentation of data is as in Figure 3B. F, Means
(±SEM) of the PTP amplitudes observed after synapsin (black
bars) or buffer (gray bars) injection.
n = 5 at +300 min and n = 3 at +480 min for synapsin-injected neurons, and n = 3 at both +300 min and +480 min for buffer-injected neurons.
Statistical analysis was performed by using two-way ANOVA followed by
Tukey-Kramer test. **p = 0.026 versus buffer
injection on PTP amplitude. n.s, Not significant.
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These observations indicate that synapsins play a major role in PTP
mechanisms, possibly by determining the size of the reserve pool or
controlling the transition of SVs between the reserve and the
releasable pools, or both. A recruitment of reserve SVs has also
been postulated to contribute to the maintenance of synaptic efficacy.
At active mammalian synapses, the exo-endocytotic cycle of SVs takes
~10-30 sec to complete (Stevens and Tsujimoto, 1995; Klingauf et
al., 1998; Murthy and Stevens, 1999). Thus, when the rate of
stimulation is increased over the recycling-refilling capacity
(i.e., >0.03-0.1 Hz), synaptic efficacy cannot be maintained. However, in most synapses, the recruitment of SVs from the reserve pool
contributes to the rapid replenishment of the ready releasable pool and
minimizes depression (Ryan et al., 1993; Stevens and Tsujimoto, 1995;
Dobrunz and Stevens, 1997). According to this model, the synaptic
depression observed in lamprey synapses injected with anti-synapsin
antibodies and subjected to prolonged 18-20 Hz stimulation is
attributable to the exhaustion of the reserve pool (Pieribone et al.,
1995). However, synapsin has also been proposed to play a role in some
post-docking event of exocytosis (Rosahl et al., 1993, 1995; Hilfiker
et al., 1998) (Fig. 2D,E). Hence,
this raises the question of whether the inability of synapses to
maintain neurotransmitter release during high-frequency stimulation is
caused by exhaustion of a synapsin-dependent reserve pool or impairment
of a post-docking function of synapsin, or both.
Synapsin neutralization induces fast and marked
synaptic depression
The replenishment time constant of release sites by reserve SVs is
on the order of several seconds (Stevens and Tsujimoto, 1995). Thus,
the application of brief (1-2 sec), high-frequency stimulations may
permit investigation of the effects of an altered post-docking function
of synapsin induced by its neutralization by minimizing the effects
caused by exhaustion of reserve pool. The amplitude of IPSCs was
analyzed during 50 Hz postsynaptic responses evoked by the first of the
four conditioning train stimuli used to initiate PTP and during 10 Hz
trains for 1 sec that do not elicit PTP (for details, see Materials and
Methods). Figures 6, A and
B, and 7A show typical 50 and 10 Hz responses
elicited from the same presynaptic neurons, before and after PTP had
been abolished by previous injection of G423 or G177 antibodies (i.e., 500 min after injection). After anti-synapsin antibody injection, the
IPSC amplitude could not be maintained during 10 and 50 Hz trains
(Figs. 6C, 7B). No comparable effect was detected
in control, noninjected, or preimmune injected neurons (Fig.
7B). The small increase in 10 Hz-induced depression observed in control neurons indicates that aging
of the preparations may also produce a small fatigue in ACh release.
The time course of the increase in depression induced by the
anti-synapsin antibody injection started within 2 hr after injection
and developed after a time course that was closely similar to that of
PTP inhibition (50 Hz train: data not shown; 10 Hz train: Fig.
7C, compare with Fig. 3B). By contrast, although
synapsin I injection was effective in enhancing PTP (Fig. 5), no
significant change in 10 or 50 Hz-induced depression could be detected
in synapsin I-injected neurons as compared with neurons injected with
synapsin buffer alone (Figs. 6D, 7D).
Taken together, these findings indicate that after neutralization of
synapsin, ACh release cannot be maintained at high levels during a
prolonged high-frequency stimulation. Interestingly, the increase in 10 and 50 Hz-induced depression was already significant at the second stimulus in the train (i.e., that elicited at a 20 or 100 msec interpulse interval) (Figs. 6 and 7, legends). This time scale is by
far smaller than the mean SV residency on the plasma membrane (2-5
sec) (Klingauf et al., 1998; Murthy and Stevens, 1999). Hence, after
synapsin neutralization, the inability of the synapse to sustain ACh
release during short high-frequency bursts may result from alteration
of a synapsin function that operates during a membrane step of the
release process.
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Figure 6.
