Department of Physiology and Biophysics, University of Washington School of Medicine, Seattle, Washington 98195-7290
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ABSTRACT |
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Singer, Joshua H. and Albert J. Berger. Contribution of single-channel properties to the time course and amplitude variance of quantal glycine currents recorded in rat motoneurons. The amplitude of spontaneous, glycinergic miniature inhibitory postsynaptic currents (mIPSCs) recorded in hypoglossal motoneurons (HMs) in an in vitro brain stem slice preparation increased over the first 3 postnatal weeks, from 42 ± 6 pA in neonate (P0-3) to 77 ± 11 pA in juvenile (P11-18) HMs. Additionally, mIPSC amplitude distributions were highly variable: CV 0.68 ± 0.05 (means ± SE) for neonates and 0.83 ± 0.06 for juveniles. We wished to ascertain the contribution of glycine receptor (GlyR)-channel properties to this change in quantal amplitude and to the amplitude variability and time course of mIPSCs. To determine whether a postnatal increase in GlyR-channel conductance accounted for the postnatal change in quantal amplitude, the conductance of synaptic GlyR channels was determined by nonstationary variance analysis of mIPSCs. It was 48 ± 8 pS in neonate and 46 ± 10 pS in juvenile HMs, suggesting that developmental changes in mIPSC amplitude do not result from a postnatal alteration of GlyR-channel conductance. Next we determined the open probability (Popen) of GlyR channels in outside-out patches excised from HMs to estimate the contribution of stochastic channel behavior to quantal amplitude variability. Brief (1 ms) pulses of glycine (1 mM) elicited patch currents that closely resembled mIPSCs. The GlyR channels' Popen, calculated by nonstationary variance analysis of these currents, was ~0.70 (0.66 ± 0.09 in neonates and 0.72 ± 0.05 in juveniles). The decay rate of patch currents elicited by brief application of saturating concentrations of glycine (10 mM) increased postnatally, mimicking previously documented changes in mIPSC time course. Paired pulses of glycine (10 mM) were used to determine if rapid GlyR-channel desensitization contributed to either patch current time course or quantal amplitude variability. Because we did not observe any fast desensitization of patch currents, we believe that fast desensitization of GlyRs underlies neither phenomenon. From our analysis of glycinergic patch currents and mIPSCs, we draw three conclusions. First, channel deactivation is the primary determinant of glycinergic mIPSC time course, and postnatal changes in channel deactivation rate account for observed developmental changes in mIPSC decay rate. Second, because GlyR-channel Popen is high, differences in receptor number between synapses rather than stochastic channel behavior are likely to underlie the majority of quantal variability seen at glycinergic synapses throughout postnatal development. We estimate the number of GlyRs available at a synapse to be on average 27 in neonate neurons and 39 in juvenile neurons. Third, this change in the calculated number of GlyRs at each synapse may account for the postnatal increase in mIPSC amplitude.
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INTRODUCTION |
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Glycine is the predominant inhibitory neurotransmitter in the
mammalian spinal cord and brain stem (Werman et al.
1967), where the timing and strength of inhibitory synaptic
transmission govern rhythmic motor output (Bracci et al.
1996
; Paton and Richter 1995
). Postnatal
maturation of glycinergic synapses is characterized by a change in
glycine receptor (GlyR)-channel subunit composition; during the first
week of life, the fetal
2 subunit is replaced by the adult
1
subunit (Malosio et al. 1991
). As a consequence of this
alteration in channel subunit composition, GlyR-channel kinetics become
faster, and the time course of glycinergic inhibitory synaptic currents
(IPSCs) becomes shorter (Krupp et al. 1994
; Singer et al. 1998
; Takahashi et al.
1992
). The GlyR-channel properties, which change postnatally
and underlie the altered IPSC decay time course, are unknown. Previous
developmental studies of GlyRs examined only the steady-state behavior
of channels as opposed to their kinetic properties under the dynamic,
nonequilibrium conditions that exist at intact synapses
(Frerking and Wilson 1996
).
The amplitude distribution of glycinergic spontaneous miniature IPSCs
(mIPSCs), like that of quantal synaptic responses throughout the CNS,
is skewed toward large amplitudes and is highly variable (e.g.,
Bekkers et al. 1990; Edwards et al.
