Division of Biology, University of California, San Diego, La Jolla, California 92093-0366
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ABSTRACT |
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Hagler, Donald J., Jr. and
Yukiko Goda.
Properties of Synchronous and Asynchronous Release During Pulse
Train Depression in Cultured Hippocampal Neurons.
J. Neurophysiol. 85: 2324-2334, 2001.
Neurotransmitter release displays at least two kinetically distinct
components in response to a single action potential. The majority of
release occurs synchronously with action-potential-triggered Ca2+ influx; however, delayed releasealso
called asynchronous release
persists for tens of milliseconds
following the peak Ca2+ transient. In response to
trains of action potentials, synchronous release eventually declines,
whereas asynchronous release often progressively increases, an effect
that is primarily attributed to the buildup of intracellular
Ca2+ during repetitive stimulation. The precise
relationship between synchronous and asynchronous release remains
unclear at central synapses. To gain better insight into the mechanisms
that regulate neurotransmitter release, we systematically characterized
the two components of release during repetitive stimulation at
excitatory autaptic hippocampal synapses formed in culture.
Manipulations that increase the Ca2+ influx
triggered by an action potential
elevation of extracellular Ca2+ or bath application of tetraethylammonium
(TEA)
accelerated the progressive decrease in synchronous release
(peak excitatory postsynaptic current amplitude) and concomitantly
increased asynchronous release. When intracellular
Ca2+ was buffered by extracellular application of
EGTA-AM, initial depression of synchronous release was equal to or
greater than control; however, it quickly reached a plateau without
further depression. In contrast, asynchronous release was largely
abolished in EGTA-AM. The total charge transfer following each
pulse
accounting for both synchronous and asynchronous
release
reached a steady-state level that was similar between control
and EGTA-AM. A portion of the decreased synchronous release in control
conditions therefore was matched by a higher level of asynchronous
release. We also examined the relative changes in synchronous and
asynchronous release during repetitive stimulation under conditions
that highly favor asynchronous release by substituting extracellular
Ca2+ with Sr2+. Initially,
asynchronous release was twofold greater in Sr2+.
By the end of the train, the difference was ~50%; consequently, the
total release per pulse during the plateau phase was slightly larger in
Sr2+ compared with Ca2+. We
thus conclude that while asynchronous release
like synchronous release
is limited by vesicle availability, it may be able to access a
slightly larger subset of the readily releasable pool. Our results are
consistent with the view that during repetitive stimulation, the
elevation of asynchronous release depletes the vesicles immediately
available for release, resulting in depression of synchronous release.
This implies that both forms of release share a small pool of
immediately releasable vesicles, which is being constantly depleted and
refilled during repetitive stimulation.
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INTRODUCTION |
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A hallmark of neurotransmitter release
is its exquisite temporal regulation. Arrival of an action potential
triggers Ca2+-dependent exocytosis of synaptic
vesicles within less than a millisecond. Phasic (or synchronous)
neurotransmitter release is central to proper brain function as
evidenced by early postnatal death of mice deficient in synaptotagmin I
whose phasic release is severely compromised (Geppert et al.
1994). When synaptic transmission is monitored from many
synapses, synchronous release of neurotransmitters is followed by a
delayed release that also exhibits Ca2+
dependence (Barrett and Stevens 1972
; Goda and
Stevens 1994
; Meiri and Rahamimoff 1972
;
Miledi 1966
). Such asynchronous release is thought to be
sustained by residual Ca2+ in the presynaptic
terminal. Consistently, asynchronous release is enhanced during
conditions that elevate intracellular Ca2+ as
observed in the course of repetitive stimulation. Asynchronous release
is thus likely to significantly contribute to synaptic transmission in
vivo where neuronal activity mostly occurs as spike trains
(Hubel 1959
; O'Keefe and Dostrovsky
1971
). The physiological relevance of asynchronous release,
however, remains to be established.
During repetitive stimulation, the increase in asynchronous release is
accompanied by a progressive decline in the size of synchronous
release. This decrease in synaptic transmission, which we refer to as
pulse train depression (PTD), has been attributed to occur primarily by
depleting a pool of release-ready synaptic vesicles (Dobrunz and
Stevens 1997; Liley and North 1953
;
Rosenmund and Stevens 1996
). Supporting the hypothesis
that depletion of the readily releasable pool (RRP) causes PTD,
conditions that promote synaptic vesicle fusion by increasing
presynaptic Ca2+ influx increase the rate of PTD
(Tsodyks and Markram 1997
; Varela et al.
