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The Journal of Neuroscience, January 15, 2001, 21(2):462-476
Quantitative Relationship between Transmitter Release and Calcium
Current at the Calyx of Held Synapse
Takeshi
Sakaba and
Erwin
Neher
Max-Planck-Institute for Biophysical Chemistry, Department of
Membrane Biophysics, D-37077, Göttingen, Germany
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ABSTRACT |
A newly developed deconvolution method (Neher and Sakaba,
2001
) allowed us to resolve the time course of neurotransmitter release at the calyx of Held synapse and to quantify some basic aspects
of transmitter release. First, we identified a readily releasable pool
(RRP) of synaptic vesicles. We found that the size of the RRP, when
tested with trains of strong stimuli, was constant regardless of the
exact stimulus patterns, if stimuli were confined to a time interval of
~60 msec. For longer-lasting stimulus patterns, recruitment of new
vesicles to the RRP made a substantial contribution to the total
release. Second, the cooperativity of transmitter release as a function
of Ca2+ current was estimated to be 3-4, which
confirmed previous results (Borst and Sakmann, 1999; Wu et al., 1999).
Third, an initial small Ca2+ influx increased the
efficiency of Ca2+ currents in subsequent
transmitter release. This type of facilitation was blocked by a high
concentration of EGTA (0.5 mM). Fourth, the release rates
of synaptic vesicles at this synapse turned out to be heterogeneous:
once a highly Ca2+-sensitive population of vesicles
was consumed, the remaining vesicles released at lower rates. These
components of release were more clearly separated in the presence of
0.5 mM EGTA, which prevented the buildup of residual
Ca2+. Conversely, raising the extracellular
Ca2+ concentration facilitated the slower population
such that its release characteristics became more similar to those of
the faster population under standard conditions. Heterogeneous release
probabilities are expected to support the maintenance of synaptic
transmission during high-frequency stimulation.
Key words:
transmitter release; synaptic depression; facilitation; release probability; vesicle pool; the calyx of Held
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INTRODUCTION |
Knowledge of the kinetics of
transmitter release and its dependence on
Ca2+ influx is important for elucidating
the molecular mechanisms mediating exocytosis from presynaptic
terminals. In particular, short-term plastic changes, such as synaptic
facilitation and depression, are likely to have presynaptic components,
the quantification of which require accurate knowledge of release
rates. Until now, the time course of transmitter release in most
mammalian synapses has been assayed mainly by analyzing postsynaptic
responses. In the neuromuscular junction, deconvolution of the EPSCs
with the miniature EPSC (mEPSC) gives an accurate estimate of the time course of transmitter release (van der Kloot 1988a,b), because the
released quanta do not interact with each other (Hartzell et al., 1975;
Magleby and Pallotta, 1981). However, this assumption cannot always be
made in central synapses. Postsynaptic factors, such as desensitization
(Trussell et al., 1993; Otis et al., 1996) or saturation of receptors
(Jonas et al., 1993; Tang et al., 1994; Auger et al., 1998), and
delayed clearance of the released transmitter from the synaptic cleft
(Barbour et al., 1994; Mennerick and Zorumski, 1995; Silver et al.,
1996) can also contribute to the time course of the EPSC. All of these
mechanisms prevent accurate characterization of the presynaptic factors
mediating the EPSC. Thus, deconvolution has so far been applied to only
a limited number of central synapses (Borges et al., 1995; Diamond and
Jahr, 1995).
To separate the presynaptic and postsynaptic factors that determine
the time course of the EPSC, we have developed a novel type of
deconvolution (Neher and Sakaba, 2001). In this method, we modeled the
residual current component caused by the delayed clearance of glutamate
from the synaptic cleft and incorporated this current into the
deconvolution algorithm. The validity of the method was tested using
fluctuation analysis and is described in detail in a recent paper
(Neher and Sakaba, 2001).
At the calyx of Held synapse, an axosomatic synapse in the brainstem,
it is possible to clamp the presynaptic terminal and the postsynaptic
target simultaneously (Borst et al., 1995; Takahashi et al., 1996),
allowing precise timing and quantification of the presynaptic
Ca2+ influx, as well as the measurement of
EPSCs at high time resolution. Basic aspects of transmitter release,
such as the cooperativity of transmitter release and its dependence on
extracellular Ca2+ concentration (Borst
and Sakmann, 1996, 1999a; Takahashi et al., 1996; Wu et al., 1999), as
well as the size of the readily releasable pool (RRP) of synaptic
vesicles (Schneggenburger et al., 1999; Wu and Borst, 1999), have
already been investigated at this synapse. However, possible
postsynaptic factors affecting the EPSCs were not taken into account.
Using our new deconvolution method, we have been able to analyze the
effects of these postsynaptic mechanisms and found that both receptor
desensitization and saturation can readily occur during intense
stimulation protocols. However, in the presence of cyclothiazide (CTZ)
and kynurenic acid (Kyn), both effects are reduced to the point that
they are no longer a serious cause of error in deconvolution (Neher and
Sakaba, 2001). Using these drugs, we now present data which show that
the size of the RRP is constant regardless of the stimulus patterns
used and also much larger than previously estimated. Additionally, we
find that the release probabilities of synaptic vesicles are heterogeneous. Possible mechanisms and biological implications of these
findings are discussed.
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MATERIALS AND METHODS |
Presynaptic and postsynaptic recordings at the calyx of Held
were performed in slice preparations of the rat brainstem at room
temperature (21-24°C) as described elsewhere (Neher and Sakaba, 2001). Briefly, a slice was transferred to the recording chamber and
perfused continuously with normal saline at a rate of ~1 ml/min. Normal saline contained (in mM): NaCl 125, KCl 2.5, CaCl2 2.0, MgCl2, 1.0, glucose 25, NaHCO3 1.25, ascorbic acid 0.4, myo-inositol 3, and Na-pyruvate 2, pH 7.3-7.4, 320 mOsm, and was
bubbled continuously with 95% O2 and 5%
CO2.
The presynaptic and postsynaptic compartments were simultaneously
clamped at a holding potential of 
80 mV (unless noted otherwise). The
presynaptic pipette (4-7 M
) was filled with a solution containing (in mM): Cs-gluconate 125-130, TEA-Cl 20, HEPES 10, Na2 phosphocreatine 5, MgATP 4, GTP 0.3, BAPTA 0.05, pH
7.2, 310 mOsm. In some experiments the concentration of BAPTA was
either increased to 0.2 mM or replaced by EGTA (0.05 or 0.5 mM). The postsynaptic pipette (2-4 M) was filled with
the same solution as the presynaptic pipette, except that BAPTA was
replaced by 5 mM EGTA.