Frequency stimulation-induced depression (50 Hz)
is increased after synapsin neutralization. Pairs of neurons, one
injected with anti-synapsin antibodies, preimmune serum, buffer alone,
or purified synapsin I and the other kept intact, were subjected to
repetitive stimulation at 50 Hz. A, B,
Representative postsynaptic responses evoked by 50 Hz stimulation
applied before (thin line) and 500 min after G423
(A) or G177 (B) injection
(thick line) (i.e., at a time when PTP was abolished).
Note the different time scales in A and B.
C, The mean amplitude of the 20 first IPSCs of the 50 Hz train
response was determined (see Materials and Methods), expressed as a
percentage of the first IPSC of the train (i.e., which amplitude is
identical to IPSC amplitude recorded under basal stimulation frequency
at 0.025 Hz), and normalized against the corresponding values
determined before injection. Mean values (±SEM) were calculated from
recordings made 430-530 min after the injection of anti-synapsin
antibodies (, 6 G423-injected neurons) or preimmune serum
(gray-filled circle, 4 neurons). denotes mean values
from 10 control, noninjected neurons in the same preparations.
D, Same kind of measurements as in C
except that synapsin I ( , n = 3) or buffer alone
(gray-filled square, n = 3) was
injected. The comparison among the extents of 50 Hz-induced depression
observed after the various treatments was performed as described in
Materials and Methods (two-way ANOVA followed by a
Fisher-Snedecor test). **p < 109 anti-synapsin versus
noninjected or preimmune. Significance of the differences observed in
the second response to 50 Hz trains was tested as described in
Materials and Methods using the Tukey-Kramer test after one-way ANOVA:
p = 0.016 and 0.004 anti-synapsin versus noninjected and
preimmune, respectively.
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Figure 7.
Frequency stimulation-induced depression (10 Hz)
is increased after synapsin neutralization. Representative postsynaptic
responses evoked by 10 Hz (A) stimulation applied
before (thin line) or 485 min after (thick
line) G423 injection. B, The mean amplitude of
the 10 IPSC responses evoked by the 10 Hz train 430-530 min after
injection of antibodies was determined (see Materials and Methods) and
normalized against the corresponding values determined before
injection. Mean values (±SEM) were calculated after the injection of
anti-synapsin antibodies (, n = 6) or preimmune
serum (gray-filled circle, n = 3). denotes mean values from nine control, noninjected neurons in the same
preparations. C, The mean amplitude of the 10th response
to the 10 Hz trains recorded in the course of a representative
experiment was normalized with respect to the amplitude of the first
IPSC of the train (4-6 trains were averaged at each time indicated;
note that SEM bars are smaller than the symbol size) and plotted as a
function of time after injection (arrow) in
G423-injected () and noninjected () neurons. D,
Same kind of measurements as in B except that synapsin I
(, n = 3) or its buffer ( ,
n = 3) was injected. Significance of the
differences observed in the responses to 10 Hz trains under the various
treatments was tested as described in Figure 6. Whole train:
**p < 109
anti-synapsin versus noninjected or preimmune. Second response to 10 Hz
trains: **p = 0.02 and 0.03, anti-synapsin versus
noninjected and preimmune, respectively.
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To better characterize fast depression (and thus the underlying
post-docking synapsin function), we next examined the
Ca2+ dependency of 10 Hz-induced synaptic
depression (three experiments). Under control conditions, no or little
synaptic depression was detected when 10 Hz trains were elicited in the
presence of a low Ca2+-containing medium
(with
Ca2+/Mg2+ = 0.14 to reduce the release probability), whereas depression could be
observed when using a high
Ca2+-containing medium
(Ca2+/Mg2+ = 0.42 or 2.1). When similar experiments were repeated 500 min after the injection of G423 (n = 3) antibody, at
a time PTP was abolished, 10 Hz-depression was dramatically enhanced
when the extracellular Ca2+ concentration
was raised to higher levels, whereas it was not affected at low
Ca2+ (Fig.
8, compare open and
closed bars). These results suggest that the
synapsin-dependent regulation that operates at the membrane stage of
the release process to prevent high-frequency-induced depression is
limiting only when the probability of release is high.
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Figure 8.
Fast depression induced by anti-synapsin antibody
injection depends on the release probability. Trains of 10 Hz were
elicited in the presence of low (Ca/Mg = 0.14), normal (Ca/Mg = 0.42), or high (Ca/Mg = 2.1) Ca2+-containing
medium. Mean amplitudes of IPSCs evoked by the 10 Hz stimulation of
noninjected (open bars) and G423-injected
(filled bars) neurons were determined as
described in Materials and Methods, under the various conditions,
410-550 min after the time of antibody injection. Means (±SEM) were
calculated from three independent experiments performed under each
condition. Significance of the differences in the responses to 10 Hz
trains between antibody-injected and control, noninjected neurons was
as follows: Ca/Mg = 0.14: non-significant (ns);
Ca/Mg = 0.42 or 2.1: **p < 109.