1990
). The source of this variability is a subject of debate
(Bennett 1995
; Frerking and Wilson 1996
),
and much discussion centers around the question of whether postsynaptic
receptors are saturated by a single vesicle of transmitter (e.g.,
Tang et al. 1994
). If they are then the majority of
quantal variability must be attributed to differences in the number of
receptors at individual synapses or between-site variability
(Faber et al. 1992
; Hestrin 1992
;
Nusser et al. 1997
). However, if the postsynaptic
receptors at a synapse are not saturated by a quantum of transmitter,
the postsynaptic response at a single release site will vary from
quantal event to quantal event, and this will account for much of the
observed quantal variability (Frerking et al. 1995
).
Rapid agonist application to outside-out patches was used to mimic
synaptic release in studies of channels underlying fast synaptic
transmission (e.g., Clements 1996; Edmonds et al.
1995
; Jonas and Spruston 1994
). This technique
permits systematic investigation of channel behavior under conditions
where the concentration and duration of agonist exposure can be
controlled. By using the rapid agonist-application technique, we
examined the contribution of GlyR-channel properties to both the time
course and amplitude variability of quantal synaptic currents recorded
in hypoglossal motoneurons (HMs) in rat brain stem slices throughout
the early postnatal period.
HMs control the tongue and subserve motor functions, including
respiration, vocalization, and deglutition (Lowe 1980).
Tongue muscle tone, particularly that of the extrinsic genioglossus and styloglossus muscles, is modulated throughout the respiratory cycle to
maintain upper airway patency (reviewed by Lowe 1980
), and loss of inspiration-related activity in these muscles is thought to
underlie respiratory pathologies such as obstructive sleep apnea
(Remmers et al. 1978
). Activation of afferent sensory
inputs to the hypoglossal motor nucleus (n. XII) elicits both
excitatory and inhibitory potentials in HMs (Kubin et al.
1993
; Lowe 1978
; Sumino and Nakamura
1974
; Withington-Wray et al. 1988
), which are
thought to coordinate tongue movements during complex behaviors such as
mastication. Glycinergic synaptic transmission to HMs then may be of
particular importance because it allows the tongue to perform a variety
of voluntary motor functions while receiving rhythmic, respiratory
input. Additionally, the hypoglossal motor nucleus (n. XII) exhibits
the highest GlyR density, as assayed by 3H-strychnine
binding, of any area in the CNS (White et al. 1990
), making it an excellent system in which to study inhibitory neurotransmission.
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METHODS |
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Brain stem slice preparation
Experiments were performed with brain stem slices from
Sprague-Dawley rats (P0-18). For simplicity, animals were placed in two age groups, neonate (P0-3) and juvenile (P11-18). HMs exhibit adult-like GlyRs and glycinergic synaptic currents by P10-14, and
motoneurons acquire adult-like electrophysiological, morphological, and
biochemical properties by the end of the second postnatal week
(Berger et al. 1996; Kalb and Hockfield
1994
). Animals were anesthetized by injection of a
ketamine-xylazine mixture (200 and 14 mg/kg im, respectively) and
decapitated. Brain stems were isolated and cut in 300-µm transverse
sections with a vibratome (Pella) in an ice-cold Ringer solution
containing (in mM) 120 NaCl, 26 NaHCO3, 1.25 NaH2PO4, 3 KCl, 20 glucose, 1 CaCl2, and 5 MgCl2. Slices were incubated at
37°C for 1 h in this Ringer solution and then maintained at room
temperature (20-22°C) in the same solution (except with 2 CaCl2 and 2 MgCl2). Solutions were bubbled
continuously with a 95% O2-5% CO2 gas mixture.
Data acquisition and analysis
Whole cell and outside-out patch recordings were obtained from
visualized HMs in brain stem slices at room temperature (20-22°C). Slices were submerged in a chamber mounted on a fixed-stage microscope (Carl Zeiss) equipped with Nomarski optics and a ×40 water-immersion objective. Slices were illuminated with near-infrared light (750-790 nm), and HMs were visualized with an infrared sensitive charge-coupled device video camera (Hamamatsu) connected to a video monitor (Sony). HMs were identified by their location within the hypoglossal nuclei (n.
XII) and by their size (15-25 µm) and multipolar shape
(Umemiya and Berger 1994).