During recordings, 0.5-1 µM TTX, 10 mM
TEA-Cl, and 50 µM
D-2-amino-4-phosphonobutyric acid (D-AP5) were
added to the normal saline to isolate the presynaptic calcium current
and to block NMDA receptors. In some cases, 0.1 mM
3,4-diaminopyridine was also added to block the presynaptic
K+ current. Bicuculine (10 µM) and strychnine (2 µM) were added in
some recordings to block possible inhibitory input. To vary the
extracellular calcium concentration, the CaCl2
concentration of the normal saline was increased to 10 mM.
In some recordings, 2 or 10 mM CaCl2
was added to a solution containing (in mM): NaCl 150, KCl 2.5, HEPES 10, and glucose 10, pH 7.3-7.4, 320 mOsm. In
most experiments, Kyn (1 mM) and CTZ (100 µM) were added, because it was shown that desensitization
and saturation of postsynaptic receptors can be avoided only in
the presence of these drugs (Neher and Sakaba, 2001). CTZ, Kyn,
D-AP5, and
2,3-dioxo-6-nitro-1,2,3,4,-tetrahydrobenzoquinoxaline-7-sulfonamide (NBQX) were purchased from Tocris (Köln, Germany). Bicuculine, strychnine, 3,4-diaminopyridine, and TEA were from Sigma (Deisenhofen, Germany). TTX was from Alamone Labs (Jerusalem, Israel).
Both presynaptic and postsynaptic cells were whole-cell-clamped with an
EPC9/2 amplifier controlled by the Pulse program (HEKA-Electronik, Lambrecht, Germany) to a holding potential of 80 mV. Currents were
low-pass-filtered at 2.9 or 6 kHz and stored at 10 or 20 kHz. The
presynaptic series resistance (Rs;
8-35 M, typically 15 M) was compensated 30-70%. Subtraction of
the presynaptic capacitive current was made with the P/5 protocol or by
scaling the capacitive current by a small voltage pulse. This
correction was applied for the calculation of plateau inward currents
during depolarizations. However, the original traces in the figures
show uncorrected records. Presynaptic leak currents of >200 pA
typically caused rundown of the postsynaptic response; therefore,
recordings from cells with such leakage were not used. The interval
between the two stimuli was 20-30 sec. The postsynaptic series
resistance (3-10 M, typically 5 M) was compensated so that the
uncompensated series resistance was ~2-3 M. Residual resistance
was compensated off-line by correcting for the deviation from the
holding potential (=Rs × IEPSC). Postsynaptic leak currents
were typically 50-100 pA. Recordings were stopped once the
uncompensated postsynaptic Rs became
>10 M or the postsynaptic leak current was >300 pA. Most of the
data that had postsynaptic Rs of >8
M were rejected later during analysis because power spectra showed
that the high-frequency signal is reduced by high
Rs.
Deconvolution of postsynaptic current records was performed as
described by Neher and Sakaba (2001). This procedure calculates release
rates 
(t) in vesicles released per millisecond. A
first-order estimate of the total size of the RRP can be obtained by
integrating the release rate between the beginning of a stimulus
protocol and the end of a depleting stimulus (all of our stimulus
protocols are concluded by such depleting stimuli; see below). However, this integral includes vesicles that have been newly recruited to the
RRP while the stimulus protocol proceeds. To correct for this effect,
we applied an iterative procedure based on the simplest possible pool
model, which is a homogeneous pool of
Nt vesicles, when completely filled,
and recovering exponentially with a time constant 
, when fully or
partially depleted. Designating n(t) as the pool
size at a given time t, this model can be represented by the
rate equation:
![<FR><NU>dn(t)</NU><DE>dt</DE></FR>=<UP>−</UP>&xgr;(t)+[N<SUB><UP>t</UP></SUB>−n(t)]/&tgr;.](/content/vol21/issue2/fulltext/462/img001.gif)
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(1)
|
This can be integrated to yield:
|
(2)
|
As initial guesses Nt,0 for
Nt we use:
|
(3)
|
where t1 is the time at which
the depleting pulse ends. The initial value
n0(t) for
n(t) is calculated according to:
|
(4)
|
With these values inserted into the right side of Equation 2, we
obtain an improved estimate
n1(t), from which we
calculate an improved estimate for Nt,
according to:
|
(5)
|
Here, we calculate a weighted average of new and previous
estimates to avoid oscillation in the iteration. Repeating these steps
iteratively two to five times results in stable values for Nt,v and
nv(t), which are then taken
as the initial pool size, and the number of remaining vesicles at a
given time t, respectively. The quantity "release rate per
vesicle", pv (see below), was calculated
according to:
|
(6)
|
with nv(t)
calculated according to Equation 2 using a stable value of
Nt,v (in place of
Nt).
This procedure clearly is an oversimplification because it has been
shown that recovery of the pool is not described by a single
exponential (Wu and Borst, 1999). However, all of our protocols are
short (<120 msec) relative to the fast time constant of recovery of Wu
and Borst (1999) (200-300 msec). Therefore, it is only the initial
rate of recovery that is of relevance for the calculation. We represent
this, for simplicity, by a single time constant.
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RESULTS |
Presynaptic effects of CTZ
In our experiments, we used CTZ to block desensitization of
postsynaptic AMPA receptors and to slow the decay of EPSCs. However, it
has been suggested that CTZ may act presynaptically to enhance transmitter release (Barnes-Davies and Forsythe, 1995; Diamond and
Jahr, 1995; Isaacson and Walmsley, 1996; Bellingham and Walmsley, 1999). In a preceding paper, we have shown that AMPA receptors desensitize strongly during synaptic transmission in the absence of CTZ
(Neher and Sakaba, 2001). Therefore, the enhancement of the synaptic
response seen in the presence of CTZ is most likely caused by block of
the desensitization of postsynaptic receptors rather than an increase
in transmitter release.
To further explore this question, we also examined the effects of CTZ
on NMDA receptor-mediated EPSCs (NMDA EPSCs), which have been used for
testing the presynaptic effects of this drug (Trussell et al., 1993;
Diamond and Jahr, 1995; Bellingham and Walmsley, 1999). The NMDA EPSC
was isolated by application of 1 µM NBQX and 10 µM glycine, whereas the postsynaptic neuron was held at
either 40 or +60 mV. Strychnine (2 µM) was also
applied to avoid responses to glycine. Trains of pulses
(depolarization to 20 mV for 2 msec with an interpulse interval of 6 msec) were applied to the presynaptic terminal. Responses to pulses did
not change significantly after the addition of CTZ (100 µM) (Fig. 1A,B).