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The time scale of the fast depression observed during high-frequency
stimulation after injection of anti-synapsin antibodies overlaps with
the time window (0-200 msec) for facilitation in the same
Aplysia synapses (Doussau et al., 1998). To determine whether synapsin neutralization primarily enhanced depression or
affected it indirectly through a decrease in facilitation, we
determined the extent of IPSC facilitation induced by paired stimuli
applied at time intervals ranging from 20 msec to 8 sec before and
360-480 min after (i.e., when PTP was abolished) injection of the
anti-synapsin antibody G423. The injection of G423 significantly decreased the amplitude of the second response of pairs elicited at all
the examined interpulse intervals (20 msec-8 sec). As shown in Figure
9A, this manifested as a
decrease in paired-pulse facilitation at the shorter time intervals and
as paired-pulse depression at longer interpulse intervals
(n = 4). Because the extent of depression was
comparable with that of the decrease in facilitation (Fig. 9A), this suggested that facilitation per se might be
unchanged after antibody injection. To dissociate facilitation from
depression, we took advantage of the observation that the effect on
depression is dramatically relieved under conditions of low release
probability (Fig. 8). Paired-pulse stimulation protocols were then
repeated using low Ca2+-containing medium
(Ca/Mg = 0.14) (Fig. 9B). Under these conditions, the
amplitudes of paired-pulse facilitation determined before and after
G423 injection were not significantly different (Fig. 9B)
(n = 3). These results suggest that facilitation per se
is not altered after neutralization of synapsin, consistent with previous observations in synapsin II or synapsin I/II knock-out mice
(Rosahl et al., 1995).
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Figure 9.
PPF is not affected after synapsin neutralization
under low release probability. Paired stimuli were elicited at various
interpulse intervals before () and 360-480 min after () the
intraneuronal injection of G423 antibody in the presence of either
normal (Ca/Mg = 0.42; A) or low (Ca/Mg = 0.14;
B) Ca2+-containing medium. The extent
of paired-pulse facilitation (means ± SEM from 4 experiments in A
and from 3 experiments in B) was calculated at the
indicated interpulse intervals. Note that values reported at the 40 sec
interval have been collected at the basal stimulation rate (0.025 Hz).
Statistical comparison of PPF extent before and after synapsin
neutralization: *p < 0.01; **p < 0.001.
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Paired-stimulation experiments in anti-synapsin-injected synapses show
that the time scale of paired-pulse depression extends over a 10 sec
range. This time scale exceeds the mean residency time of SVs at plasma
membrane but remains in the range of the replenishment kinetics of the
readily releasable pool of SVs (Stevens and Tsujimoto, 1995; Klingauf
et al., 1998; Murthy and Stevens, 1999). Accordingly, the inability to
maintain high synaptic efficacy after synapsin neutralization in
synapses subjected to twin stimulations applied at long intervals may
be attributable to an altered SV trafficking between the reserve pool
and the releasable pool. To investigate this possibility while
minimizing the effects of an alteration of synapsin function at
membrane level, we examined the effects of a prolonged increase in the
stimulation rate from 0.025 to 0.5 Hz for 20 min. Under these
conditions, the IPSC amplitude decreased and reached a stable level
(plateau) at ~50% of initial amplitude in ~10-15 min (Fig.
10A), as reported
earlier (Mothet et al., 1996). In a series of five experiments, 0.5 Hz-induced depression was examined before and after injection of G423
antibodies and in noninjected neurons in the same preparations (a
typical example is illustrated in Fig. 10A). In
anti-synapsin antibody-injected neurons subjected to repeated periods
of 0.5 Hz stimulation, the development of depression was significantly
accelerated as compared with control recordings made before injection
or in noninjected neurons (Fig.
10A,C). However, the extent of
depression was virtually unchanged, because the plateau level reached
during 0.5 Hz stimulation was not modified significantly (Fig.
10A,B). When the stimulation frequency was returned to 0.025 Hz, the initial phase of recovery from
depression was significantly faster in the antibody-injected neurons (Fig. 10A,D).
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Figure 10.
Depression and recovery from 0.5 Hz stimulation
periods are accelerated after synapsin neutralization.