Recordings were made with borosilicate glass pipettes (Clark
Electromedical) containing (in mM) 130 CsCl, 10 NaCl, 4 MgCl2, 10 HEPES, 10 EGTA, 4 ATP-Mg, and 0.4 GTP-Tris for
whole cell recording or 140 N-methyl-D-glucamine
Cl (NMDGCl), 5 HEPES, 1 EGTA, 3 ATP-Mg, 0.3 GTP-Tris or 120 Cs-methanesulfonate, 4 CsCl, 4 NaCl, 10 HEPES, 10 EGTA, 5 lidocaine
N-ethyl bromide (QX-314), 4 ATP-Mg, and 0.4 GTP-Tris for
patch recording. The measured liquid junction potential of the
CsCl-rich and NMDGCl internal solutions was ~2-3 mV, and, because it
was small, holding potentials were not compensated for it. The
Cs-methanesulfonate pipette solution had a measured liquid junction
potential of ~35 mV, and the holding potentials of patch current
recordings were corrected off-line. Pipette resistances were 3-5 M
for whole cell and 7-10 M
for patch current recording. Pipette
solutions' pH was adjusted to 7.3 by CsOH, and osmolarity was adjusted
to 305 mOsm by sucrose. Series resistance during whole cell recording
was <10 M
and was compensated 70-98%; experiments were terminated
if the series resistance increased by >25%.
The extracellular solution for whole cell recording was composed of (in
mM) 120 NaCl, 26 NaHCO3, 1.25 NaH2PO4, 3 KCl, 20 glucose, 2 CaCl2, and 2 MgCl2. Osmolarity was adjusted to
315 mOsm by addition of sucrose. Bicuculline methiodide (10 µM,
Sigma), 6,7-dinitro-quinoxaline (DNQX, 10 µM, Research
Biochemicals), and D()-2-amino-5-phosphono-pentanoic acid
(APV, 25 µM, Research Biochemicals) were added to block
GABAA, AMPA, and
N-methyl-D-aspartate receptor-mediated currents,
respectively. TTX (0.5-1.0 µM, Calbiochem) and CdCl2
(100 µM) were added to block action potential- and calcium
channel-dependent synaptic transmission. For patch recording, the
extracellular solution contained either (in mM) 120 NaCl, 20 TEA-Cl, 10 HEPES, 2 CaCl2, 2 MgCl2 or 132 N-methyl-D-glucamine Cl, 10 HEPES, 11 glucose, 2 CaCl2 and 1 MgCl2; pH was adjusted to 7.4 with
NaOH, and osmolarity was adjusted to 315 mOsm by addition of sucrose.
Additionally, TTX (0.5 µM) and CdCl2 (50 µM) were added
to block Na+ and Ca2+ currents, respectively.
Glycine (Sigma) at varying concentrations was used to elicit
GlyR-channel-mediated currents.
Voltage-clamp recordings were made with an Axopatch 200B amplifier
(Axon Instruments), and the recording chamber was perfused at ~3
ml/min. Holding potential (Em) was 70 mV for
whole cell recording and
70 or
35 mV for patch current recording.
The calculated reversal potential (Erev) of
Cl
was
0.4 mV for the NMDGCl pipette solution and
71
mV for the Cs-methanesulfonate pipette solution; single channel
conductance (g) was calculated from the single channel
current (i) as g = i/(ECl
Em). Rapid solution exchange was accomplished with either a high-voltage piezoelectric stack translator (Physik Instrumente, model P-244.40 with an E-470 power supply) or
piezoelectric bimorph element (Piezo Systems) to move theta glass flow
pipes (Hilgenberg) across a membrane patch. To ensure uniform flow, a
syringe pump (Harvard Apparatus) was used (flow rate ~0.1 ml/min). Rapid solution exchange (10-90% rise time <400 µs for bimorph, <200 µs for stack translator) was confirmed at the end of each experiment by switching between the control patch solution and one
diluted 50% with distilled H2O and monitoring the open-tip current after patch rupture. Illustrated patch currents and IPSCs represent the average of five or more trials and are digitally filtered
at 2 kHz for display purposes.
Signals were filtered at 2-5 kHz and digitized at 5-10 kHz (pCLAMP,
Axon Instruments or WCP, Strathclyde Electrophysiology Software). A
software package developed in our laboratory and using the detection
algorithm described by Cochran (1993) was used to identify spontaneous
mIPSCs. A minimum of 200 events was recorded in each neuron. Whole cell
and ensemble patch current decays were fit by a Chebychev algorithm
(pClamp). Spontaneous mIPSCs and ensemble patch currents were best fit
by two exponentials, and the mean time constant,
decay,
was calculated from the time constants and their relative amplitudes,
decay =
fastafast +
slowaslow.
