Contribution of Single-Channel Properties to the Time Course and Amplitude Variance of Quantal Glycine Currents Recorded in Rat Motoneurons

Joshua H. Singer and Albert J. Berger

Department of Physiology and Biophysics, University of Washington School of Medicine, Seattle, Washington 98195-7290


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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.


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 alpha 2 subunit is replaced by the adult alpha 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.


    METHODS
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ABSTRACT
INTRODUCTION
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 MOmega for whole cell and 7-10 MOmega 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 MOmega 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, tau decay, was calculated from the time constants and their relative amplitudes, tau decay = tau fastafast + tau 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.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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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Fig. 1. Spontaneous miniature inhibitory postsynaptic current (mIPSC) amplitude distributions are highly skewed and variable throughout postnatal development. A1: amplitude histogram of several hundred mIPSCs (10-90% rise time <= 1 ms) recorded in a neonate (P0) hypoglossal motoneuron (HM). Mean amplitude = 38 pA. A2: distribution of recording noise does not overlap that of the mIPSCs, illustrating that quantal synaptic currents can be resolved clearly, as illustrated in A3. B1-B3: as in A, amplitude histogram from a juvenile (P16) HM. Mean amplitude = 80 pA. Note the increase in mean quantal size with postnatal development. C: cumulative probability distributions reflect the developmental increase in quantal amplitude, as illustrated by a shift of the juvenile distribution toward larger amplitudes. Shown are averages of distributions from n = 8 neonate and n = 13 juvenile motoneurons.

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 alpha 2/beta heteromers in neonates and alpha 1/beta 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 (sigma 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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Fig. 2. Estimation of glycine receptor (GlyR) single-channel conductance from nonstationary variance analysis of spontaneous mIPSCs. A1 and B1: 5 mIPSCs (top traces) recorded in a neonate (A1) and a juvenile (B1) motoneuron are scaled to the peak of the mean mIPSC (bottom traces, the mean mIPSC is a heavy, white line). A2 and B2: difference currents resulting from subtraction of mean peak-scaled mIPSCs from the mean mIPSC. Note the amplitude fluctuations around the baseline after the mIPSC peak (arrow) that result from channel variance. C1 and C2: current-variance relationship for mIPSCs. Single-channel current is taken as the slope of a line fit to the initial 25% of the data: i = 3.8 pA for the neonate (C1) and 3.2 pA for the juvenile (C2) neuron.

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 (sigma 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)
&sfgr;(<IT>t</IT>)<SUP><IT>2</IT></SUP><IT>=</IT><IT>iI</IT>(<IT>t</IT>)<IT>−</IT><IT>I</IT>(<IT>t</IT>)<SUP><IT>2</IT></SUP><IT>/N</IT> (1)
where i is the mean single-channel current, I(t) is the macroscopic patch current, and N is the number of channels in the patch. From this fit, Popen is calculated as IP/Ni. Representative current-variance plots and their fits are illustrated in Fig. 3B.



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Fig. 3. GlyR-channel Popen is high and does not change with postnatal development. A1 and A3: 5 current responses to 1-ms steps into glycine (1 mM) recorded in outside-out patches excised from neonate (A1) and juvenile (A3) HMs (light traces) with average responses (darker traces) overlaid. A2 and A4: single patch current responses from those grouped in A1 and A3 show single channels closing along the decay phases. These closings (in ovals) illustrate the single GlyR-channel conductance of the channels contributing to the macroscopic patch current and are amplified in the figure insets. The observed single-channel current amplitude is identical to that predicted by nonstationary variance analysis of these currents. B1 and B2: current-variance relationships from traces in A fit with the parabolic relationship: sigma (t)2 = iI(t- I(t)2/N. From this fit, Popen is calculated as I/Ni. For the neonate patch in A1, i = 3.1 pA, n = 13 channels, and Popen =0.72; i = 2.7 pA, n = 20 channels, and Popen = 0.76 for the juvenile patch in A3.

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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Fig. 4. GlyR channels do not desensitize rapidly, and patch-current decay in the absence of glycine becomes faster with postnatal development. A1 and A2: currents from neonate and juvenile patches illustrate an almost complete absence of fast (5-ms paired-pulse interval) desensitization (glycine 10 mM). B1 and B2: glycine patch currents desensitize slowly and almost completely during a long (8 s) exposure to glycine (10 mM). C: patch-current decay becomes faster postnatally. Neonate (C1) and juvenile (C2) patch current responses to 1-ms pulses of 10 mM glycine are normalized to the same peak amplitude (C3).

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, tau desens, was calculated; tau 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 (tau 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: tau fast = 12.1 ± 0.9 ms (65 ± 3%) in neonate and 7.7 ± 0.8 ms (70 ± 2%) in juvenile patches; tau 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 tau 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 alpha 2 and adult alpha 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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Fig. 5. Concentration-response relationship for patch current responses to brief (1 ms) glycine pulses show a postnatal increase in agonist potency. A1 and A2: currents elicited by 0.5 and 10 mM glycine in neonate and juvenile patches; 0.5 mM glycine activates a smaller proportion of available receptors in the neonate patch. Insets: patch-current decay is independent of agonist concentration. Currents are normalized to the same peak amplitude and show identical rates of decay. All scale bars as in A1. B: concentration-response relationship for n = 21 neonate and n = 18 juvenile patches. Points were fit by a logistic equation, I/Imax = 1 /[1 + (EC50/c)h]. Note the 33% reduction in calculated EC50 with postnatal development.


    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 tau 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 ATPgamma 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 approx  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 sigma 2 = Ntot f(1 - f ) × i2, where m is the mean quantal current; CV, or sigma /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).


    ACKNOWLEDGMENTS

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.


    FOOTNOTES

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.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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