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1 Institut de Neurobiologie de la Méditerranée (INMED) INSERM U29, 163, route de Luminy, 13273 Marseille cedex 09, France
2 Dipartimento di Fisiologia Umana e Farmacologia, Università La Sapienza, Roma, P le A. Moro 5, I-00185, Italy
3 Istituto Neurologico Mediterraneo (Neuromed), Pozzilli (IS), Italy
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Abstract |
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(Received 18 July 2005;
accepted after revision 24 August 2005;
first published online 25 August 2005)
Corresponding author P. Bregestovski: Institut de Neurobiologie de la Méditerranée, INSERM U29, 163, route de Luminy, 13273 Marseille cedex 09, France. Email: pbreges{at}inmed.Univ-mrs.fr
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Introduction |
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To answer this question, we analysed the effects of Cai on glycinergic inhibitory postsynaptic currents (IPSCs) in hypoglossal motoneurones (HMs) from rat and mouse brainstem acute slices. HMs are characterized by rhythmic activity and they are involved in a variety of motor functions, including breathing, chewing, sucking, swallowing and phonation (Peever & Duffin, 2001). HMs receive synaptic information from various parts of the brain due to dendritic arborization, and exhibit two remarkable features. The first is a low endogenous Ca2+-buffering capacity, which determines both the selective vulnerability of HMs to Ca2+-related excitotoxicity and the rapid dynamics of Cai (Lips & Keller, 1998; Palecek et al. 1999). The second is powerful glycinergic synaptic inputs, which present the major inhibitory drive in this brainstem nucleus (Umemiya & Berger, 1995; Singer et al. 1998; Donato & Nistri, 2000; Singer & Berger, 2000). HMs thus provide a convenient model for examination of the action of intracellular Ca2+ on glycinergic synapses.
We report here that Cai elevation in HMs modulates glycinergic synaptic transmission by two independent mechanisms of opposite sign: (i) a decrease in glycinergic IPSCs due to the reduction in presynaptic glycine release induced, predominantly, by retrograde action of endogenous cannabinoids; (ii) potentiation of postsynaptic GlyRs. Under normal physiological conditions in HMs, the postsynaptic effect is masked by powerful presynaptic inhibition.
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Methods |
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Experiments were carried out on brainstem slices from postnatal (P5–P9) Sprague-Dawley or Wistar rats and C57BL/6J or Swiss mice. Animals were anaesthetized with ether and killed by decapitation in agreement with the European Directive 86/609/EEC requirements. The brainstem was rapidly removed and placed in an oxygenated ice-cold saline buffer. Transverse 250-µm-thick brainstem slices were cut using a DSK (Dosaka, Japan) or HM 650V (Microm, Germany) vibrating microslicer in ice-cold oxygenated solution containing (mM): NaCl, 125; KCl, 3.5; CaCl2, 2; MgCl2, 1.3; NaH2PO4, 1.25; NaHCO3, 26; and glucose, 10; equilibrated at pH 7.3 with 95% O2 and 5% CO2. Prior to recording, slices were incubated at room temperature (22–25°C) for at least one hour to allow recovery. Experiments were performed in the same solution at room temperature.
Electrophysiological recordings
For patch-clamp recordings brainstem acute slices were visualized through a x40 water-immersion objective using an upright microscope (Axioskop, Zeiss, Germany). HMs were identified by their location in the hypoglossal nucleus (n.XII), their large somata (25–40 µm) and their dendritic arborization.
During measurements, slices were superfused with oxygenated saline (1.0–1.5 ml min–1). Whole-cell borosilicate glass pipettes (Hilgenberg or Clark Capillaries) with a tip resistance of 2–5 M
were filled with a KCl-based intracellular solution containing (mM) KCl, 135; MgCl2, 1; Hepes, 10; Mg-ATP, 4; Na-GTP, 0.3; equilibrated at pH 7.3 with KOH. Different BAPTA concentrations (0.1–20 mM) were used as indicated in the text. To obtain about 50 nM free Cai in the internal solution, corresponding concentrations of CaCl2 (0.03–6 mM) were added (calculating using developed by Dr F. Mendez (Gottingen) the ‘Patcher's Power Tools’ subroutine of the ‘Igor’ program). In some experiments an alternative CsCl-based intracellular solution was used, containing (mM) CsCl, 140; MgCl2, 2; Hepes, 10; MgATP, 2; Na2GTP, 0.4; CaCl2, 0.11; and EGTA, 0.5 (pH 7.3).