Specifically, the EPSC amplitude in the presence of CTZ was ~90% of
the control EPSC amplitude for both the first (0.93 ± 0.07) and
second (0.91 ± 0.04) pulse (n = 5 cell pairs).
Also, the paired-pulse ratio of the EPSCs (0.99 ± 0.08) did not
change substantially in the presence of CTZ (Fig.
1B). Therefore, we found no evidence for presynaptic
effects mediated by CTZ in our experimental conditions. However, we
cannot entirely exclude the possibility that CTZ modulates transmitter
release under current-clamp conditions, for example, by modulating ion
channels, instead of modulating the release machinery.
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Figure 1.
The effect of cyclothiazide (CTZ)
on the NMDA EPSC. A, The presynaptic terminal was
depolarized from 80 to +80 mV and repetitively shifted back to +20 mV
every 8 msec for 2 msec. After repeating this stimulus six times, the
terminal was held at 0 mV for 10 msec
(VPre). The Ca2+
currents (middle) and the NMDA EPSCs
(bottom) before and after the application of CTZ (100 µM) were superimposed. The NMDA EPSC was isolated by
application of 1 µM NBQX and 10 µM glycine,
and the postsynaptic neuron was held at +60 mV. Strychnine (2 µM) was also applied to avoid responses to glycine.
B, Left to right, The EPSC
amplitude evoked by the first pulse (1st pulse), the
second pulse (2nd pulse), the EPSC amplitude achieved at
the end of the stimulus protocol (total response), and
the paired-pulse ratio [ratio between the second pulse and the first
pulse (2nd/1st)]. In all cases,
quantities indicated were measured both before and after the
application of CTZ. Ratios of these values before and after the
application of CTZ are plotted (n = 5 cell
pairs).
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Transmitter release rates estimated from deconvolution
In the presence of CTZ, a large current component of the EPSC (the
residual current component) arises from the delayed clearance of
glutamate from the synaptic cleft. Therefore, simple deconvolution of
the EPSC with the mEPSC overestimates release rates. Thus, the residual
current component must be subtracted from the EPSC before deconvolution
can be performed. Also, Kyn, a fast competitive glutamate antagonist,
must be included in the bath solution to avoid receptor saturation and
clamp errors. To calculate the residual current, we used a simple model
of glutamate diffusion [for a detailed description, see Neher and
Sakaba (2001)]. The protocol shown in Figure
2A, referred to as the
"fitting protocol, " was used to determine the free parameters of
this model (n, the exponent of the power law of glutamate
channel activation; rD, the
diffusional distance; and nD, the
exponent of the diffusion law). In the fitting protocol, the
presynaptic terminal was depolarized from 80 to +70 mV to activate
Ca2+ channels maximally. At this potential
there was almost no Ca2+ influx, because
the driving force of Ca2+ was very low.
From +70 mV the voltage of the terminal was shifted to 0 mV several
times. Durations of these repolarizations were adjusted so that the
resulting Ca2+ influx evoked EPSCs that
varied several-fold in amplitude. The EPSCs decayed during episodes
when the presynaptic terminal was held at +70 mV. The rate of quantal
release calculated by variance analysis was <10
msec
1
during such decay phases of EPSCs (Neher and Sakaba, 2001), suggesting low rates of asynchronous release. Because the EPSC decay is largely shaped by the delayed clearance of glutamate and not by asynchronous release, we could set parameters of the glutamate diffusion model from
the decay phases of the EPSCs (Fig. 2A, dotted
line close to the EPSC trace). Parameters of the diffusion model
were determined for each cell pair using a fitting protocol like that
described in Figure 2A. Then, deconvolution was
performed after subtracting the residual current component (Fig.
2A). Once parameters were determined, they were used
for other traces in a given cell pair.
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Figure 2.
The deconvolution method to estimate quantal
release rates at the calyx of Held. A, The fitting
protocol to determine parameters of the glutamate diffusion model. The
presynaptic voltage (VPre) was shifted
from 80 to +70 mV and then repolarized to 0 mV (1-10 msec in
duration) several times. IPre denotes
presynaptic voltage-clamp current, which represents mainly
Ca2+ current, not corrected for leak and capacitance
current. Durations of these repolarizations were adjusted so that the
size of EPSCs varied several-fold. Filtered variance
(variance, solid line) was used to
estimate release rates during the EPSC decay (Neher and Sakaba,
2001). The dotted line in the variance trace indicates
an estimate of AMPA receptor channel noise (Neher and Sakaba, 2001).
The difference between the total variance and channel noise should
arise from fluctuations associated with quantal release. Parameters of
the glutamte diffusion model, which accounts for the residual current
caused by delayed clearance of glutamate (Neher and Sakaba, 2001), were
determined from EPSC decay phases. After subtracting the residual
current component (dotted line on the EPSC
trace) from the total EPSCs, release rates (release
rate, bottom) were calculated by deconvolution
of the remaining EPSC with the mEPSC. The presynaptic patch
pipette contained 0.05 mM BAPTA. B,
Release rates during trains of pulses. The presynaptic terminal was
depolarized from 80 to +70 mV and repetitively shifted back to 5 mV
every 10 msec for 4 msec. After repeating this stimulus six times, the
terminal was held at 5 mV for 20 msec to deplete the RRP. Release
rates were calculated after subtracting the residual current component
(dotted line in the EPSC trace) from the
total EPSC. Variance associated with EPSCs and channel variance
(dotted line) are also shown. The data were obtained
from the same cell pair as shown in A.
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To investigate the kinetics of transmitter release, we applied trains
of pulses as shown in Figure 2B. The presynaptic
terminal was depolarized to +70 mV to activate
Ca2+ channels fully while keeping
Ca2+ influx minimal. The terminal was then
repetitively shifted back to a fixed potential every 10 msec (in the
case of Fig. 2B, to 5 mV) for 4 msec to allow a
finite amount of Ca2+ influx. After six
such stimuli, the terminal was held at 5 mV for 20 msec to deplete
completely the RRP of synaptic vesicles (hence, the name "depleting
pulse"). We did not see a significant inactivation of the presynaptic
Ca2+ current in the protocol that we used
(Forsythe et al., 1998; but see Borst and Sakmann, 1999b) (Fig.
2B, Ipre). Compared
with the first pulse, the Ca2+ current
amplitudes during the second and third pulses were 100 ± 1% and
97 ± 1%, respectively. We did not see facilitation of the
Ca2+ current as has been reported
previously (Borst and Sakmann, 1998; Cuttle et al., 1998). This is
because Ca2+ channels were activated
maximally in our protocols, whereas facilitation of
Ca2+ current was seen only when
Ca2+ channels were partially activated
(Borst and Sakmann, 1998; Cuttle et al., 1998).