A, Representative experiment from a series of 5. Presynaptic neurons were stimulated either at basal stimulation
frequency (0.025 Hz) or at 0.5 Hz for 20 min periods (horizontal
bars). IPSCs were evoked in both G423-injected () and
control, noninjected () neurons. Note the faster depression and
recovery rates that occurred after antibody injection.
B, The IPSC plateau levels (means ± SEM) reached
during 0.5 Hz stimulation, determined as described in Materials and
Methods, are reported for antibody-injected neurons
(filled bars) and control, noninjected neurons
(open bars). C, D, The
amplitude of IPSCs evoked during 0.5 Hz stimulation
(C) and after the stimulation frequency was
returned to 0.025 Hz (D) was averaged for periods
of 30 sec (0-2 min) or 60 sec (2-16 min), and the corresponding
plateau values were subtracted. Means ± SEM of normalized
IPSC-plateau values determined in five experiments are presented
before () and +480 min after () the injection of G423. denotes values from control, noninjected neurons. Statistical analysis
was performed by using two-way ANOVA followed by the Tukey-Kramer
test. *p < 0.05 antibody versus noninjected or
before injection. n.s, Not significant.
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DISCUSSION |
Our data show that after injection of specific anti-synapsin
antibodies or purified synapsin, several forms of short-term plasticity
are strongly modified. This raises the questions of whether our
observations can be ascribed to an effect on the synapsin-dependent reserve pool or to alteration in SV trafficking and how antibodies interfere with synapsin function.
Post-tetanic potentiation is believed to involve activity-dependent
recruitment of reserve SVs (for review, see Zucker, 1996, 1999). The
implication of synapsin in PTP has remained an unclear issue: decreased
PTP, concomitant with long-lasting depression, has been found in
synapsin II or synapsin I/synapsin II but not synapsin I mutant mice
(Rosahl et al., 1995). Because we found that PTP amplitude increases
after synapsin injection and is virtually abolished after synapsin
neutralization, our findings unambiguously implicate the synapsins as a
major determinant for this form of synaptic plasticity. The expression
of PTP is generally believed to involve a fast recruitment of SVs from
the reserve pool to the releasable pool (Zucker, 1996, 1999) that can
be achieved, at least in part, through
Ca2+-dependent phosphorylation of synapsin
(Fig. 11, step 5) (Greengard et al., 1993). However, the enhancement of PTP amplitude after injection of dephosphorylated synapsin and the relative long delay for
the appearance of the changes in PTP expression suggest that antibody-
or synapsin-dependent variations in the size of SV pools are involved.
In fact, in recent years the synapsins have been suggested to have a
structural role by favoring the assembly of the reserve pool of SVs
during synaptogenesis and regulating its size in mature neurons (Lu et
al., 1992; Li et al., 1995; Pieribone et al., 1995; Rosahl et al.,
1995; Takei et al., 1995; Valtorta et al., 1995; Ryan et al., 1996;
Hilfiker et al., 1998; Fiumara et al., 2001). Thus, it is possible that
the synapsin-induced enhancement of PTP results from an increase in the
size of the reserve pool at steady state (Fig. 11, box A,
balance between steps 5 and 6) that, in
turn, will increase the supply of reserve SVs to the readily releasable
pool in response to a conditioning stimulus (Fig. 11, step
5), whereas the antibody would elicit the opposite effect by
inducing depletion of the synapsin-dependent reserve pool of SVs.
Moreover, the ability of exogenous synapsin to increase PTP indicates
that the mechanism by which synapsins recruit SVs to the reserve pool
is not saturated at physiological synapsin concentrations.
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Figure 11.
Synapsin-dependent regulation of SV pools and
trafficking during short-term plasticity. Numbers refer
to the following steps in the release process: 1,
recycling of fused SVs; 2, refilling of free release
sites or docking; 3, priming of docked SVs;
4, SV fusion; 5, SV trafficking between
synapsin-dependent reserve pool and membrane-bound pool;
6, sequestration of recycled SV in the
synapsin-dependent reserve pool. Synapsin molecules are shown as
black ribbons; open arrows denote
proposed synapsin-dependent steps. Box A highlights the
putative mechanisms involved in the expression of PTP and depression at
low rate of stimulation. Box B highlights the putative
mechanisms involved in the occurrence of depression at a high rate of
stimulation and synchronization of release. For more details, see
Discussion.
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Several of our data show that after synapsin neutralization, synaptic
efficacy cannot be maintained during and after repetitive stimulations
elicited at time intervals ranging from the millisecond to the second
time scale. This manifests by the appearance of increased depression,
decreased facilitation, and PTD. As detailed below, depending on the
stimulation rate, the underlying synapsin-dependent mechanisms altered
by antibody injection seem to be different.