Data are presented as means ± SE unless otherwise noted. Statistical significance was determined with analysis of variance (ANOVA) for between-group comparisons. Miniature IPSC amplitude distributions were compared with a Kolmogorov-Smirnoff test. To assay changes in EC50 and Hill coefficient (h) values for statistical differences, the sigmoid logistic equations fit to the concentration-response data were linearized by a logarithmic transformation, and differences in the slopes and intercepts were tested for statistical significance by analysis of covariance (ANCOVA). Changes were considered significant if P < 0.05.
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RESULTS |
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Quantal current amplitude is highly variable
We recorded glycinergic mIPSCs in neonate (n = 8)
and juvenile (n = 13) HMs. To reduce the possibility
that recorded events were altered significantly by electrotonic
filtering, we limited our analysis to events with a 10-90% rise time
1 ms (no correlation between mIPSC amplitude and half-width or rise
time was observed in populations of mIPSCs selected in this fashion;
data not shown); the lack of correlation between these parameters
suggests but does not provide unequivocal support for the absence of
electrotonic filtering of synaptic currents (Soltesz et al.
1995
; Spruston et al. 1993
). Amplitude
distributions were highly skewed (Fig. 1,
A and B): mean amplitude = 42 ± 6 pA,
skewness = 1.5 ± 0.3 in neonates, and mean amplitude = 77 ± 11 pA, skewness = 1.9 ± 0.3 in juveniles. The
postnatal change in quantal amplitude is statistically significant
(Fig. 1C, P < 0.05 by the
Kolmogorov-Smirnov test); the skewness of neonate and juvenile
amplitude distributions is not statistically different
(P = 0.3 by ANOVA).
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Glycinergic mIPSC amplitude distributions were highly variable in both neonate and juvenile HMs. To quantify this variability, we calculated a CV for each distribution: CV = SD/mean. The mean CV was 0.68 ± 0.05 for neonate and 0.83 ± 0.06 for juvenile motoneurons (Fig. 1, A and B); this difference was not statistically significant (P = 0.10 by ANOVA). Our observations of these spontaneous mIPSCs led us to ask two questions. First, what accounts for the postnatal increase in quantal amplitude; second, what is the source of the variability between quantal currents?
Estimation of GlyR-channel conductance at the intact synapse
GlyR-channel subunit composition is different in neonate and
juvenile HMs; from our previous work (Singer et al.
1998) we concluded that GlyRs are primarily
2/
heteromers
in neonates and
1/
heteromers in juveniles. We considered the
possibility that the conductance of synaptic GlyR channels increases
postnatally, thereby increasing the amplitude of recorded mIPSCs. To
estimate the single-channel conductance of ligand-gated channels
underlying quantal glycinergic synaptic currents, we analyzed the
current variance associated with GlyR-channel opening. For each neonate and juvenile HM, spontaneous mIPSCs were aligned along their rising phases and averaged. The average mIPSC from each neuron was scaled to
the peak amplitude of each individual mIPSC comprising it, and then the
peak-scaled average current was subtracted from the individual mIPSCs.
The resultant difference currents are the result of random channel
fluctuation around the mean (Robinson et al. 1991
;
Sigworth 1980
; Traynelis et al. 1993
) and
are illustrated in Fig. 2, A
and B. The average mIPSCs were binned intervals of 5% of
peak current amplitude along the current decay phase, and their
variance (
2, equivalent to the difference current
squared) was plotted against the binned current. The initial 25% of
the relationship between variance and current was fit with a straight
line, the slope of which is the mean single-channel current
(i) representing the weighted average of the channels'
various subconductance states (Traynelis et al. 1993
).
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The mean single-channel current of the GlyR at an intact synapse was
3.4 ± 0.4 pA for neonates (n = 5) and 3.2 ± 0.7 pA for juveniles (n = 6), corresponding to a mean
single-channel conductance of 45-50 pS (48 ± 8 pS in neonates
and 46 ± 10 pS in juveniles; this difference is not significant,
P = 0.9 by ANOVA). That Gly-R single-channel
conductance does not change with postnatal development is in keeping
with our previous study of HM GlyR single-channel properties
(Singer et al. 1998). Further, our calculated mean single-channel conductance of 48 pS in neonate motoneurons is virtually
identical to the reported GlyR single-channel main conductance of 43 pS
that was measured directly in neonatal dorsal horn neurons (Takahashi and Momiyama 1991
). The observed differences
in mIPSC amplitude between neonate and juvenile HMs then are not the
result of changes in GlyR single-channel conductance.