The release rate was calculated by deconvolution as described above
(Fig. 2B). It increased during the first episode of
presynaptic Ca2+ influx and decreased
immediately after cessation of the Ca2+
influx. Both the EPSC and the peak release rate were depressed during
successive stimuli, suggesting that the first pulse released most of
the RRP. Furthermore, the depleting pulse at the end of the protocol
evoked almost no release (Fig. 2B). Variance analysis also indicated that the release rate at the end of a depleting pulse is
quite low (~6
msec1)
(Neher and Sakaba, 2001).
A readily releasable pool of synaptic vesicles at the calyx
of Held synapse
Using this deconvolution method, we were also able to estimate the
size of the RRP of vesicles. Our expectation is that a constant number
of vesicles should be released by a sufficiently strong stimulus,
regardless of the stimulus pattern, if pool depletion is the main
mechanism of synaptic depression (Neher, 1998a; but see Hsu et al.,
1996). A protocol similar to that of Figure 2B was
used, except that the magnitude of the repolarization was varied in
each train of pulses to elicit varying amplitudes of Ca2+ current. Because the terminal was
depolarized first to +70 mV, the half-time of the
Ca2+ current rise during repolarization
was the same regardless of the level of repolarization. At the end of
all stimulus protcols, the presynaptic terminal was held at 5 mV for
20 msec to deplete the RRP completely (Fig.
3A,
VPre).
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Figure 3.
Relationship between release rate and
Ca2+ influx. A, A protocol similar to
that described in Figure 2B was used, except that
pulse durations were kept constant and the magnitude of repolarization
was varied in each train of pulses, thereby varying amounts of
Ca2+ influx. Four sweeps were superimposed.
Different colors represent the different sweeps in the
traces for VPre,
IPre, EPSC, and the
Release Rate. The data were obtained from the same cell
pair as shown in Figure 2. Details of the presynaptic inward
currents evoked by the first pulse in each record are shown in the
inset between the two bottom-most traces. These records
have not been corrected for leak and capacitive transients (see
Materials and Methods). B, The number of vesicles
released during the stimulation protocol was calculated by integrating
the release rate. The release rate was integrated from the end of the
depleting pulse to the beginning of the protocol. Therefore, this
figure shows the number of remaining vesicles. We assumed that new
synaptic vesicles were recruited to the RRP with a time constant of 1 sec, calculating the integral according to Equation 2. The individual
traces correspond to those of A, using the same
color code. C, The release rates per
vesicle during the first three pulses, calculated according to Equation 5, are shown. The traces correspond to those of
A, using the same colors.
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In cases of larger Ca2+ influxes (for
example, the green trace in Fig. 3A), the peak
release rate during the initial stimulus was large, and the release
rate was depressed during subsequent stimuli. In contrast, during
smaller initial Ca2+ influxes (for
example, the red trace in Fig. 3A), the peak
release rate during the initial stimulus was very small, and the
release rate was facilitated by subsequent stimuli (Fig.
3A). For all stimulus patterns, the release rate declined to
a low value at the end of the depleting pulse. The number of vesicles
released during stimulation was calculated by integrating the release
rate and considering also partial pool refilling during the stimulus, as described in Materials and Methods (Fig. 3B). A constant
number of vesicles (~2000) were released during all stimulus
patterns. Although the number varied among different cell pairs (from
800 to 4400), the number was almost constant within a single cell pair.
The average number of releasable vesicles from 11 cell pairs was
2408 ± 309.
The above estimation assumes that recovery of the RRP of synaptic
vesicles occurs with a time constant of 1 sec (see Materials and
Methods). This value comes from experiments in which double depolarizations to 10 mV for 20 msec were applied to the presynaptic terminal with an interpulse interval of 500 msec. Each pulse was strong
enough to deplete completely the RRP of synaptic vesicles. The number
of vesicles released by the second pulse was 47% (41-49%; n = 3 cell pairs) of the number released by the first
pulse. Thus, the entire RRP of synaptic vesicles should be refilled in
~1 sec assuming that refilling proceeds at a constant rate for all
vesicles. However, it has been shown that a subset of vesicles of the
RRP at the calyx synapse recovers faster (Wu and Borst 1999).
Therefore, we also estimated the size of the RRP assuming a much faster
recovery time constant. To determine this faster recovery time
constant, we examined the release rate at the end of the depleting
pulse and during a short time interval after the depleting pulse. We found that the release rate approached values of 10 msec1
(9.8 ± 1.9 msec1;
n = 11 cell pairs; see also Fig. 5A). This
value should reflect the rate at which (after depletion) new vesicles
are recruited to the RRP and immediately released, because the
Ca2+ concentrations after strong
depolarization are expected to be high (Wu and Borst, 1999).
Integrating this rate for individual experiments yields recovery time
constants between 200 and 1000 msec (typically 200400 msec), similar
to values found in a previous study (Wu and Borst, 1999). Using
200400 msec as a time constant, we recalculated the size of the RRP
to be 2246 ± 303 vesicles (n = 11 cell pairs)
with pool size estimates from individual traces fluctuating typically
by ±10% around the mean pool size of a given cell pair. Thus, the
effect of pool recovery turned out to be minor, probably because the
stimulation protocol was relatively short (<100 msec) compared with
the recovery time constant. For the following analyses, we assumed fast
recovery ( = 200400 msec). However, the results did not
differ significantly when the recovery time constant was assumed to be
1 sec.
The pool size that we estimated was larger than previous estimates
(600-800 vesicles) (Schneggenburger et al., 1999; Wu and Borst, 1999).
These previous studies assumed that the size of the mEPSC was constant
under the experimental conditions examined. However, this assumption is
usually not warranted at the calyx of Held, because postsynaptic AMPA
receptors desensitize in the absence of CTZ and saturate in the
presence of CTZ, unless a fast blocker of glutamate binding such as Kyn
is added (Neher and Sakaba, 2001). Also, an appreciable fraction of
release may occur desynchronized during bursts of strong stimuli, such
that the cumulative amplitude of EPSCs is not a good measure of pool
size. It should be noted that our pool size estimate depends on the
assumption that under Kyn, mEPSC size is reduced by the same factor
(0.5×) as the macroscopic EPSC. Recordings in the presence of Kyn show
that mEPSCs are clearly reduced. Unfortunately, an accurate measurement
of mEPSC amplitude is not possible, because the smallest mEPSCs are
lost in noise. Our pool size estimate would be proportionally smaller
if mEPSC amplitude under Kyn were larger than 0.5 times the control value.