1997
). Recent studies indicate that presynaptic mechanisms other than vesicle depletion may also contribute significantly to
synaptic depression. Accordingly, in cultured hippocampal neurons, repetitive stimulation releases only ~77% of the RRP, the size of
which is determined by the application of hypertonic solution (Rosenmund and Stevens 1996
). In addition, models of
depression that take into account depletion only cannot completely
match the properties of experimentally derived PTD data (Dittman
and Regehr 1998
; Matveev and Wang 2000
).
Potential presynaptic mechanisms include inhibition by presynaptic
autoreceptors (Scanziani et al. 1997
; Takahashi
et al. 1996
), Ca2+-channel inactivation
(Forsythe et al. 1998
; Patil et al.
1998
), action potential conduction failures (Brody and
Yue 2000
; Hatt and Smith 1976
; Streit et
al. 1992
), and modulation of
Ca2+-sensitive exocytosis machinery
(Bellingham and Walmsley 1999
; Hsu et al.
1996
; Wu and Borst 1999
).
The purpose of this study is to investigate the relation between
synchronous and asynchronous release during PTD and how PTD affects the
number of readily releasable vesicles in autaptic excitatory synapses
formed in cultured hippocampal neurons. We demonstrate that during
repetitive stimulation the decline of synchronous release is matched by
increased asynchronous release. In addition, our findings are
consistent with recent models of depletion and refilling that are based
on a small subset of the RRPcomprised of on average one or fewer
vesicles
which we term the "immediately releasable pool" (IRP).
Our results suggest that this IRP is shared by synchronous and
asynchronous release; thus depletion by elevated asynchronous release
can reduce future synchronous release.
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METHODS |
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Unless otherwise noted, all chemicals are from Sigma (St. Louis, MO).
Hippocampal neuronal cultures
Cultured hippocampal neurons were prepared from P0 to P2 rats as
previously described with a few modifications (Bekkers and Stevens 1991; Segal and Furshpan 1990
).
Dissection solution consisted of Hank's balanced salt solution (Life
Technologies, Gaithersburg, MD) supplemented with 25 mM HEPES, pH 7.35. The dentate gyrus was removed, and the remaining hippocampal tissue was
proteolyzed in dissection solution plus 20 U/ml papain (Worthington,
Freehold, NJ), 0.5 mM EDTA, 1.5 mM CaCl2, 1 mM
L-cysteine, and 0.1 µg/ml DNAase
for 30 min at 37°.
Tissue pieces were triturated in culture media (CM)
Basal Media Eagle
(Life Technologies) plus 1 mM HEPES pH 7.35, 1 mM Na-pyruvate, 50 U/ml
penicillin, 50 µg/ml streptomycin, 6 mg/ml glucose, 10% fetal bovine
serum (Hyclone, Logan, UT or Omega Scientific, Tarzana, CA), mito serum
extender (Fisher Scientific, Pittsburgh, PA), and 2% B27 (Life
Technologies)
and 0.5 ml of cell suspension was plated at 1-2 × 104 cells/ml onto 12-mm coverslips that had been
preplated with astrocytes. One day after plating, 4 µM cytosine
-D-arabinofuranoside (araC) was added to prevent
astrocyte proliferation. Every 3-4 days thereafter, cells were fed
with 0.1 ml of CM. Astrocytes were prepared from extra cells obtained
from each dissection. After a week, the confluent glial cells were
shaken at 260 rpm overnight to enrich for type I astrocytes. Usually,
the cells were then passaged and grown for an additional week to select
for adherent, quickly spreading astrocytes. Astrocytes were plated at 6 × 103/ml onto coverslips sprayed with microdots
of collagen and poly-D-lysine. After 2-4 days, 4 µM araC
was added to limit the size of the astrocyte islands. Neurons were
added 3-6 days after astrocyte plating, after aspirating most of the
media. Cultures were used for experiments 8-14 days after plating.
Recording solutions
Medium Ca2+ extracellular bath solution
(MC-EBS) contained (in mM) 135 NaCl, 5 KCl, 3 CaCl2, 2 MgCl2, 10 D-glucose, 5 HEPES-NaOH (pH 7.3), and 0.1 picrotoxin plus 1 µM glycine. Osmolarity was adjusted with sorbitol to 315-325 mOsm.