GlyR-channel open probability
We examined next the possible contribution of stochastic
GlyR-channel behavior to the total mIPSC amplitude. Specifically, we
wished to determine the GlyR channels' open probability
(Popen) under conditions that approximated those
at an intact synapse, where channels are exposed presumably to
transient, high concentrations of agonist (Clements
1996). By using a piezoelectric bimorph element to move theta
glass flow pipes (see METHODS), brief pulses (1 ms) of
glycine (1 mM) were applied to patches excised from neonate and
juvenile HMs (EM =
70 mV, NMDGCl internal and
external solutions). We chose this glycine concentration because it is
thought to approximate the transmitter concentration at other fast,
central mammalian synapses (Clements et al. 1992
;
Jones and Westbrook 1995
; Maconochie et al.
1994
).
Short (1 ms) pulses of glycine (1 mM) elicited patch currents that
closely resembled mIPSCs (Fig. 3,
A1 and A3, average traces). Figure
3A shows patch current responses to 1-ms pulses of glycine (1 mM). The mean and variance (2) of 10 responses were
calculated and binned in intervals of 5% of mean peak current
amplitude (IP) along the current decay phase. Binned current and variance were plotted, and the current-variance relationship was fit with the parabolic equation (Eq. 1)
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(1) |
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Popen was 0.66 ± 0.09 in neonates (n = 8) and 0.72 ± 0.05 in juveniles (n = 13); the difference in Popen was not statistically significant (P = 0.15 by ANOVA). Calculated values for i were 3.1 ± 0.4 pA in neonates and 2.5 ± 0.2 pA in juveniles (not statistically different, P = 0.92 by ANOVA). These values of i correspond to mean single-channel conductances of 44 ± 6 pS and 35 ± 3 pS for neonate and juvenile GlyR channels, respectively. These values are almost identical to the most prevalent single-channel closings observed in the decay phase of macroscopic patch currents (Fig. 3, A2 and A4, insets) as well as those estimated for GlyR channels at the intact synapse (see the previous section and Fig. 2C) and are not significantly different from the latter (P = 0.14 by ANOVA). The calculated number of channels (N) in neonate patches (39 ± 14 channels) was however much smaller than that in patches from juvenile HMs (154 ± 59 channels). This difference, although large, was not statistically significant (P = 0.06 by ANOVA). We conclude that the probability of GlyR-channel opening is quite high when concentrations of agonist approximate those at the intact synapse.
Glycinergic patch currents lack a significant rapidly desensitizing component
We considered next the possibility that mIPSC amplitude variance
resulted in part from some or all of the synaptic GlyR channels rapidly
entering into and slowly recovering from a desensitized state. To look
for fast channel desensitization on the time scale of a synaptic
current, we used a paired-pulse, rapid-application protocol; a second
1-ms glycine pulse was applied to patches 5 ms after the initial pulse,
and a paired-pulse ratio (PPR) was calculated from the resultant
currents as I2/I1
(EM = 35 mV).
To permit such closely spaced pulses of transmitter (resulting from
closely spaced translations of the theta glass flow pipes), we used a
high-voltage piezoelectric stack translator rather than a bimorph
element to move the theta glass perfusion apparatus during these
experiments; the stack translator expands and relaxes much more quickly
than the bimorph element does. Additionally, we used an NaCl- rather
than an NMDGCl-based perfusate because the former is less viscous, and
therefore the flow streams can be moved more quickly. Finally, we used
a Cs-methanesulfonate- rather than an NMDGCl-based internal pipette
solution because it improved patch stability in the changed perfusate
(see METHODS; patch currents are outward in these
experiments). Neither the rise times nor the decay rates of glycinergic
patch currents were affected by these changes (data not shown). To
ensure maximal activation of the GlyR channels in the patch, a
saturating (10 mM) concentration of glycine was used (Berger et
al. 1997).
If a significant number of the GlyR channels in the patch desensitized after the initial glycine pulse, the PPR would be much less than 1. The PPR, however, was 0.95 ± 0.02 in neonates (n = 7) and 0.94 ± 0.01 in juveniles (n = 8, Fig. 4A). At subsaturating glycine concentrations (0.5 mM), the absolute amplitude of the second pulse was as expected greater than that of the first; PPR = 1.43 ± 0.12 in neonates (n = 3) and 1.19 ± 0.04 in juveniles (n = 5). Thus GlyR-channel-mediated currents show virtually no rapidly desensitizing component throughout postnatal development, and GlyR-channel desensitization does not contribute to quantal amplitude variability.