Relationship between Ca2+ influx and the release
rate per vesicle
If the observed synaptic depression was caused simply by the
depletion of a homogeneous RRP, then the release rate per vesicle (pv) should remain constant at constant
Ca2+ influx or even increase because of
facilitation. We calculated pv by determining
the pool size (see above), calculating the cumulative release as a
function of time, and then dividing the release rate () by the
number of remaining synaptic vesicles (see Materials and Methods). For
a homogeneous pool, for which the release rate is expected to decay
exponentially, this rate per vesicle pv is
expected to be constant and equal to the rate constant (or the inverse
of the time constant) of the release process (Burrone and Lagnado,
2000). We analyzed the traces shown in Figure 3 and calculated
pv (Fig. 3C). It is seen that for large Ca2+ currents,
pv decreased with successive pulses, despite
the fact that the Ca2+ current amplitude
stayed constant. For a homogeneous vesicle pool, we would instead
expect an increase in pv because of
facilitation; however, this was observed only for weak stimuli in which
the first pulse elicited little release. In each cell pair, the peak release rates during the first three pulses were plotted against the
presynaptic Ca2+ current amplitude
(ICa) (Fig.
4A), and the
relationship was fitted with a Hill function:
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(7)
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or a power function:
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(8)
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when the relationship did not show saturation. The release rate
per vesicle during first pulses could be fitted with a third or fourth
power function of Ca2+ current (Eq. 8)
(n = 3.69 ± 0.36; five cell pairs). Corresponding values for the second and third pulses, however, showed saturation at
larger calcium influxes, and the release rates were lower than those
observed during the first pulses when the amplitude of
Ca2+ current was >1.5 nA. In contrast,
pv showed facilitation during the second and
third pulses when Ca2+ influxes were
smaller (<1 nA of Ca2+ current). Similar
trends were apparent in data pooled from different cell pairs (Fig.
4B). pv values during the
second and third pulses could be fit empirically with a Hill
coefficient (n) of 34 (second pulse: n = 3.66 ± 0.11; third pulse n = 3.07 ± 0.44;
five cell pairs), and max values during second
and third pulses were 0.21 ± 0.02 msec1 and
0.19 ± 0.03 msec1,
respectively (five cell pairs). Third pulses had slightly smaller exponents. However, because the relationship exceeds the
supralinear range, third pulses can also be fitted with the same
exponent as the first and second pulses. Specifically,
pv decreased from an initial value of
0.28 ± 0.05 msec1
during the first pulse to 0.20 ± 0.03 msec1 and
0.16 ± 0.08 msec1
during second and third pulses, respectively, when the amplitude of the
Ca2+ current was relatively large
(1460 ± 80 pA). In contrast, facilitation of the rate (0.1-0.2
msec1) was
observed during smaller Ca2+ influxes
(when the Ca2+ current was <1 nA) (Fig.
4B).
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Figure 4.
Relationship between Ca2+
influx and the peak release rate per vesicle. A, The
peak release rates per vesicle during the first (+), second ( ), and
third pulses ( ), calculated from the data shown in Figure 3, are
plotted against peak Ca2+ current. The data could be
fitted with a Hill function (Eq. 7). First pulses were fitted with
max = 0.76 msec1, K = 1320 pA, n = 3.19. Second pulses were fitted with
max = 0.35 msec1, K = 686 pA, n = 3.31. Third pulses were fitted with
max = 0.34 msec1, K = 704 pA, n = 2.36. B, Summary of the
relationship between the peak release rate per vesicle during the first
(continuous line), second (broken line),
and third (dotted line) pulses and the amplitude of
Ca2+ current (n = 7 cell pairs).
Presynaptic patch pipettes contained 0.05 mM BAPTA.
C, Summary of the relationships between the peak release
rate per vesicle and the amplitude of Ca2+ current
during the first (continuous line), second
(broken line), and third (dotted line)
pulses. The presynaptic patch pipette contained 0.05 mM
EGTA.
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We also examined the relationship between pv
and the Ca2+ influx in the presence of
0.05 mM EGTA (Fig. 4C) (four cell pairs). Qualitatively, results similar to those obtained in the presence of
0.05 mM BAPTA were observed. The relationship
between the peak release rate during the first pulses and
Ca2+ current was fitted with a power
relationship of n = 2.81 ± 0.62, which seemed to
be slightly lower in the presence of 0.05 mM EGTA in comparison to 0.05 mM BAPTA. Peak
pv during the second and third pulses could be
fitted with Hill coefficients of 2.94 ± 0.18 and 2.28 ± 0.49 and maximal rates of 0.17 ± 0.02 and 0.21 ± 0.03 (msec1),
respectively. Slight differences in cooperativity (n)
between 0.05 mM BAPTA and 0.05 mM EGTA are most likely because of the increased
ability of BAPTA to be saturated by short
Ca2+ influxes because of its faster
Ca2+ binding kinetics. Faster binding
increases the apparent cooperativity, as has been demonstrated in a
recent modeling study (Bertram et al., 1999).
These results indicate that the release probabilities of synaptic
vesicles are heterogeneous; decreases in peak release rates per vesicle
during second and third pulses cannot be explained by changes in the
Ca2+ stimulus, because peak
[Ca2+] values during second and third
pulses are probably larger than those during first pulses, because of
residual Ca2+ and, possibly, saturation of
Ca2+ buffers. Rather, the observed
differences are probably a consequence of preferred depletion of a
subset of synaptic vesicles that has a high probability of release.
Separating fast- and slow-releasing synaptic vesicle pools
by Ca2+ chelators
The dependence of secretion rates on
Ca2+ influx is complicated (Fig. 4),
because facilitation and preferred depletion of a subset of synaptic
vesicles with high release probability occur simultaneously. Because it
has been demonstrated in other synapses that exogenous
Ca2+ chelators block facilitation by
reducing residual Ca2+ (Kamiya and Zucker,
1994; Atluri and Regehr, 1996), it is expected that heterogeneous
release processes may be observed more clearly in the presence of an
exogenous Ca2+ chelator. Therefore, we
examined vesicular release rates in the presence of exogenous
Ca2+ chelators introduced via the patch
pipette. Specifically, we compared the effects of the slow
Ca2+ chelator EGTA with those of the fast
Ca2+ chelator BAPTA (Adler et al., 1991).
Concentrations of 0.5 mM EGTA and 0.2 mM BAPTA
were used, because the introduction of higher concentrations made it
impossible to fully deplete the RRP, such that
pv could not be calculated (data not shown).