2-Amino-5-phosphonopentanoic acid [(±)APV, 50 µM] was included to
prevent long-term plasticity. High Ca2+ EBS (HC)
was modified MC-EBS with 10 mM CaCl2 and 0.5 mM
MgCl2. Low Ca2+ EBS (LC)
contained 1 mM CaCl2 and 4 mM
MgCl2. The recording chamber was perfused with a
constant flow rate of 0.5-1 ml/min. Pipette solution contained (in mM)
127.7 K-gluconate, 16.4 KCl, 8.4 NaCl, 2.9 MgCl2,
9.4 HEPES-KOH (pH 7.2), 0.2 EGTA, 2 ATP, 0.5 GTP, and 10 creatine
phosphate and 25 U/ml creatine phosphokinase. Drugs were prepared as
stock solutions at the following concentrations in the indicated
solvents: EGTA-AM, 5-nitro-2-(3-phenyl-propylamino) benzoic acid
(NPPB), and cyclothiazide (CTZ) at 100 mM in DMSO, (S)--methyl-4-carboxyphenylglycine (MCPG) at 50 mM in 0.1 N NaOH, apamin at 1 mM in 50 mM acetic acid, and TEA
(tetraethylammonium) at 1 M in H2O. Final
dilutions in EBS are stated in the text.
Data acquisition
Standard whole cell procedures were followed. Recordings were
obtained with Axopatch 200B (Axon Instruments, Foster City, CA),
filtered at 2 kHz and digitized at 2-5 kHz. Traces were stored and
analyzed with programs written by DH using Visual BASIC 5.0 (Microsoft,
Redmond, WA) and ComponentWorks 1.1 (National Instruments, Austin, TX).
Neurons were held at 70 mV; series resistance was compensated to
80%. Leak currents were not subtracted and recordings with a leak
>0.2 nA were excluded. There was at least a 45-s delay between trains
of 20 stimuli, which consisted of 1-ms step depolarizations to 30 mV.
All experiments were performed at room temperature (22-25°C).
Analysis of synchronous release
Excitatory postsynaptic current (EPSC) amplitudes were measured
by subtracting an average baseline value1-10 ms before each pulse
from the peak following the stimulus artifact. For cells in
HC-EBS, the mean initial EPSC amplitude was 6.7 ± 1.2 (SE) nA,
ranging from 0.5 to 31.5 nA, with a standard deviation of 7.6 and a
median of 3.4 nA (n = 39). No relationship between the rate of PTD and the initial peak EPSC amplitude was found (not shown).
If short-term potentiation was observed in a given cell, responses were
included only after the initial peak EPSC amplitude stabilized
(typically after a single pulse train). Recovery of synchronous release
following PTD was assayed by applying a train of 20 pulses followed by
a test pulse at varying time intervals. Recovery was calculated by
normalizing the amplitude of the test EPSC to the amplitude of the
first EPSC of the depressing train. To adjust for incomplete depression
(average depression to 9 ± 1% of initial EPSC amplitude,
n = 7), the amplitude of the last EPSC of the train was
subtracted from both values before normalization.
Analysis of asynchronous and total release
Asynchronous and total release were estimated by integrating the
current in the appropriate time window following each pulse 40-50 ms
for asynchronous and 5 ms to immediately before the next pulse for
total release. A baseline current measured before the first pulse of
the train was subtracted from current traces before integration. Total
release was normalized to the first response, similarly to synchronous
release. Asynchronous release was normalized to the last response as
asynchronous release on the first pulse was small in most cases. The
asynchronous fraction of release
the relative contribution of
asynchronous release to total release
for responses to 20-Hz
stimulation, which is shown in Fig. 5F, was approximated by
dividing the average current 40-50 ms after each pulse by the average
current 5-50 ms after each pulse.
It should be noted that our method of estimating the amount of
asynchronous releaseby measuring charge transfer
has some limitations. First, a portion of the measured currents could be due to
presynaptic currents, which are expected to be insensitive to EGTA-AM.
EGTA-AM, however, greatly reduces the measured currents (see Figs. 6
and 10), suggesting that the presynaptic currents do not contribute
significantly to the estimate of asynchronous release. A second concern
is that accumulation of extracellular glutamate and desensitization of
glutamate receptors may affect the size of the measured currents. The
contribution of these phenomena to the measurement of asynchronous
release is difficult to estimate, and so we compared our method with a
straightforward count of individual asynchronous release events. In a
cell with an initial EPSC amplitude of 1 nA, in medium
Ca2+
i.e., under conditions in which individual
miniature EPSCs (mEPSCs) can be resolved
we find that the charge
transfer and number of mEPSCs increases with matching time courses
during the pulse train and that the charge transfer per release event
closely matches the average charge transfer measured for well-isolated
individual releases (data not shown).