Membrane currents were recorded at 3–10 kHz with an Axopatch 200A (Axon Instruments, USA) or EPC-9 (HEKA Elektronik, Germany) amplifiers. Stored data were analysed using pClamp software (Axon Instruments) or the PulseFit program (HEKA Elektronik). The series resistances, ranging between 5 and 15 M as estimated from slow transient cancellation, were compensated by 30–80% depending on the cell. The holding potential (Vh) was –70 mV. IPSCs were elicited by stimulating (2–100 V, 200–300 µs) at 0.5–1 Hz with glass bipolar electrodes. The stimulus intensity was adjusted to obtain about 20–50% failures. Stimulating electrodes were obtained by pulling theta-glass tubes to a final tip diameter of 1–2 µm and filling with external solution. Pairs of stimuli (at 50 ms intervals) were applied.
To isolate strychnine-sensitive glycinergic synaptic currents all experiments were performed in the presence of blockers of glutamate (CNQX, 10 µM; AP-5, 40 µM) and GABA (bicuculline, 20 µM) receptors unless otherwise mentioned. Control experiments (n= 5) showed that, at the concentrations used, bicuculline did not affect the amplitude of glycinergic IPSCs (see supplementary material). To record glycinergic miniature IPSCs (mIPSCs) TTX (1 µM) was added to the external solution in addition to the antagonists of glutamate and GABA receptors. Cannabinoid (CB) receptor agonist WIN55,212-2 (2,3-dihydro-5-methyl-3-(4-morpholinymethyl)pyrrolo[1,2,3de]-1,4-benzoxazin-6-yl]-1naphthalenylmethanone mesylate) and antagonist SR141716A (N-piperidini-5(4chlorophenyl)-4-methyl-3pyrazole-carboxamide) were bath-applied at concentrations of 3–5 µM. These concentrations are far above affinity constant (Ki) for CB1 receptors (Howlett et al. 2002) and similar concentrations have been used in a number of studies on brain slices (Hajos et al. 2001; (Jennings et al. 2001; Diana et al. 2002; Marsicano et al. 2002; Yoshida et al. 2002). Adding WIN55,212-2 or SR141716A did not change leakage currents or input resistance of recorded HMs.
To increase the cytosolic Ca2+, the recorded neurones were repetitively depolarized to 0 mV (1 s pulse duration). Each single depolarization was preceded by a 100 ms step to –100 mV to allow complete activation of voltage-dependent Ca2+ channels, and was followed by electrical stimulation to produce an IPSC after a delay of 100–300 ms. This protocol was repeated 20–40 times (30 times in most of experiments) and properties of averaged glycinergic IPSCs were compared in control and following depolarizing series.
For external application, agonists were applied via a puff pipette close to the cell soma. For glycine application, pipettes contained either low (30 µM) or high (0.5–5 mM) concentrations of the agonist. Glutamate and NMDA were applied at a concentration of 200 µM. Glycine and NMDA were dissolved in the standard external solution (Ca2+ 2 mM; Mg2+ 1.3 mM). In experiments with application of glutamate, to maximize the influx of Ca2+ the puff pipette contained Mg2+-free extracellular solution with elevated Ca2+ (10 mM). In some experiments, at NMDA application cells were concomitantly depolarized (–20 mV) to give relief from Mg2+ block (Nowak et al. 1984). Effects of NMDA and glutamate were studied with AP-5 omitted from the external solution.
Antagonists and agonists were bath-applied via a gravity-driven perfusion system (unless otherwise specified). AP-5, bicuculline and WIN55,212-2 were from Tocris (UK), SR141716A was the gift from Sanofi–Aventis and TTX was from Latoxan (France). All the other chemicals were from Sigma (USA).