An experimental situation to be considered is the depression induced by
sustained 0.5 Hz stimulation (Fig. 10). After synapsin neutralization,
it is characterized by both accelerated kinetics and no change in the
plateau level as compared with controls. At active synapses, when the
rate of stimulation is increased over the SV cycle rate (Fig. 11, steps
1-4) (~10-30 sec), synaptic efficacy
depresses and reaches a level that is determined both by the
refilling-recycling capacity and by the concomitant recruitment of
reserve SVs (Ryan et al., 1993; Stevens and Tsujimoto, 1995; Dobrunz
and Stevens, 1997). Because the reserve pool is limited, when
repetitive stimulations are prolonged, transmitter release is likely to
reach a level determined entirely by the recycling capacity. In view of
this model, the unchanged plateau level that characterizes 0.5 Hz
depression indicates that at moderate stimulation frequency, synapsin
does not affect the rate of SV cycling (Fig. 11, steps
1-4). This is consistent with the
unchanged endocytosis and repriming rates observed in synapsin
knock-out mice (Ryan et al., 1996). The marked effects of synapsin
neutralization during the early phase of 0.5 Hz-depression, as well as
the appearance of PTD, are compatible with the notion that synapsin
controls either the size of the reserve pool (Fig. 11, step
6) or the supply of reserve SVs to the releasable
pool to minimize depression (Fig. 11, step 5). However, we
cannot exclude the possibility that 0.5 Hz-depression is also
contributed to by the alteration of a synapsin function that operates
at the membrane level, as discussed below for fast depression (Fig. 11,
step 3). After synapsin neutralization, we found that ACh
release returns faster to its initial level when the stimulation rate
is slowed from 0.5 to 0.025 Hz. This is reminiscent of the faster
replenishment observed at inhibitory synapses in mice lacking synapsin
I (Terada et al., 1999). A likely possibility is that when the
stimulation rate is slowed down, the amount of recycled SVs (determined
by the preceding higher rate stimulation period) exceeds the
replenishment need during a time window in the order of the recycling
time. In intact neurons, these extra SVs are sequestered in the reserve
pool (Fig. 11, step 6). After synapsin
neutralization, this process is no longer possible, and thereby all
recycled SVs may directly enter the releasable pool (Fig. 11, step
2), leading to faster recovery of synaptic efficacy to its
initial level. Obviously, the synapsin-dependent mechanisms involved in
minimizing the depression induced by moderate stimulation rates share
several similarities with PTP mechanisms (Fig. 11, box
A).
A different experimental situation is represented by the fast synaptic
depression induced by synapsin neutralization in Aplysia synapses subjected to short trains of high-frequency (10-50 Hz) stimulation. A disruption of the reserve pool and the subsequent impaired refilling of the releasable pool is unlikely to be involved because the fast kinetics of depression indicates that the SVs involved
should be made available for exocytosis in a time scale (10-100 msec)
that is significantly shorter than the estimated time (several seconds)
needed for reserve SVs to replenish the releasable pool (Stevens and
Tsujimoto, 1995). Taking into account an average residency time of SVs
at the presynaptic membrane of 2-5 sec (Klingauf et al., 1998; Murthy
and Stevens, 1999), this suggests that fast depression results from
disruption of a synapsin action that operates during a membrane step of
the release process (Fig. 11, box B). At variance with
previous deductions made in mice lacking synapsin I (Rosahl et al.,
1995), our data do not support the possibility that this
synapsin-dependent membrane action is related to facilitation
mechanisms. Indeed, although paired-pulse facilitation is depressed
after synapsin neutralization under control conditions (Fig. 9), it is
not affected under low Ca2+ conditions
that maximize facilitation and minimize depression. Most likely,
synapsin plays a modulatory role in the mechanism by which
membrane-bound SVs attain fusion competence or fuse after stimulation
(Fig. 11, steps 3-4). Because fast
depression develops only when the release probability and stimulation
rates are high (Fig. 8), the membrane action of synapsin becomes
limiting under conditions of exhaustion of the releasable pool. The
fact that these regulatory actions of synapsin are not enhanced after
injection of exogenous synapsin suggests that, at variance with PTP,
the involved molecular event(s) may already be saturated by the levels of endogenous synapsin.
The implication of synapsin at a plasma membrane step of the
exocytotic process is also supported by the slower IPSC rise time
induced by anti-synapsin antibodies (Fig. 2). This effect is similar to
that |
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ABSTRACT
INTRODUCTION