Measurement of hypertonic-evoked response
Hypertonic solution was applied as previously described
(Stevens and Tsujimoto 1995). A 3-s puff of MC-EBS plus
0.5 M sucrose was applied at the end of a train of 100 pulses at 20 Hz,
and the peak of the hypertonic response (HR) arrived 1 s after the end of the train. Cells in which the peak EPSC amplitude at the end of
the train did not depress to <5% of the initial peak EPSC size were
excluded (1 of 5 cells tested). HR currents were integrated using
200-ms bins; the value from the bin with the largest charge transfer
the peak of the integrated response
was used as a measure of
the RRP (Stevens and Tsujimoto 1995
). Control
measurements of the HR were obtained >60 s before and after each test
HR measurement ("test" HR refers to the HR 1 s after the pulse
train). To correct for possible rundown in these measurements, the test
HR measurement was normalized to the average of the preceding and
subsequent control HR measurement. To correct for the baseline shift
due to residual asynchronous release following the pulse train, the integrated charge of a control pulse train was subtracted from that of
a pulse train plus hypertonic solution application. On average, the
uncorrected test HRs were 76 ± 6% of the control HRs. Following
each hypertonic solution application or control pulse train, there was
at least a 60-s delay before subsequent stimulation to allow for
recovery of the RRP.
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RESULTS |
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Extracellular Ca2+ and frequency dependence of PTD
We first examined the basic properties of the decline of
synchronous release (peak EPSC amplitude) during pulse trains by varying extracellular Ca2+ and stimulation
frequency. The effects of changing extracellular Ca2+and thus release probability
on PTD caused
by repetitive stimulus trains in low (1 mM), medium (3 mM), or high (10 mM) extracellular Ca2+ are shown in Fig.
1A. Relative to high
Ca2+, synchronous release to the first pulse was
decreased to 84 ± 6% (n = 8) in medium
Ca2+ and 34 ± 4% (n = 6)
in low Ca2+. While the rate of PTD was fast in
high Ca2+, PTD was slowed in lower
Ca2+. Moreover, facilitation of the second EPSC
of the train relative to the first was observed in low
Ca2+. When stimulus frequency was varied
5, 10, and 20 Hz
higher stimulus frequency resulted in a faster rate of
depression (Fig. 1, B-D), in agreement with previous
studies (Dittman and Regehr 1998
; Galarreta and
Hestrin 1998
; Janz et al. 1999
; Tsodyks
and Markram 1997
; Varela et al. 1999
). At 20 Hz
in high Ca2+, the EPSC amplitude usually decayed
to zero within 10-20 pulses (Figs. 1A and
2B).
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RRP recovers quickly following complete depression
PTD has been generally interpreted to result from depleting the
RRP of vesicles (Liley and North 1953) (see
INTRODUCTION). The observed increase in the rate of PTD
caused by a higher stimulus frequency and conditions that increase
release probability is consistent with this model. The depletion model
would then predict that the lack of synaptic response observed for PTD
at higher stimulus frequencies reflects the emptied RRP
(Rosenmund and Stevens 1996
). To address whether the RRP
is indeed emptied following PTD, we first applied a prolonged pulse
train (100 pulses) at 20 Hz (in medium Ca2+) to
achieve a fully depressed state, where the synchronous release is
negligible or nonexistent. Immediately after this depression, the size
of the RRP was measured by applying extracellular solution made
hyperosmolar with 0.5 M sucrose (Rosenmund and Stevens
1996
). Surprisingly, 62 ± 6% (n = 4) of
the RRP was available for release just 1 s after the end of the
train, the shortest time at which the RRP can be reliably assayed by
hypertonic solution application (Fig. 2D). The recovery of
RRP was also matched by the recovery of the EPSC amplitude tested at
various times following PTD (Fig. 2D). The refilling of the
RRP has been well characterized as a single exponential recovery with a
time constant ranging from 3 to 11 s (Dobrunz and Stevens
1997
; Pyle et al. 2000
; Rosenmund and
Stevens 1996
; Stevens and Tsujimoto 1995
;
Stevens and Wesseling 1998
). The lower limit of 3 s
is achieved by a Ca2+-dependent acceleration of
refilling that occurs following repetitive stimulation; however, this
effect saturates at ~10 pulses at 10 Hz, such that more pulses or
higher stimulus frequencies produce approximately the same rate of
refilling (Stevens and Wesseling 1998
). Figure
2D, - - -, shows the predicted recovery from an emptied
RRP that would be observed under accelerated conditions. Note that the
observed recovery of the RRP attained within 1 s following PTD is
much faster than the prediction. While this could indicate an
exceptionally accelerated refilling, it is plausible that the fast
recovery results from incomplete depletion of the RRP. In support of
this latter proposal, at the end of a pulse train substantial
asynchronous release occurs even though synchronous release is
completely depressed (see following text). Thus factors other than
depletion likely contribute to the depression of the synchronous EPSC.