The results presented here have been obtained on 70 HMs from rat and 79 HMs from mouse brain slices. As data obtained on the two preparations were similar they have been pooled (unless otherwise specified).
Statistical analysis was done in Origin (OriginLab Corporation, USA) and SigmaStat (Systat Software Inc., USA) programs using Student's paired t test or Wilcoxon's signed-rank test for non-parametric data. The Kolmogorov–Smirnov statistical test was used to assess differences in distribution of glycinergic mIPSC intervals. Results are given as means ±S.E.M.
Calcium imaging
Monitoring of Cai was conducted using a customized digital imaging microscope allowing simultaneous monitoring of whole-cell currents and fluorescence signals. Fluo-5F (50–100 µM) or, in some experiments, Fura-2 (50–200 µM) (both from Molecular Probes, Netherlands) were included in the internal solution and allowed to diffuse into the neurone for at least 15 min before the beginning of the records. The single (480-nm) or dual (350/380-nm) wavelength excitation was achieved using a 1 nm bandwidth polychromatic light selector equipped with a 100 W xenon lamp (Polychrome II, TILL Electronics, Germany). Fluorescence was visualized using an upright microscope (Axioskop, Zeiss, Germany) and a 12-bit charge-coupled device (CCD) camera equipped with an image intensifier (PentaMax, Princeton Instruments, USA). Fluorescence signals were acquired with variable sampling rate (from 0.2 to 10 Hz), using MetaFluor software (Universal Imaging Co, USA). Because of high Cai transients in HMs, fluorescence signals recorded with Fura-2 showed a tendency to saturation. To avoid this problem, a lower sensitivity Ca2+ dye, Fluo-5F, was used in the experiments described here. To ensure that Ca2+ signals were recorded within the dynamic range of Fluo-5F, the membrane permeability of HMs was artificially increased by a series of electrical discharges at the end of some experiments. Under these conditions the estimated maximal change in fluorescence (
F) was 2–5-fold higher than that induced by depolarization-induced Cai transients.
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Results |
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Properties of glycinergic IPSCs were studied in visually identified HMs. IPSCs were evoked with a theta-tube glass pipette placed either close (100–150 µm) to the recorded neurone or on the surface of small neurones (<10 µm) in the reticular formation, ventral to and immediately outside the hypoglossal nucleus, allowing extracellular monosynaptic stimulation (Umemiya & Berger, 1995). In the presence of antagonists of GABA and glutamate ionotropic receptors, stable IPSCs (range in different cells 50–800 pA, n= 149) were recorded at –70 mV, and their activity was completely abolished by strychnine (3 µM; Fig. 1A). IPSC deactivation kinetics was best fitted by a single exponential. The decay time constant varied in different experimental conditions from 6 to 20 ms, consistently with previously described kinetic properties (Takahashi et al. 1992; Singer et al. 1998; Singer & Berger, 1999; Donato & Nistri, 2000).
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When the recording pipette contained 0.1 mM BAPTA and 0.1 mM Fluo-5F, depolarization caused a rapid Cai rise, whose amplitude was proportional to the number of depolarizing pulses (Fig. 1B). In these conditions, after 10–30 depolarizing pulses, the quasi-maximum values for Cai-induced relative fluorescence amplitudes (Fig. 1C) were reached. For this reason a depolarization protocol employing 30 pulses (otherwise mentioned) was chosen in the subsequent investigations.