We will refer to experiments using 0.5 mM EGTA or 0.2 mM BAPTA as performed in high buffering conditions.
In contrast to the case of low Ca2+
buffering (control conditions in the presence of 0.05 mM
EGTA and 0.05 mM BAPTA), facilitation of both the EPSC and
the release rate was largely blocked during low
Ca2+ influxes (<1 nA of the
Ca2+ current) in the presence of 0.5 mM EGTA (Fig.
5A,B).
At larger Ca2+ influxes,
pv decreased with successive pulses (Fig.
5A,B). In all protocols, the
release rate dropped to values on the order of 10 msec1 at
the end of the depleting pulse. This suggests that the RRP was
completely depleted (Fig. 5A). pv
values are plotted against the amplitude of the
Ca2+ current in Figure 5B. On
average, pv of the first pulse is very similar
to that obtained in low buffering conditions (Fig. 5C), and
the power relation was fit with n = 2.84 ± 0.31. However, facilitation during the second and third pulses was almost
entirely blocked in the presence of elevated EGTA (Fig.
5B,C). For four cell pairs studied,
the relationship between pv and
Ca2+ current during second and third
pulses (Fig. 5B,C) could be fitted empirically with a Hill coefficient (n) of 2.74 ± 0.35 and 2.30 ± 0.08 and max of 0.17 ± 0.03 and 0.135 ± 0.01 msec1,
respectively. At larger Ca2+ influxes,
pv during second and third pulses approached
saturation and was lower than that of first pulses. Averaging among
four cell pairs (Fig. 5C), pv
values at a mean Ca2+ current amplitude of
1522 ± 60 pA were 0.26 ± 0.06 msec1
(first pulse), 0.11 ± 0.003 msec1
(second pulse), and 0.08 ± 0.01 msec1
(third pulse). As has been described before,
pv values during second and third pulses were
twice as large (second pulse: 0.2 msec1;
third pulse: 0.16 msec1) in
low buffering conditions (0.05 mM BAPTA), when
the mean amplitude of Ca2+ current was
comparable (1460 pA). Thus, 0.5 mM EGTA appears
to slow down the release of all vesicles by reducing residual
Ca2+ and blocking facilitation, regardless
of whether vesicles have lower or higher release probabilities.
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Figure 5.
The effect of 0.5 mM EGTA on release
rates per vesicle. A, The same protocols as described in
Figure 3 were used. However, the presynaptic patch pipette contained
0.5 mM EGTA. Release rates observed during the depleting
pulse are shown at two different magnifications. Note that the
Ca2+ tail current at the end of the depleting pulse
(indicated by a short artifact in the inset) does not
lead to any changes in release rate. B, The relationship
between the peak release rate per vesicle and the amplitude of the
Ca2+ current. The data are from the cell pair shown
in A. The peak values of release rate per vesicle during
the first (continuous line), second (broken
line), and third (dotted line) pulses were
plotted against the amplitude of Ca2+ current. First
pulses could be fitted with a power relationship (Eq. 8) with
n = 2.43. Second and third pulses were fitted with
a Hill function (second pulse: max = 0.12 msec1; K = 739 pA; n = 3.42; third pulse:
max = 0.13 msec1; K = 204 pA; n = 0.916). C, Summary of
the relationship between the peak release rate per vesicle during
the first (continuous line), second (broken
line), and third (dotted line) pulses and the
amplitude of Ca2+ current. The presynaptic patch
pipette contained 0.5 mM EGTA (n = 4 cell pairs).
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Because BAPTA binds to Ca2+ (4.5 × 108
M1/sec1)
much faster than EGTA (2.7 × 106
M1
sec1)
(Adler et al., 1991; Naraghi, 1997), BAPTA should more strongly affect
transmitter release. The same stimulus protocols as shown in Figures 3
and 5 were used to examine the relationship between pv and Ca2+
influx in the presence of 0.2 mM BAPTA (Fig.
6) (three cell pairs). With this higher
concentration of BAPTA, pvs during the first pulse were reduced to <0.05
msec1 when
the amplitude of Ca2+ current was ~1 nA.
In contrast, pv values of >0.1
msec1 were
obtained both during low buffering conditions (Fig. 4) and under high
EGTA concentrations (Fig. 5C). Thus, 0.2 mM BAPTA was effective in blocking exocytosis of
the subset of synaptic vesicles with higher release probabilities. This
inhibition seems to be partially overcome by larger
Ca2+ influxes, when current amplitude was
>1 nA (Fig. 6). Interestingly, facilitation of
pv was still observed during second and third pulses when Ca2+ influx was small (Fig.
6). Such findings can be interpreted as "pseudofacilitation," which
occurs because of saturation of the exogenous fast
Ca2+ chelator (Neher, 1998b; Bertram et
al., 1999). This phenomenon has already been studied in detail at
cortical synapses (Rozov et al., 2001).
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Figure 6.
The effect of 0.2 mM BAPTA on release
rates per vesicle. The same stimulus protocols as described in Figure 3
were used, and the peak values of release rate per vesicle during the
first (continuous line), second (broken
line), and third (dotted line) pulses were
plotted against the amplitude of the Ca2+ current
(n = 3 cell pairs).
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Although the protocols described so far readily display facilitation
and the reduction of pv during second and
third pulses, a quantitative interpretation of Figures 4 and 5,
B and C, is somewhat complicated by the fact that
each value of a given curve represents the situation after some
prerelease, which is different between different data points. In the
following experiments, we used different protocols to better
characterize the individual vesicle subsets.
Heterogeneity of transmitter release observed during
long depolarization
Because the presence of EGTA effectively prevents the buildup of
residual Ca2+ during long-lasting stimuli
(Wu and Borst, 1999), we expected that the use of 0.5 mM
EGTA would allow us to resolve distinct components of transmitter
release using long depolarizations of the presynaptic terminal.
Specifically, the terminal was depolarized from 80 to 70 mV for 4 msec, and then the voltage was shifted back to 10 mV for 50 msec to
allow maximum Ca2+ influx (Fig.
7A). During this protocol, the
amplitude of the Ca2+ current was
1634 ± 126 pA (n = 4 cell pairs) and did not show strong inactivation (Forsythe et al., 1998; Borst and Sakmann, 1999b).
More precisely, the Ca2+ current at the
end of the 50 msec pulse was 84.0 ± 2.8% of the Ca2+ current during the first 5 msec.
Release rates were calculated from deconvolution of the evoked EPSC
(Fig. 7B). They increased rapidly after activation of the
Ca2+ current and displayed a biexponential
decay (Fig. 7). After the decay phase, a small amount of release (<10
msec1)
persisted. Again, this release probably reflects the release of both
the remaining releasable vesicles and the newly recruited vesicles.