Factors that could potentially contribute to PTD
We next addressed mechanisms other than depletion that might
potentially contribute to PTD in cultured hippocampal neurons. To
determine whether desensitization of AMPA-type glutamate receptors plays a role in PTD, repetitive stimulation was applied in the presence
of CTZ to block AMPA-receptor desensitization. CTZ (100 µM) had no
effect on the rate of PTD despite a threefold increase in the initial
EPSC amplitude (Fig. 3A). A
small increase in the normalized amplitude of the second response was
observed in CTZ at 20 Hz; this difference was not observed for 5 or 10 Hz and was not statistically significant (n = 4, P = 0.23, 2-tailed, paired t-test). We also
examined the potential contribution of metabotropic glutamate receptors
in modifying presynaptic glutamate release during repetitive
stimulation (Scanziani et al. 1997; Takahashi et
al. 1996
). Bath application of 250 µM MCPG, a metabotropic glutamate receptor antagonist, was without effect on PTD (Fig. 3B). AMPA receptor desensitization and activation of
metabotropic receptors therefore do not contribute significantly to PTD
in our recording conditions.
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It has been suggested that PTD could arise from action potential
propagation failures at axonal branch points during repetitive stimulation (Brody and Yue 2000). For example,
Ca2+ that builds up during repetitive stimulation
could activate Ca2+-dependent
K+ or Cl
channels.
Ensuing hyperpolarization could then impede action potential
propagation or simply reduce the extent of depolarization at the
synapse (Lüscher et al. 1996
). Such modulation of
action potential propagation could result in a progressive reduction of
EPSC amplitudes. To test this possibility, PTD was measured in the
presence of blockers of Ca2+-dependent
K+ or Cl
channels (Fig.
3, C-E). Apamin (1 µM), which blocks the small conductance K+ channels, had no effect on the
rate of PTD. Nevertheless, 1 mM TEA, which blocks the big conductance
channels, greatly increased the rate of PTD. The TEA-dependent
increase in PTD rate most likely resulted from higher release
probability caused by enhanced Ca2+ influx,
similar to the effects of elevating extracellular
Ca2+ (Katz and Miledi 1966
;
Kusano et al. 1967
) (see following text). NPPB (10 µM), a chloride channel blocker, had no effect on PTD; when applied
at a higher concentration (100 µM), NPPB slightly increased the rate
of PTD (n = 3, data not shown). A reduction in the rate
of PTD was not observed by blocking
Ca2+-dependent K+ or
Cl
channels. Our results, therefore are
inconsistent with activation of Ca2+-dependent
K+ or Cl
channels
contributing to PTD by promoting action potential propagation failures.
Depression of EPSC amplitude depends on accumulation of intracellular Ca2+
The dependence of PTD on the levels of extracellular
Ca2+ and stimulus frequencyboth of which
determine the intracellular Ca2+ concentration
attained during repetitive stimulation (Tank et al.
1995
)
suggests that the accumulation of intracellular
Ca2+ contributes to the depression of synaptic
strength. To test this hypothesis, PTD was measured following bath
application of 100 µM EGTA-AM to chelate intracellular
Ca2+. EGTA-AM caused a slight but significant
decrease in the initial EPSC size (78 ± 5% of control;
P < 0.002, 2-tailed, paired t-test; Fig.
10C), presumably because of the slight reduction in peak
Ca2+ concentration brought about this slow
chelator (Atluri and Regehr 1996
). In contrast to
responses recorded in control condition, which gradually decreased
toward zero, EGTA-AM greatly attenuated the extent of PTD, most
noticeably later in the train when responses reached a plateau (Fig.
4E). Furthermore, the stimulus
frequency and extracellular Ca2+ dependence of
PTD were greatly reduced in the presence of EGTA-AM (compare Figs. 1
and 4, A-D). A buildup of intracellular
Ca2+ during repetitive stimulation is thus likely
to play a role in the mechanism of PTD.
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Decrease in synchronous release is matched by increase in asynchronous release
We next characterized the properties of asynchronous release
during repetitive stimulation in relation to the depression of synchronous release. Asynchronous release was examined by measuring late releasethat which follows the decay of synchronous release
by integrating the current between 40 and 50 ms after each pulse (see
METHODS). As expected from its Ca2+
dependence, more asynchronous release was initially observed in high
extracellular Ca2+ (Fig. 5,
A-C). Elevated
Ca2+, as well as higher stimulus frequency, also
promoted a faster rise in asynchronous release during the pulse train.
Once a maximal level was reached, asynchronous release remained
constant or even declined slightly. This plateau level was higher with
increased stimulus frequency or Ca2+
concentration (Fig. 5, A-C).
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We then examined the relative changes in synchronous and asynchronous
release with increasing stimulus number. The fractional contribution of
asynchronous release to the total releasesum of synchronous and
asynchronous release (see METHODS)
was initially minimal
at each extracellular Ca2+ concentration tested.
In low Ca2+ the contribution of asynchronous
release increased gradually during the pulse train (Fig.