Depolarization-induced Cai rise in HMs caused a significant decrease in glycinergic IPSC amplitude. IPSC depression was followed by a recovery of the amplitude within 3–5 min (Fig. 2A and B). As illustrated in Fig. 2C and D, the inhibition become more prominent with elevation of a number of depolarisations, and reached the quasi-maximum values after 20–30 pulses. These results were observed in rat and mouse HMs. In the rat HMs, the depolarization-induced inhibition was recorded with different Ca2+ buffers in the pipette solution. With BAPTA 0.1 mM, the mean depression was 49 ± 3% (n= 10, P < 0.001, t test; values are given throughout as mean ±S.E.M.); with BAPTA 0.5 mM or 1 mM and with EGTA 0.5 mM depression was, respectively, 44%± 4% (n= 5, P= 0.002, t test; Fig. 2B), 38 ± 3% (n= 10, P < 0.001, t test) and 41 ± 3% (n= 12, P < 0.001, t test). A similar degree of inhibition was observed in the mouse HMs; at recordings with BAPTA 0.1 mM in the pipette the mean depression was 51 ± 2% (n= 30, P < 0.001, t test). These results demonstrate that even at relatively strong Cai buffering (BAPTA 1 mM), a series of depolarizations caused a substantial inhibition of glycinergic IPSCs.
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Ca2+-induced potentiation of glycine-induced currents
Ca2+-dependent depression of glycinergic IPSCs could be presynaptic in origin, due to inhibition of neurotransmitter release. Alternatively, it could be induced postsynaptically if the rise in Cai caused downregulation of GlyRs expressed in HMs, i.e. these receptors exhibit properties different from those described in spinal neurones and in HEK 293 cells expressing GlyRs (Fucile et al. 2000).
To address this point, we analysed the whole-cell currents evoked by pressure application of exogenous glycine from a puff pipette positioned close to the recorded HMs. Depending on the agonist concentrations and duration of application pulses, at a holding potential (Vh) of –70 mV, glycine induced inward currents (IGly) in the range 50–1000 pA.
When the Cai was increased by repeated depolarization, systematic modulation of IGly was not observed (not shown). A previous study (Fucile et al. 2000) demonstrated that, in spinal cord neurones, conditioning pulses of glutamate induce the potentiation of responses to exogenously applied glycine. As HMs express both NMDA and Ca2+-permeable AMPA receptors (O'Brien et al. 1997; Essin et al. 2002), we used glutamate receptor ionic channels as a tool to elevate Cai during whole-cell recordings.
When a second puff pipette, containing glutamate (200 µM), was positioned close to the motoneurone's soma, a conditioning application of glutamate caused a large and reversible increase in the peak amplitude of IGly (Fig. 4A and B). No changes in the decay time of glycine-evoked currents were observed. This is in agreement with the fact that at puff application the decay time of IGly is determined by the relatively slow rate of glycine washout from the vicinity of recorded HM. On average, when recording with 0.1 mM BAPTA in the intracellular solution, after glutamate application IGly increased to 154 ± 8% (n= 6, P= 0.001, t test) and recovered to the control value (110 ± 3%) within 2–3 min (Fig. 4A–C). The extent of potentiation increased with prolongation of the conditioning glutamate pulse (Fig. 4B). Similar augmentation of IGly was observed with a conditioning application of NMDA (200 µM) dissolved in standard extracellular solution. After NMDA pulses (5–10 s), the IGly amplitude reversibly increased to 127 ± 3% (n= 3, P= 0.013, t test; Fig. 4D).
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These results indicate that glutamate- or NMDA-dependent elevation of glycine-evoked currents is a Ca2+-dependent phenomenon. They also demonstrate that glycine receptors expressed in HMs are potentiated by Cai, similarly to GlyRs previously described (Fucile et al. 2000), raising the question of the mechanism of Ca2+-dependent depression of glycinergic IPSCs shown in Fig. 2.
Presynaptic nature of Ca2+-dependent depression at glycinergic synapses
Depression of glycinergic IPSCs in HMs resembles the phenomenon called depolarization-induced suppression of inhibition (DSI), described originally for GABAergic synapses in cerebellar Purkinje cells (Llano et al. 1991). DSI is initiated postsynaptically due to a rise in Cai and expressed presynaptically as a decrease in neurotransmitter release. This phenomenon is accompanied by an increase in the failure rate (Vincent et al. 1992; Diana & Marty, 2003) and also an elevation in the paired-pulse ratio (PPR) (Yoshida et al. 2002; Diana & Marty, 2003).