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Whole cell glycine currents desensitize slowly, usually after hundreds
of milliseconds (Agopyan et al. 1993; Melnick and
Baev 1993
); this property of GlyR channels is preserved in
excised patches (Harty and Manis 1998
). Lengthy
application of glycine (10 mM, 8 s) elicits GlyR desensitization,
as evidenced by the profound reduction in patch-current amplitude in
the presence of agonist illustrated in Fig. 4B. Current
desensitization was biexponential, and a mean time constant of
desensitization,
desens, was calculated;
desens
= 1,172 ± 157 ms in neonate (n = 7) and 1,097 ± 81 ms in juvenile (n = 4) patches, a time
course that is far slower than the decay of either mIPSCs or patch
currents elicited by rapid application of glycine.
Patch current decay time course becomes faster with postnatal development
We demonstrated previously that the decay time course of
glycinergic mIPSCs recorded in HMs becomes faster with postnatal development (decay = 14.2 ± 2.4 ms in neonates and
6.3 ± 0.7 ms in juveniles, a change of 56%), concomitant with a
developmental change in GlyR subunit expression and resultant
single-channel, steady-state kinetics (Singer et al.
1998
). Here we find that the decay time course of the patch
current response (glycine, 10 mM) was accelerated markedly in patches
from older animals (Fig. 4C). The current decay time course
was best fit with two exponentials:
fast = 12.1 ± 0.9 ms (65 ± 3%) in neonate and 7.7 ± 0.8 ms (70 ± 2%) in juvenile patches;
slow = 71.4 ± 4.9 ms (35 ± 3%) for neonates and 42.0 ± 2.8 ms (30 ± 2%)
for juveniles. Thus with postnatal development mean
decay is reduced from 33.4 ± 3.4 ms (neonates,
n = 24) to 17.3 ± 1.1 ms (juveniles,
n = 17), a change of 48%.
GlyR-channel concentration-response relationship
In a heterologous expression system, fetal 2 and adult
1
GlyR channels exhibit almost identical concentration-response
relationships for glycine under steady-state conditions
(Schmieden et al. 1992
; see Akagi and Miledi
1988
). The ability of glycine to activate fetal and adult GlyR
channels under the nonequilibrium conditions presumed to exist at an
intact synapse (Clements 1996
; Frerking and
Wilson 1996
) was not examined, however. Because variability in
the number of GlyRs activated from event to event at an individual synapse might contribute to the amplitude variability of mIPSCs, we
examined the nonequilibrium dose-response relationship of GlyRs by
applying brief (1 ms) pulses of varying concentrations of glycine (0.1, 0.2, 0.5, 1.0, and 10.0 mM) to outside-out patches. Current amplitude
was normalized to the patch response to 10 mM glycine (Imax), a suprasaturating dose in both neonate
and juvenile patches as determined by analysis of patch-current
responses to long (50 ms) glycine pulses and evidenced by the fact that
the amplitude of the current response did not change with increasing
agonist pulse duration at this concentration (n = 18 neonate and 14 juvenile patches). By using the nonstationary variance
analysis described previously, we estimated the GlyR channels' maximal
open probability (Popen,max) under saturating
concentrations of agonist (glycine, 10 mM) and found it to be 0.79 ± 0.04 in neonates and 0.86 ± 0.02 in juveniles
(n = 8 and 9, neonates and juveniles, respectively); the difference in Popen,max was not
statistically significant (P = 0.15 by ANOVA).
Representative patch currents from two different HMs are illustrated in
Fig. 5A. The resulting
concentration-response relationship (Fig. 5B) was fit by a
logistic equation: I/Imax = 1/[1 + (EC50/c)h] where
c is the agonist concentration, h is the Hill
coefficient, and EC50 is the agonist concentration that
elicits a half-maximal response. In neonate patches (n = 21) the EC50 = 0.869 mM, and h = 1.9; in
juvenile patches (n = 18) the EC50 = 0.580 mM, and h = 1.8 (The postnatal change in
EC50 is statistically significant P < 0.05 by ANCOVA; h does not change postnatally. See
METHODS for details on this statistical analysis). In
neither group was the patch current decay time course dependent on
agonist concentration (see Harty and Manis 1998). Thus
over the first 2 wk of postnatal development the ability of glycine to
activate the GlyR under nonequilibrium conditions in enhanced by
~33%.
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DISCUSSION |
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Channel deactivation governs the time course of glycinergic synaptic events
We described previously a postnatal change in the decay time
course of quantal and unitary evoked glycinergic synaptic currents that
results from a developmentally regulated alteration of GlyR-subunit composition (Singer et al. 1998). We and others
(Singer et al. 1998
; Takahashi et al.