5F). In high Ca2+, however, there was
a complete or nearly complete shift to asynchronous release, especially
at higher stimulus frequency (Fig. 5F). Despite the
differences in the relative contributions of synchronous and asynchronous release at different extracellular
Ca2+, at higher stimulus frequency the total
release per pulse tended to eventually reach the same value, regardless
of extracellular Ca2+ concentration (Fig.
5E).
The sensitivity of asynchronous release to intracellular Ca2+ is illustrated in Fig. 6. Bath application of EGTA-AM largely blocked the rise in asynchronous release that normally develops during repetitive stimulation (Figs. 6A and 10C). In contrast, synchronous release reached a higher plateau in EGTA-AM relative to control (Fig. 4E). To determine whether the sustained synchronous release observed in EGTA-AM can be accounted for by the reduction in asynchronous release, we compared the total release per pulse before and after EGTA-AM treatment. In high extracellular Ca2+, by the end of the train total release is approximately equal between that measured with or without EGTA-AM despite greatly reduced synchronous release in control conditions (Figs. 4E and 6C). This demonstrates that part of the decrease in synchronous release during a pulse train is accompanied by a matching increase in asynchronous release.
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Effect on PTD of increasing asynchronous release with extracellular Sr2+
The preceding results suggest that depression of synchronous
release involves a shift to asynchronous release. To examine how
tightly coupled these two forms of release are, we substituted extracellular Ca2+ with
Sr2+, a manipulation that increases asynchronous
release while decreasing synchronous release.
Sr2+ substitution of high
extracellular Ca2+ decreased the initial EPSC
amplitude by half while increasing the initial asynchronous release by
about twofold (Figs. 7, A and
B, and 10A). At 20 Hz, the rate of depression of
the EPSC amplitude was similar in Ca2+ and
Sr2+. At 5 and 10 Hz, however, the initial
depression on the second pulse (paired-pulse depression: PPD) was
significantly greater in
Sr2+ than in
Ca2+ (extent of depression was 71 ± 11 and
21 ± 5% greater for 5 and 10 Hz, respectively, n = 6, P < 0.01 for both, 2-tailed, paired t-test). As in the presence of Ca2+,
asynchronous release reached a plateau for each of these frequencies tested; however, the plateau levels reached in
Sr2+ were somewhat higher than in
Ca2+ (for example, in Fig. 7B, 20-Hz
plateau was 45 ± 15% greater in
Sr2+, n = 6, P < 0.03, 2-tailed, paired t-test). As a
result, by the end of the train, the total release evoked per pulse was
greater in Sr2+ than in
Ca2+by 31 ± 5% at 20 Hz (Fig.
7C: P < 0.01, 2-tailed, paired
t-test) and 48 ± 17% at 5 Hz (Fig. 7F:
P < 0.04, 2-tailed, paired t-test).
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We also examined the effects of Sr2+ substitution
of medium Ca2+. The initial EPSC amplitude was
decreased to one-fifth of the original size, and the initial
asynchronous release was slightly elevated. Although PPD was observed
at 10 and 20 Hz in Ca2+, we found facilitation at
those frequencies in Sr2+ (Fig.
8, A and D),
consistent with a reduction in release probability in
Sr2+ (Xu-Friedman and Regehr
2000). Subsequent PTD was largely unaffected (compare Figs.
1C and 8D). In contrast to
Sr2+ replacement of high
Ca2+, the increase in asynchronous release with
Sr2+ replacement of medium
Ca2+ was not sufficient to cause a greater total
release per pulse (Fig. 8, C and F). At 20-Hz
stimulation, the differences in asynchronous and total release between
Ca2+ and Sr2+ became very
small by the end of the train.
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Effect on PTD of prolonged Ca2+ influx
While testing for a possible contribution of
Ca2+-dependent K+ channels,
we found that TEA caused a large increase in the rate of PTD. This
result is readily explained by the broadening of the action potential
caused by blockade of K+ channels and the
subsequent increase in Ca2+ influx (Katz
and Miledi 1966; Kusano et al. 1967
). TEA
increased both the initial synchronous and asynchronous release (Figs.
9, A and B, and
10B). Total release per
pulse was in general greater in TEA; however, at 20 Hz this difference
quickly diminished (Fig. 9, C and F). As
discussed in the preceding text, the plateau level of asynchronous
release later in the pulse train increases with higher stimulus
frequency (Fig. 5A). In the presence of TEA, the frequency
dependence of the plateau reached by asynchronous release was greatly
reduced (Fig. 9E). In addition, TEA caused an increase in
the maximal asynchronous release (Fig. 9B). During 20-Hz
stimulation, asynchronous release was 49 ± 17% greater in TEA on
the 10th pulse (n = 5, P < 0.05, 2-tailed, paired t-test) but only 23 ± 17% greater on
the 20th pulse (P > 0.25, 2-tailed, paired
t-test).