To clarify whether the Ca2+-dependent depression of glycinergic IPSCs was of presynaptic origin, we analysed changes in these parameters before and after Cai elevation in HMs. To measure the PPR, two consecutive pulses separated by 50 ms were delivered to the stimulating pipette. In the majority of neurones, we observed an increase in IPSC amplitude evoked by the second stimulus (I2) with respect to the first one (I1) (Fig. 5A and B). In whole-cell recordings with 0.5 mM EGTA in the intracellular solution, the average value of the PPR, expressed as the ratio I2/I1, was 1.13 ± 0.05 (n= 10, range 0.91–1.34; Fig. 5C). When a series of depolarizing steps was delivered to the recorded neurone, the PPR significantly increased. In the first minute after depolarization-induced Ca2+ influx, PPR increased to 1.36 ± 0.08 (n= 10, P= 0.002, Wilcoxon test) and recovered to the control value within 5 min (1.11 ± 0.05; Fig. 5C). The failure rate of glycinergic IPSCs also increased after the series of depolarizing pulses. On average, the failure rate increased from 34 ± 4% to 62 ± 6% (n= 10, P= 0.002, Wilcoxon test; Fig. 5D).
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Involvement of endocannabinoids in Ca2+-dependent depression of glycinergic IPSCs
Several recent studies have demonstrated that the retrograde signalling for DSI is mediated by endo-cannabinoids, whose synthesis is stimulated by the depolarization-induced rise in Cai (Maejima et al. 2001; Ohno-Shosaku et al. 2001; Wilson & Nicoll, 2001). To clarify whether such a mechanism could also be responsible for Ca2+-dependent glycinergic DSI, we analysed the effects of the cannabinoid (CB) receptor agonist WIN55,212-2 and the CB receptor antagonist SR141716A on IPSCs.
The CB receptor agonist WIN55,212-2 (3 µM) caused decrease in IPSCs and partially or completely occluded glycinergic DSI (Fig. 7A). In 13 recorded cells, in low-Ca2+-buffering conditions (0.5 mM EGTA or 0.1 mM BAPTA), IPSC suppression by WIN55,212-2 was 35 ± 3% (P < 0.001, t test; not shown). In these cells, control DSI achieved 46 ± 3% (Fig. 7B) while, in the presence of WIN55,212-2, the DSI value was, on average, only 12 ± 5% (n= 13, P < 0.001, t test; Fig. 7B). In two cells WIN55,212-2 was not effective while in the other 11 cells the DSI values varied from –13 to +19% (Fig. 7B). Administration of WIN55,212-2 caused augmentation of the mean PPR value from 1.20 ± 0.10 to 1.45 ± 0.13 (n= 13, P < 0.001, Wilcoxon test; not shown), suggesting the presynaptic origin of glycinergic IPSC suppression by this CB receptor agonist.
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These results indicate that, in the hypoglossal nucleus, the release of glycine is modulated presynaptically through the Ca2+-induced cannabinoid machinery (Wilson & Nicoll, 2002; Freund et al. 2003; Diana & Marty, 2004), while the extent of this modulation varied for different HMs.
Similarly to the control conditions (Fig. 3A and B), in the presence of SR141716A (3–5 µM), glycinergic IPSCs exhibited depolarization-induced prolongation of decay kinetics (Fig. 7D and E). The mean decay time of IPSCs before depolarization was 9.2 ± 0.5 ms; after depolarization it was 11% longer (10.3 ± 0.6 ms; n= 16, P < 0.001, Wilcoxon test) and 5 min later it had recovered to the control value (9.1 ± 0.4 ms; Fig. 7E).
This weak but significant reversible increase of glycinergic IPSC decay time might reflect prolongation of postsynaptic glycine receptor channel openings during Cai elevation (Fucile et al. 2000). To check this hypothesis, we analysed glycinergic IPSCs when presynaptic modulation was suppressed by the CB receptor antagonist SR141716A and the Cai rise was transiently induced by activation of NMDA receptor channels.