1992
) also demonstrated a postnatal change in the kinetics of
native GlyR channels by recording single-channel activity in response
to steady-state application of low (5-10 µM) concentrations of
glycine. These experiments, although informative, are lacking in two
important ways. First, they do not permit an examination of the role of
fast desensitization (with a time course similar to a synaptic current)
in synaptic transmission. Second, and perhaps more importantly, it is
thought that the concentration of transmitter in the synaptic cleft is
high (i.e., 1 mM) and its presence is brief, such that receptors are
not in equilibrium with transmitter throughout the duration of a
synaptic current (Clements 1996
; Frerking and
Wilson 1996
). Thus analysis of steady-state, single-channel
behavior will not yield a complete understanding of postsynaptic
GlyR-channel function. We used therefore the rapid agonist-application
technique to examine the behavior of GlyRs in outside-out patches from
HMs under conditions similar to those at the intact synapse.
We find that the decay rate of patch currents after a 1-ms
application of glycine becomes faster with postnatal development (48 vs. 56% reduction in decay, patch currents vs. mIPSCs).
Because patch current decay after removal of glycine closely mimics the IPSC decay, we see no evidence for glycine rebinding during the course
of an IPSC. We demonstrated previously that transmitter uptake does not
shape the time course of glycinergic synaptic currents (Singer
et al. 1998
). Further, as GlyR-channel desensitization is very
slow and patch currents display virtually no paired-pulse desensitization, we conclude that fast channel desensitization is not
an important determinant of the time course of glycinergic synaptic
transmission. The decay rate of patch responses to brief glycine pulses
and by analogy that of glycinergic synaptic currents reflect channel
deactivation alone. We conclude that the decay phase of a glycinergic
mIPSC is limited by channel closure rather than transmitter rebinding
or desensitization. In this important respect, inhibitory glycinergic
neurotransmission differs from inhibitory GABAergic neurotransmission,
which is influenced significantly by receptor desensitization
(Galarreta and Hestrin 1997
; Jones and Westbrook
1995
). These results are in keeping with the only other study
of GlyR-channel nonequilibrium kinetics (Legendre 1998
).
In our experiments, whereas the time course of patch currents and
mIPSCs were biexponential and changed similarly with postnatal development, patch currents were slower than mIPSCs. This difference was particularly evident in the slow component of current decay. We
considered the possibility that receptors in outside-out patches are
different from those at intact synapses. Glycine channels associate
with the cytoskeleton through the linker protein gephyrin (Prior
et al. 1992; Schmidtt et al. 1987
), and
separating the channels from their cytoskeletal anchors could induce
functional changes in channel behavior (Rosenmund and Westbrook
1993
). To address this issue we examined patch currents
recorded with the cytoskeleton-stabilizing agents phalloidin (2 µM)
or taxol (10 µM) included in the patch pipettes (n = 3 and 5 respectively, data not shown) and found them to be identical to
control currents.
The phosphorylation state of the GlyR may affect its kinetic properties
(Agopyan et al. 1993). We therefore included ATP
S in
the internal solution during patch-current recording but did not
observe any effect on the current time course (n = 5, data not shown). Despite the results of these experiments, we cannot eliminate the possibility that GlyRs in excised patches behave differently from those at intact synapses. Additionally, extrasynaptic receptors, with kinetic properties that may differ from those of
channels underlying the mIPSC (e.g., Nusser et al.
1998
), may be included in the outside-out patches, and this
might account for the slower time course of patch currents relative to mIPSCs.
Importantly, the discrepancy between patch current and mIPSC decay time
course that we observed is consistent with results from other
rapid-application studies of inhibitory synaptic transmission. Several
groups reported that GABAergic patch currents are slower than mIPSCs in
brain slices (Galarreta and Hestrin 1997; Mellor and Randall 1997
; Puia et al. 1994
; Tia
et al. 1996
).
GlyR-channel properties are not the primary source of quantal amplitude variance
The Popen, max of mammalian GlyR
channels, estimated by nonstationary variance analysis of patch
currents, is high, ~0.85 for both neonate and juvenile channels. At
saturating concentrations of glycine then most of the channels in a
patch or at a postsynaptic site will open. GlyR agonist affinity,
however, is fairly low (EC50 0.5-1.0 mM),
particularly for neonate channels. What then is the
Popen of a channel at an intact synapse?