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DISCUSSION |
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We studied the properties of synchronous and asynchronous release
during repetitive stimulation to gain insight into the underlying mechanisms of PTD and the pool of vesicles that supplies the two components of release. In view of its extracellular
Ca2+ and frequency dependence, the progressive
decline in synchronous release is consistent with depletion of
available vesicles. Nevertheless, RRP recovery following complete
depression of synchronous release was faster than the expected
refilling of an empty pool, suggesting that the RRP is not fully
depleted during PTD. Thus it is plausible that depletion of the RRP is
not the only source of depression of synchronous release. We have also
shown that synchronous and asynchronous releases are coordinately
regulated such that manipulations that increase the rate of decline of
synchronous release elevate asynchronous release. The sum of
synchronous and asynchronous release was similar between control and
EGTA-AM, despite the differential effects of EGTA-AM on the two
components of release. We use these data to advance a model describing
an immediately releasable vesicle which suppliesand is depleted
by
both forms of release.
Possible factors governing depression of synchronous release
Since PTD likely results from the combination of
several processes, we have sought to rule out several potential
mechanisms to simplify the analysis of neurotransmitter release
properties during PTD. We have ruled out desensitization of AMPA
receptors because CTZ has no significant effect on PTD under the
present experimental conditions as reported for several other
preparations (Bellingham and Walmsley 1999;
Dittman and Regehr 1998
; Dobrunz and Stevens
1997
; Galarreta and Hestrin 1998
; Varela
et al. 1997
). Metabotropic glutamate receptors do not
contribute to PTD as an antagonist for these receptors had no effect,
similar to a previous demonstration in hippocampal slices
(Dobrunz and Stevens 1997
). Furthermore, earlier work
showed that PPD was not affected by MCPG in hippocampal cultures
(Maki et al. 1995
). We were unable to test the role of
Ca2+ channel inactivation in PTD; however, such a
role is expected to be minor since the fractional reduction in
Ca2+ currents is relatively small compared with
the extent of PTD (Forsythe et al. 1998
). Furthermore,
"closed-state" Ca2+ channel inactivation is
not sensitive to buffering of Ca2+ (Patil
et al. 1998
) unlike the PTD observed here.
Another potential mechanism contributing to PTD that deserves attention
is alteration of action potential (AP) propagation; this could include
a reduction in AP amplitude or actual AP conduction failure at axonal
branch points (Brody and Yue 2000; Hatt and Smith
1976
; Streit et al. 1992
). If AP amplitude is
progressively reduced, or if the frequency of AP conduction failures
increases with repetitive stimulation, it is expected that the EPSC
amplitude would decrease during a pulse train. Although AP conduction
in hippocampal axons is highly reliable, even for pairs of pulses (Mackenzie and Murphy 1998
; Stevens and Wang
1995
), a reduction in AP amplitude in the second of a pair of
stimuli has been reported with a recovery time constant of ~8 ms
(Brody and Yue 2000
). Such a fast recovery rate should
preclude significant effects on PTD at the frequencies tested in the
current study. Nonetheless, a potential mechanism for reducing AP
amplitude is activation of Ca2+-dependent
K+ or Cl
channels
(Lüscher et al. 1996
). A blockade of these
channels might be expected to decrease the rate of PTD. PTD, however,
was either unaffected or increased, suggesting that this type of
mechanism is unlikely to be involved in the EGTA-sensitive component of PTD. Presently, we cannot completely exclude the possible contribution to PTD by channels that are insensitive to the drugs tested. Finally, it has also been proposed that high extracellular
Ca2+ promotes conduction failure by reduced
excitability resulting from charge screening effects (Brody and
Yue 2000
). That the Ca2+ dependence of
PTD is sensitive to buffering intracellular Ca2+
with EGTA demonstrates that changes in membrane excitability by charge
screening effects is not a significant issue.
Coordinate regulation of synchronous and asynchronous release
We find that manipulations that increase
Ca2+ influx accelerate both the depression of
synchronous release and the increase in asynchronous release with
subsequent stimulus pulses. We also observe that the sum of synchronous
and asynchronous release at the end of the train was similar
between control and EGTA-AM. These results indicate an interesting
scheme whereby elevating asynchronous release depresses synchronous
release by depleting the vesicles immediately available for synchronous
release. Such a proposal implies that the two forms of release share a
common pool of releasable vesicles. Our observations are similar to
those reported recently for inhibitory synapses in nucleus
magnocellularis where the enhanced contribution of asynchronous release
during high-frequency synaptic transmission may play a significant role in auditory processing (Lu and Trussell 2000). The fact
that interdependent regulation of synchronous and asynchronous release
also occurs at hippocampal excitatory synapses suggests that it might
be a general property expressed by all central synapses.