Potentiation by NMDA of glycinergic IPSCs in the presence of a CB receptor antagonist
In the absence of a CB receptor antagonist, when the intracellular solution contained 0.1 mM BAPTA, application of NMDA (5–10 s pulses, Vh=–70 mV) resulted in a strong suppression of glycinergic IPSCs, similar to that caused by depolarization (Fig. 8A and B). On average, in six neurones analysed, the amplitude of glycinergic IPSCs after the NMDA pulse decreased to 47 ± 8% (P= 0.001, t test; Fig. 8B).
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In the presence of SR141716A (3 µM), following NMDA pulses, synaptic currents exhibited both significant increases in peak amplitudes and prolongation of decay time constants (Fig. 8D–F). Both parameters reverted almost completely to control values within 1–2 min. On average, in the first minute following NMDA application, the amplitude of IPSCs increased to 123 ± 6% of control (n= 6, p = 0.012, t test; Fig. 8E). In these experiments, to give relief from Mg2+ block, cells were concomitantly depolarized to –20 mV during NMDA applications. This facilitation of Ca2+ influx resulted in a particularly powerful prolongation of the IPSC decay time constant (by 80 ± 17%; n= 6, P= 0.006, Wilcoxon test; Fig. 8F). After the NMDA pulse, the IPSC amplitude and decay kinetics recovered to, respectively, 103 ± 5% and 119 ± 13% of control (n= 6; Fig. 8E and F).
When the same experiments were repeated with the BAPTA 20 mM internal solution, the NMDA-induced potentiation was absent: both IPSC amplitude (103 ± 6%, n= 6) and decay time constant (105 ± 4%, n= 6) measured following NMDA application were equal to control values (Fig. 8G, H and J).
Our observations demonstrate that GlyRs at glycinergic synapses can be strongly modulated by Ca2+ influx through glutamate receptor channels. This postsynaptic modulation could cause an increase in glycinergic transmission. However, in the HMs this phenomenon is masked by a powerful retrograde inhibition mediated partially by the endocannabinoid system.
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Several arguments indicate that the glycinergic DSI has a presynaptic origin and is largely mediated by the endo-cannabinoid system. First, we find that depolarization: (i) increases PPR and (ii) increases the failure rate of stimulation-induced IPSCs. Second, depolarization caused a decrease in the frequency of mIPSCs without changes in their amplitude. Third, SR141716A, a selective antagonist of the brain CB receptors, decreased glycinergic DSI without changes in IPSC amplitude. Finally, the CB receptor agonist WIN55,212-2 decreased the amplitude of IPSCs and occluded glycinergic DSI. The mechanism of Ca2+-induced feedback regulation of inhibitory and excitatory synaptic transmission via retrograde endo-cannabinoid signalling is widely used in the mammalian nervous system (Jennings et al. 2001; Wilson & Nicoll, 2001; Yoshida et al. 2002; Azad et al. 2003).
Endocannabinoids can modulate the function of ionic channels independently on CB receptors. It has been shown that the acid-sensitive potassium (K+) channel TASK-1 is directly blocked by submicromolar concentrations of anandamide (Maingret et al. 2001). Moreover, anandamide, as well as WIN55,212-2, can also inhibit the activity of glycine receptors in heterologous systems (Lozovaya et al. 2005). While for HMs, a direct modulation of ionic channels by endocannabinoids is not excluded, several arguments suggest that this mechanism plays a minor role in glycinergic DSI. First, presynaptic inhibition of K+ channels should lead to membrane depolarization and, consequently, stimulation of neurotransmitter release. In our experiments we observed the opposite effect. Second, postsynaptic inhibition by a CB receptor agonist of K+ TASK-1 channels should result in changes of both the input resistance and level of background current. However, WIN55,212-2 caused inhibition of glycinergic IPSCs without affecting the membrane properties of HMs. Third, SR141716A does not prevent the effects of anandamide and WIN55,212-2 on TASK-1 channels (Maingret et al. 2001), while in HMs this CB receptor antagonist effectively suppressed the action of WIN55,212-2. Finally, postsynaptic inhibition of ionic channels should be observed without changes in failure rate and PPR. In our experiments, these parameters were clearly modified by WIN55,212-2.