If we assume that the intrasynaptic glycine concentration is 1 mM,
based on estimates from central, glutamatergic synapses (Clements 1996), we can calculate the fraction of
channels at a synapse, f, that will open in response to
a single quantum of glycine; f = N/Ntot =
Popen, max × (I1
mM/Imax), where N is
the number of open channels, Ntot is the
number of channels at the synapse, I1mM is
the average current elicited by 1 mM glycine, and
Imax is the average current elicited by 10 mM glycine.
In neonate patches I1mM/Imax = 0.57, and in juvenile patches I1 mM/Imax = 0.71; f, then, is 0.45 for neonate channels and 0.61 for juvenile channels (assuming a linear relationship between open probability and current amplitude). Because GlyR-channel Popen is high, f will remain constant from event to event as long as the intrasynaptic glycine concentration remains constant. At glycinergic synapses then approximately one-half of the postsynaptic receptors at a single release site will respond to a single quantum of transmitter.
Given the measured mIPSC mean amplitudes and observed
GlyR-channel properties, we can estimate the contribution of channel activation to the CV of mIPSC amplitude distributions, as detailed by
Hestrin (1992). If quantal glycinergic currents arise from activation
of independent channels with identical behavior, their peak amplitudes
will be described by binomial statistics: m = Ntot fi and
2
= Ntot f(1
f ) × i2, where
m is the mean quantal current; CV, or
/m, will be [(1
f )/Ntot f ]1/2.
We determined the mean single-channel current of the GlyRs that underlie mIPSCs: i = 3.4 pA in neonates and 3.2 pA
in juveniles; Ntot, or the number of
available channels at a synapse, was calculated to be 27 channels for
neonates and 39 channels for juveniles, based on a measured mean
quantal amplitudes, m, of 42 and 77 pA and
f = 0.45 and 0.61 for neonates and juveniles,
respectively. The CV resulting from channel activation is therefore
0.21 in neonate HMs and 0.13 in juveniles. Some of this variability may arise from the fact that GlyR channels are not saturated by a single
quantum of transmitter, based on our assumption of an intrasynaptic glycine concentration of 1 mM. These factors contribute to ~10-25% of the mIPSC amplitude distributions' total variance. Quantal variance
resulting from GlyR-channel variance is greater for neonate than
juvenile mIPSCs, as would be expected from the neonate GlyR's lower
affinity for glycine (i.e., higher EC50) under
nonequilibrium conditions.
The primary source of quantal variability, then, is likely differences in the number of receptors between glycinergic synapses, and the difference in quantal amplitude between neonate and juvenile HMs is most probably due to a postnatal increase in the number of GlyR channels at glycinergic synapses. We cannot exclude, however, the possibility that some of the largest mIPSCs arise from synchronous release of multiple quanta and that the extent of quantal variability is overestimated.
Although the ultrastructure of these synapses on HMs was not studied,
Alvarez et al. (1997) described large differences in the size of
postsynaptic clusters of GlyRs on mature cat spinal motoneurons; this
reflects presumably large differences in the number of GlyRs at
individual synapses. In neonatal synapses, where GlyR clustering
mediated by the linker protein gephyrin is not complete (Bechade
et al. 1996
; Kirsch and Betz 1998
), it is quite
likely that similar, large differences in postsynaptic receptor number
also exist. Additionally, the possibility exists that the postsynaptic
receptor complex is poorly defined at immature synapses, and its
geometry is not optimized for signal detection. This would make neonate
synapses more sensitive to variation in intrasynaptic glycine
concentration. This assertion is supported qualitatively by the
observation that the number of channels contributing to macroscopic
patch currents tends to be much larger in patches excised from juvenile
as opposed to neonate HMs. Further, the calculated number of receptors
at a juvenile synapse is 44% larger than that at a neonate synapse
(n = 39 vs. 27).
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ACKNOWLEDGMENTS |
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We thank Drs. D. Koh and J. Isaacson for helpful advice, Drs. J. Dempster and W. Satterthwaite for analysis software, and P. Huynh for technical assistance.
J. H. Singer was supported by a predoctoral fellowship from the National Science Foundation. This work was made possible by Javits Neuroscience Award NS-14857 and National Heart, Lung, and Blood Institute Grant HL-49657 to A. J. Berger.
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FOOTNOTES |
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Present address and address for reprint requests: J. H. Singer, NIH/NINDS, 36 Convent Dr., MSC-4156, Bldg. 36, Rm. 5B21, Bethesda, MD 20892-4156.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 29 October 1998; accepted in final form 4 January 1999.
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REFERENCES |
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