It is of interest to note that our results differ from a previous study
in cultured hippocampal neurons in which synchronous release monitored
in the presence of EGTA-AM did not reach a significantly higher plateau
at the end of the stimulus train compared with control (Cummings
et al. 1996). Likely reasons for the apparent disagreement are
the lower range of extracellular
Ca2+ levels and shorter stimulus
trains used in the earlier study.
Model for a common pool supplying synchronous and asynchronous release
Based on the coordinate regulation of the two components of release we propose that synchronous and asynchronous release share a common pool of releasable vesicles. We describe a model that has been refined within the framework of our observations and those made by others.
RELEASE OCCURS FROM AN IRP CONSISTING OF A SINGLE QUANTUM.
Studies of transmitter release properties at single central synapses
indicate that neurotransmitter release is limited to at most a single
quantum (Arancio et al. 1994; Edwards et al. 1976
; Stevens and Wang 1995
). A plausible
explanation for the release of just a single vesicle is that the IRP,
which supplies synchronous release, represents a small subset of the
RRP consisting of a single, most primed vesicle (Matveev and
Wang 2000
). In our model, synchronous release occurs
exclusively from the IRP, and asynchronous release can also gain access
to the IRP (see Fig. 11). This model
accounts for features of short-term plasticity of synchronous release.
An IRP that is limited to one vesicle will produce prominent PPD under
conditions of high release probability as found in this and other
studies (Fig. 1) (Brenowitz et al. 1998
; Dittman
and Regehr 1998
; Thomson 1997
; Varela et
al. 1997
). Paired-pulse facilitation occurs with low initial
release probability if, at rest, the fraction of synapses with an
occupied IRP is less than one and refilling accelerates upon
stimulation. Our model accounts for the increased PPD of synchronous
release
in particular for 5- and 10-Hz stimulation
in high
extracellular Sr2+. Although it would be expected
that greatly reduced initial release probability should result in less
depression, we observe increased PPD (compare Figs. 1D and
7D). Assuming asynchronous release can also gain access to
the IRP, elevated levels of asynchronous release would deplete the IRP,
thereby preventing that synapse from later releasing synchronously with
the action potential (Fesce 1999
).
|
MECHANISMS UNDERLYING ADDITIONAL INCREASES IN ASYNCHRONOUS
RELEASE.
We observed that while asynchronous release increases to match a
portion of the depression of synchronous release, asynchronous release
could be increased further in the presence of TEA or by extracellular
substitution of Ca2+ by
Sr2+. Both TEA and Sr2+ are
expected to greatly increase the residual divalent concentrations; TEA
because of prolonged Ca2+ influx (Katz and
Miledi 1966; Kusano et al. 1967
), and
Sr2+ because of slower clearance
(Xu-Friedman and Regehr 1999
). This suggests that while
synchronous release may be limited to a single, most primed vesicle
(IRP), asynchronous release can, under special conditions
such as in
the presence of TEA or high
Sr2+
access a slightly larger
pool. We suggest two alternative mechanisms to account for this greater
access. In the first scenario, asynchronous release can access vesicles
other than the IRP in the RRP. Such non-IRP vesicles normally have a
much lower affinity for Ca2+ or
Sr2+ and require much greater concentrations to
undergo fusion. Asynchronous release, therefore can access these
vesicles under the extreme conditions achieved with TEA or
Sr2+. In the second case, very high
Ca2+ or Sr2+ further
accelerate the rate of priming, thereby increasing the occupancy of the
IRP. In this mechanism, synchronous and asynchronous release share the
identical pool of releasable vesicles. In either case, the implication
of our model is that under normal conditions the IRP is the primary
supplier of vesicles for both synchronous and asynchronous release.
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ACKNOWLEDGMENTS |
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We thank Dr. Silvio Varon for generously providing equipment support, G. Ramirez for excellent technical assistance, and Dr. Marla Feller and members of the Goda lab for helpful comments on the manuscript.
This work was supported by grants from the National Institute of Mental Health (MH-57710) and the Whitehall Foundation to Y. Goda. D. J. Hagler was supported by a training grant from the National Institutes of Health.
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FOOTNOTES |
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Address for reprint requests: Y. Goda, Div. of Biology, University of California, San Diego, 9500 Gilman Dr., La Jolla, CA 92093-0366 (E-mail: ygoda{at}biomail.ucsd.edu).
Received 17 October 2000; accepted in final form 13 February 2001.
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REFERENCES |
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