Therefore, we applied a correction to take into account this refilling
of vesicles to the RRP. Then, the cumulative release was calculated by
integrating the corrected release rate according to Equation 5, and the
fraction of released vesicles was plotted against time (Fig.
7C). The cumulative plot shows two phases of release that
could be fitted with a double exponential. From four cell pairs, the
faster time constant was 2.40 ± 0.55 msec, and the slower one was
20.91 ± 4.7 msec. The fast component contributed 53.0 ± 4.3% to the total fit. However, it is possible that the proportion of
the faster component may be even larger, because the exponential fit
assumes that the slow component begins instantaneously after the
depolarization. Also, our estimates for the low release rates after the
initial decay cannot be considered very accurate in the presence of a
large residual current (Neher and Sakaba, 2001). Even so, the analysis
shows that there is a strong deviation from a monoexponential time
course of transmitter release at this synapse and suggests that there
are at least two classes of vesicles with differing release
probabilities (high and low). Interestingly, the time constants
indicate that one class of vesicles is five to ten times more likely to
release than the other. In this respect, the two components may be
somewhat similar to those observed in hippocampal neurons, which were
identified using the postsynaptic channel blocker MK-801 (Hessler et
al., 1993; Rosenmund et al., 1993). It is also important to mention
that although our data were adequately fitted with a double
exponential, we cannot exclude the presence of more components. Indeed,
the distribution of release probabilities might be continuous, as has
been suggested for other synapses (Dobrunz and Stevens, 1997; Huang and
Stevens, 1997; Isaacson and Hille, 1997; Murthy et al., 1997; Silver et
al., 1998).
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Figure 7.
Time course of quantal release during a
long-lasting depolarization in the presence of 0.5 mM EGTA
in the presynaptic patch pipette. A, The presynaptic
terminal was depolarized from 80 to +70 mV for 4 msec and repolarized
to 10 mV for 50 msec (VPre,
top) to elicit a Ca2+ current
(IPre, middle). The
evoked EPSC (bottom) is shown. The dotted
line in the bottom panel indicates the estimated
residual current component. B, Release rate estimated
from the evoked EPSC is plotted against time. Starting at the time
point of 0, the presynaptic terminal was held at 10 mV.
C, The cumulative fraction of released vesicles plotted
against time. The release rate was integrated after correcting for the
effect of refilling of synaptic vesicles to the RRP and was normalized
to the total pool size. The data could be fitted with a double
exponential, with time constants of 2.01 msec (45%) and 16.64 msec
(dotted line).
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We also performed a similar set of experiments (long depolarization) in
low buffering conditions (0.05 mM BAPTA) (Fig.
8). In such experiments, the cumulative
amount of release plotted against time could not be readily separated
into the two components. In the five cell pairs that we studied, the
faster time constant was 3.17 ± 0.85 msec, and the slower one was
6.55 ± 3.55 msec (Fig. 8C). The faster component
contributed significantly more (76.0 ± 12.5%) to the total fit
than in high buffering conditions, whereas its time constant was
slightly slower. This may well be a consequence of the difficulty in
separating components, combined with restricted time resolution of the
deconvolution procedure. The time constant of the slow component, at a
given Ca2+ current amplitude, was faster.
Contrary to release rates, the amplitudes of the presynaptic
Ca2+ currents (1895 ± 154 pA) were
not substantially different from those of cell pairs in the presence of
0.5 mM EGTA. Similar results were also observed
in the presence of 0.05 mM EGTA (five cell pairs;
data not shown). These results suggest that the accumulation of
presynaptic Ca2+ accelerates the release
of the slow component. Therefore, the two components of release are not
as clearly separated as in the case of 0.5 mM
EGTA, which lowers residual Ca2+.
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Figure 8.
Time course of quantal release during long pulses
in the presence of 0.05 mM BAPTA in the patch pipette.
A, The presynaptic terminal was depolarized from 80 to
+70 mV for 4 msec and was held at 10 mV for 20 msec
(VPre) to elicit presynaptic
Ca2+ current
(IPre). The dotted
line shows an estimate of the residual current component.
B, Release rate plotted against time. Starting at the
time point of 0, the presynaptic terminal was held at 10 mV.
C, The cumulative fraction of vesicles released is
plotted against time. The data could be fitted with double exponentials
with time constants of 1.09 msec (68%) and 5.92 msec (dotted
line).
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Ca2+ dependence of the
slow component
To examine the Ca2+ dependence of the
slow component, we applied depolarizing pulses (test pulse, 10 msec in
duration) of various amplitudes after depleting most of the faster
component by prepulses of a fixed amplitude (depolarization to 10 mV
for 4-6 msec; constant duration within each cell pair). The duration
of the prepulse was adjusted to deplete most of the faster component,
considering that the time constant of the faster component is 2-3 msec
(Fig. 7C). At the end of the protocol, the presynaptic
terminal was held at 10 mV for 20 msec to completely deplete the RRP
(Fig. 9A). Subsequently, a
similar sequence of pulses was applied without prepulses to examine the
dependence of the fast component on Ca2+
influx (Fig. 9B) in the same cell pair.
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Figure 9.
Prepulse experiment for studying the
Ca2+ dependence of the slower component.
A, Depolarizing pulses (test pulse; 10 msec in duration)
of various amplitudes (from 20 to +30 mV) were applied after a fixed
prepulse (depolarization to 10 mV for 5 msec). In all protocols, the
terminal was finally depolarized to 10 mV for 20 msec, to deplete the
RRP. From top, VPre,
IPre, EPSC, and
release rate are shown. B, The same
protocol as described in A except that the prepulse was
omitted. The same cell pair as in A was used.
C, The peak release rate per vesicle during the test
pulse in the presence () or absence (+) of the prepulse, is plotted
against the amplitude of Ca2+ current. The data are
from the same cell pair as in A and B. D, The time-to-peak of the release rate per vesicle
during the test pulse in the presence () or absence (+) of the
prepulse is plotted against the amplitude of Ca2+
current. The data are from the same cell pair as A and
B. Note that the time-to-peak cannot exceed 10 msec,
because the pulse duration is 10 msec.
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Experiments were performed with high concentrations of EGTA (0.5 mM) to allow a clear separation of the fast and the slow components (Figs. 9,
10A,B).