The fact that SR141716A did not completely suppress glycinergic DSI might indicate the possibility of an additional mechanism of Ca2+-induced feedback regulation. Firstly, glycinergic neurotransmission in HMs could be regulated via a putative, as yet unidentified, cannabinoid receptor. Indeed, besides the cloned G-protein-coupled CB1 and CB2, it has been suggested that other cannabinoid-sensitive receptors (CBX) are present in the brain (Di Marzo et al. 2000; Breivogel et al. 2001; Hajos et al. 2001; Hajos & Freund, 2002). Alternatively, endocannabinoid-independent retrograde inhibition via activation of GABAB or metabotropic glutamate receptors, similar to that described for glutamatergic EPSCs (Zilberter et al. 1999) or GABAergic IPSCs (Zilberter, 2000) in rat neocortex neurones, could be involved. The GABAB modulation pathway is particularly attractive as corelease of glycine and GABA from ‘mixed’ presynaptic terminals is well documented (Jonas et al. 1998; O'Brien & Berger, 1999). Moreover, in the auditory brainstem nucleus (Lim et al. 2000) and in HMs (O'Brien et al. 2004), GABA mediates presynaptic inhibition of glycinergic synapses via GABAB receptors.
Ca2+-induced potentiation
Our observations demonstrate that in glycinergic synapses, postsynaptic GlyRs are potentiated by Cai and this modulation strongly depends on the source of Ca2+ rise. Influx of Ca2+ through NMDA receptor channels was much more effective for potentiation of GlyRs than that through voltage-gated Ca2+ channels.
The organization of synaptic inputs and specificity of the cytosolic buffering system in HMs might account for these remarkable differences in Cai action. From light- and electron-microscopic observations, the vast majority (88–98%) of axonal projections on HMs forms axodendritic synapses (Zhang et al. 2001; Zhang et al. 2003). As levels of Ca2+ are regulated in a highly local environment, this would suggest that Ca2+ signals initiated in the soma and backpropogating to dendritically localized glycinergic synapses would be rapidly attenuated. Consequently, a depolarization-induced Cai rise in the vicinity of glycinergic synapses might be much weaker than those recorded in the soma. This is particularly critical for HMs, which exhibit a low concentration of endogenous Ca2+ buffers (Ca2+ binding ratio about 40) in the cytosol (Lips & Keller, 1998), which accounts for rapid relaxation times of Cai after depolarization. Analysis of Cai transients in HMs demonstrated that peak amplitudes of voltage-induced Cai rise are more than two-fold higher in the soma than in dendrites monitored 80 µm from the soma. Moreover, the decay time constant of Cai in dendrites is faster (Ladewig et al. 2003).
On the other hand, a high efficacy of Ca2+ entry through NMDA receptor channels might suggest that glutamatergic and glycinergic synapses are in close proximity on dendrites of HMs. It has been demonstrated on young brainstem slices that NMDA and non-NMDA receptors are colocalized on glutamatergic synapses in HMs; furthermore, these neurones exhibit a markedly higher proportion of NMDA receptor-mediated mEPSCs than do non-NMDA ones (O'Brien et al. 1997). On the basis of indirect evidence, the colocalization of NMDA and glycine receptors has been proposed in neurones of spinal cord (Fern et al. 1996) and brainstem hypoglossal nucleus (Berger et al. 1998). Thus, activation of dendritically situated NMDA receptors would produce a high local rise in Cai, and consequent potentiation of neighbouring GlyRs in glycinergic synapses.
The action of NMDA might be presynaptic in origin, as in some brain areas NMDA receptors are found on both excitatory and inhibitory terminals (Casado et al. 2000; Duguid & Smart, 2004). However, several lines of evidence indicate that the potentiation of IPSCs observed in our study is of postsynaptic origin and results from the increase in GlyR activity. First, NMDA similarly potentiated glycinergic IPSCs and ionic currents induced by exogenous application of glycine. Second, the increase in IPSC amplitude is associated with prolongation of decay time, as is expected for an increase in the apparent affinity of GlyR for the agonist (Fucile et al. 1999, 2000; Li & Pearce, 2000). Finally, the effect of NMDA was absent in the strong buffering of postsynaptic Ca2+.