The peak release rate per vesicle (pv) during
the test pulse was plotted against the amplitude of the
Ca2+ current (Fig. 9C; pooled
data in Fig. 10A). The dependence of the release rate
on the Ca2+ current was less steep when
prepulses were present than when prepulses were absent. At 2 nA of
Ca2+ current, pv
during the test pulse in the presence of a prepulse was ~0.1
msec1,
whereas it was >0.2
msec1 in
the absence of the prepulse (Fig. 10A). The
time-to-peak of pv was also measured for each
test pulse and plotted against the amplitude of
Ca2+ current (Figs. 9D,
10B). The time-to-peak was significantly longer when
prepulses were applied. By increasing the
Ca2+ influx, differences between the
time-to-peak between test pulses with and without prepulses became
smaller. These findings seem to suggest that some buildup of
Ca2+ is necessary to activate release of
the slower component.
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Figure 10.
Prepulse experiment in the presence of 0.5 mM EGTA. A, B, The peak
release rate per vesicle (A) and the time-to-peak
of the release rate per vesicle (B) during the
test pulses (Fig. 9) are plotted against the amplitude of
Ca2+ current. Presynaptic patch pipettes contained
0.5 mM EGTA. The data are averages from five cell pairs.
Note that the time-to-peak cannot exceed 10 msec, because the pulse
duration is 10 msec (the same for D). C,
D, The peak release rate per vesicle
(C) and the time-to-peak of the release rate per
vesicle (D) during test pulses are plotted
against the amplitude of Ca2+ current. Presynaptic
patch pipettes contained 0.05 mM BAPTA. The data are
averages from six cell pairs.
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For comparison, the same experiments were performed in low buffering
conditions (0.05 mM BAPTA) (Fig.
10C,D). Then, a more complex behavior was
observed. The pv during the test pulse was facilitated by the prepulse when the amplitude of
Ca2+ current was <800 pA. In contrast,
the peak pv during the test pulse was
depressed by the prepulse when the amplitude of
Ca2+ current became >1 nA. In the
presence of the prepulse and for Ca2+
currents between 1.5 and 2.0 nA, the peak pv
during the test pulse was larger (0.2 msec1) in
the low buffering condition (Fig. 10C) than in high
buffering conditions (0.1 msec1)
(Fig. 10A). These results suggest that accumulation
of presynaptic Ca2+ facilitates the
release of the slow component and that this facilitation is prevented
by higher concentrations of EGTA. The relationship between the
time-to-peak of pv and the amplitude of the
Ca2+ current also supports this idea (Fig.
10D). The time-to-peak during the test pulse in the
presence of the prepulse was the same or even shorter than in the
absence of the prepulse. Again, this is in contrast to findings
under high buffering conditions (Fig. 10B) and
suggests that accumulation of presynaptic
Ca2+ facilitates the release of the slow component.
Recruitment of the slow component by elevating
extracellular Ca2+
The accumulation of presynaptic Ca2+
accelerates the release of the slow component. Then, sufficient
concentrations of presynaptic Ca2+ should
be able to activate release of the slow component with rates comparable
to those of the fast component in normal conditions. We tested this
expectation by elevating the extracellular
Ca2+ concentration to 10 mM
and examining the release rates under low buffering conditions (0.05 mM BAPTA). The presynaptic terminal was depolarized from
80 to +70 mV and repolarized every 10 msec to 10 mV for 4 msec to
allow Ca2+ influx into the terminal (Fig.
11). The average amplitude of the Ca2+ current was 2650 ± 167 pA
during the first pulse (n = 6 cell pairs). Inactivation
of the Ca2+ current (Forsythe et al.,
1998; Borst and Sakmann 1999b) was not prominent with the pulse
protocol used (Fig. 11). The evoked EPSC was depressed strongly during
the second pulse, and the release rate was constant and low during
further stimuli. The ratio of cumulative release evoked by the second
pulse to the first pulse was 0.14 ± 0.02 (n = 6 cell pairs). The time-to-peak of the release rate during the first
pulse was 1.21 ± 0.39 msec, and the rate began to decay during
the pulse (Fig. 11). The decay could be fitted by a single exponential
with a time constant of 1.71 ± 0.72 msec. The peak release rate
per vesicle during the first pulse was 0.51 ± 0.06 msec1,
indicating that most of readily releasable synaptic vesicles could be
depleted within the first 4 msec of depolarization. The result is
consistent with the idea that the slow component can be released quite
rapidly once presynaptic Ca2+
concentrations reach a certain level. Therefore, the slow component will most likely contribute to nerve-evoked release when
Ca2+ concentration builds up during trains
of action potentials.
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Figure 11.
Release rates observed in the presence of 10 mM extracellular Ca2+. Extracellular
Ca2+ was increased to 10 mM to augment
the amplitudes of the Ca2+ currents. The presynaptic
terminal was depolarized from 80 to +70 mV and repolarized every 10 msec to 10 mV for 4 msec. After repeating this stimulus six times,
the terminal was held at 5 mV for 20 msec to deplete the RRP
(VPre). Release rates
(bottom) were calculated from the evoked EPSC.
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DISCUSSION |
The calyx of Held offers unique possibilities for the study of
synaptic transmission by allowing an analysis of the relationship between Ca2+ influx and rates of secretion
under voltage clamp in both presynaptic and postsynaptic compartments
(Borst et al., 1995; Takahashi et al., 1996). A new variant of
deconvolution, which we developed (Neher and Sakaba, 2001), allows a
direct calculation of rates of release in the presence of CTZ and Kyn,
under which quantal size was shown to be constant over a wide range of
stimulation protocols examined (Neher and Sakaba, 2001). Because Kyn is
displaced by glutamate in its sustained presence (Diamond and Jahr,
1997), one may argue that under Kyn, quantal size changes in an
activity-dependent manner, especially at high release rates. This
possibility was not examined directly in the companion paper (Neher and
Sakaba, 2001). It would compromise some of our results, because we
assume constant mEPSC size throughout. However, we think this is very unlikely, because we did not observe any increase in quantal sizes immediately after strong stimulation ["late noise" protocol in Neher and Sakaba (2001)], and because Kyn depressed EPSCs without changing their time course significantly, when we used presynaptic stimulation protocols similar to those used here (data not shown). Such
changes should occur if displacement takes place.
Using the deconvolution method, we show that the peak rate of
release varies according to a power function of the amplitude of
Ca2+ current with an exponent of 3-4,
when a first short pulse of Ca2+ influx is
applied after a pause of at least 10 sec (Borst and Sakmann, 1996,
1999a; Schneggenburger et al., 1999; Wu et al., 1999). We also show
that the Ca2+ influx becomes more
efficient in eliciting release when it is preceded by low-level
Ca2+ influx (Figs. 3,
TOP
ABSTRACT
INTRODUCTION