The most relevant feature of Ca2+-induced postsynaptic potentiation is the reversible increase in duration of glycinergic IPSC decay time. We observed this, with different extents of prolongation, after a depolarization-induced Cai rise and after application of NMDA. An early study on HMs demonstrated that GlyR deactivation is the main determinant of glycinergic IPSC decay (Singer & Berger, 1999). As deactivation kinetics correlate with the apparent affinity of GlyR (Fucile et al. 2000; Li & Pearce, 2000), IPSC prolongation is consistent with a Ca2+-induced decrease in the EC50 for glycine.
Prolongation of the IPSC decay during glycinergic DSI could be of presynaptic origin. This might result from the lower probability of liberation during glycinergic DSI and, consequently, more scattered latencies of individual vesicular release events in comparison with the control. Some presynaptic effects on IPSC decay time constant are not entirely excluded, while several observations do not support this suggestion. First, the CB receptor antagonist SR141716A, which decreased DSI, should also suppress changes in IPSC's decay time. Our results indicate that modulation of glycinergic currents' kinetics was similar in the two conditions (Fig. 3A and 7D). Second, following NMDA application, similar prolongation of IPSCs was observed in control conditions when the amplitude of IPSCs was strongly depressed (Fig. 8A–C), as well as in the presence of SR141716A when the amplitude of IPSCs was augmented (Fig. 8D–F). For a presynaptic mechanism, the kinetics should follow the directions of amplitude variations. Finally, NMDA and depolarization caused similar suppression of amplitudes while NMDA-induced Cai rise resulted in much more pronounced prolongation of IPSC decay time. In the case of changes in the latencies of individual vesicular release, the effect should be similar in the two conditions.
The increase in glycinergic IPSC peak amplitude, although limited, is somewhat surprising because glycine-activated currents are not much affected by Ca2+-dependent potentiation at saturated concentrations of glycine (Fucile et al. 2000). Augmentation of IPSCs could indicate that GlyRs at glycinergic synapses of HMs are not saturated by transmitter release; this has been shown at glycinergic synapses in the zebrafish hindbrain (Rigo et al. 2003).
The double modulation of glycinergic synaptic transmission described here may be used for selective modification of neuronal activity under different physiological conditions. In particular, at early stages of postnatal development in some brain regions, such as hippocampus and neocortex, glycinergic (Ito & Cherubini, 1991) and GABAergic (Dammerman et al. 2000) responses are excitatory. Moreover, GlyRs may be activated in a non-synaptic fashion (Flint et al. 1998), suggesting that, before the establishment of mature chloride equilibrium, Ca2+-dependent potentiation of glycinergic currents potentially represents a positive feedback in the regulation of excitation, and may have a role in synaptic stabilization (Kirsch & Betz, 1998). This mechanism, however, would not be applicable for HMs as, immediately after birth, these motoneurones exhibit inhibition by synaptic glycinergic and GABAergic inputs (Marchetti et al. 2002).
Ca2+-induced potentiation might be critical at close colocalization of GlyRs with voltage-gated or receptor-operated Ca2+-permeable ion channels. This might be the case for the giant synapse of the calyx of Held, which contains presynaptic GlyRs; glycine spillover has been observed to activate these presynaptic receptors, regulating glutamate release (Turecek & Trussell, 2001). We suggest that, in a similar manner, the process of Ca2+-dependent potentiation of GlyRs might be relevant in the physiological regulation of synaptic activity.
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References |
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) and in the presence of 3–5 µM SR141716A (
). The average values are shown by 
(Control) and • (SR141716A). Note that the effect of SR141716A varied markedly in the different cells. D, superimposed traces of glycinergic IPSCs recorded in the presence of SR141716A (3–5 µM) before (Control) and after (Depo) depolarization; each trace is the average of 30 single IPSCs. Note decay time prolongation of glycinergic IPSCs after depolarization. E, averaged data from 16 neurones for decay time of IPSCs before (Control), in the first min after (Depo) and 5 min after (Recovery) depolarization. Data are mean ±S.E.M.*Significant difference (P < 0.001).
