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NEUROSCIENCE |
1 Neuroscience Center and Department of Bio- and Environmental Sciences, Physiology, PO Box 65 (Viikinkaari 1), 00014 University of Helsinki, Finland
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(Received 4 April 2007;
accepted after revision 7 June 2007;
first published online 14 June 2007)
Corresponding author S. E. Lauri: Neuroscience Center and Department of Bio- and Environmental Sciences, Physiology, PO Box 65 (Viikinkaari 1), 00014 University of Helsinki, Finland. Email: sari.lauri{at}helsinki.fi
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Introduction |
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Apart from fast effects on synaptic transmission and plasticity, increasing evidence suggests a role for KARs in synaptic development (Tashiro et al. 2003; Marchal & Mulle, 2004; Lauri et al. 2005, 2006). In particular, at hippocampal mossy fibres, kainate receptors can regulate motility of axonal filopodia (Tashiro et al. 2003), a phenomenon implicated in the early steps of synapse formation. At the same synapse, lack of KAR subunits GluR5 and GluR6 during development perturbs establishment of mature pre- and postsynaptic functions (Marchal & Mulle, 2004), supporting a role for KARs in synaptic maturation.
In the area CA1, pharmacological activation of GluR5-containing KARs strongly depresses glutamatergic transmission (Chittajallu et al. 1996; Vignes et al. 1998; Clarke & Collingridge, 2002). This depression is due to presynaptic receptors acting via a G-protein-mediated signalling mechanism to regulate glutamate release (Frerking et al. 2001; Lauri et al. 2006). Interestingly, although KARs are present and can be pharmacologically activated to depress transmission at CA3–CA1 synapses both in the neonate and young adult rats, physiological activation of these receptors has only been detected during early postnatal development (Lauri et al. 2006). Thus, in the neonatal CA1, presynaptic KARs are tonically activated by ambient glutamate and maintain a low probability of glutamate release at the immature synapses (Lauri et al. 2006). This tonic activation is down-regulated during development and in response to induction of neonatal LTP suggesting a role for presynaptic KAR activity in the maturation of glutamatergic synapses also in the area CA1.
To further understand the role of KARs in synaptic development and maturation we have here studied the effects of long-term activation and inhibition of GluR5 subunit-containing KARs on synaptic inputs to CA1 pyramidal neurons. Long-term agonist treatment in hippocampal slice cultures was used to mimic the receptor mechanism that is physiologically activated during early development. Prolonged activation of GluR5-containing KARs caused a specific and enduring increase in the number of functional glutamatergic synapses in area CA1. Further, chronic blockade of the physiological activation of GluR5 during development disturbed formation of functional glutamatergic connections to CA1 pyramidal neurons. These data suggest that GluR5 subunit-containing KARs regulate stabilization and/or formation of glutamatergic synapses in area CA1.
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Methods |
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Organotypic cultures were prepared according to the method of Stoppini et al. (1991). In brief, at postnatal day 9–10 (P9–10) Wistar rats were killed by decapitation without anaesthesia in accordance with the University of Helsinki animal welfare guidelines. The brain was rapidly removed and transferred into ice-cold preparation solution (MEM–Hepes (Gibco BRL) containing 28 mM glucose and 25 µg ml–1 chloramphenicol). Hippocampi were dissected under sterile conditions and cut transversally into 350 µm thick slices using a McIllwain tissue-chopper (Mickle Laboratory, Surrey, UK). Hippocampal slices were then transferred into the culture medium containing: 50% minimal essential medium with Hepes, 24% heat-inactivated horse serum, 24% Earle's balanced salt solution (all from Gibco BRL), 1% L-glutamine and 1% chloramphenicol. Slices were placed on Millicell-CM 0.4 µm membrane inserts (Millipore, Bedford) in 6-well culture trays with 1 ml of the above medium. The slices were cultivated at the interface of a culture medium at +35°C in 5% CO2. The medium was changed 1 day after plating, every second or third day thereafter and a day before experiments. Appropriate drugs were applied in the culture medium 16–24 h before electrophysiological recordings, except for LY382884, which was present throughout the cultivation. In these experiments, media was changed and fresh drug added every other day. Slices were used 13–15 days in vitro (DIV14).
Electrophysiology
For electrophysiological recordings, the slices were placed in a submerged recording chamber and constantly superfused with an extracellular solution containing the following (mM): 124 NaCl, 3 KCl, 1.25 NaH2PO4, 1 MgSO4, 26 NaHCO3, 15 D-glucose, 2 CaCl2 and 20 sucrose (bubbled with 5% CO2–95% O2 at 22°C). Sucrose was added to the extracellular solution to adjust osmolarity to be compatible with the slice culture medium.
Whole-cell voltage clamp recordings from CA1 pyramidal neurons were obtained by using Multiclamp 700A or Axoclamp 200B amplifier (Axon Instruments, Foster City, CA, USA). Patch electrodes (3–5 M
) contained the following (mM): 130 CsMeSO4, 10 Hepes, 0.5 EGTA, 4 Mg-ATP, 0.3 Na-GTP, 5 QX314, 8 NaCl and 20 sucrose (300 mOsmol l–1, pH 7.2). For recording of NMDA-receptor mediated responses, 10 mM BAPTA was included in the electrode filling solution.
AMPA receptor-mediated spontaneous miniature EPSCs (mEPSCs) were recorded in the presence of 1 µM TTX and 100 µM picrotoxin (PiTX) at –70 mV. NMDA receptor-mediated mEPSCs were pharmacologically isolated with TTX, PiTX and 20 µM NBQX and recorded at a holding potential of –40 mV. Spontaneous miniature IPSCs (mIPSCs) were recorded in the presence of 1 µM TTX and 20 µM NBQX at –75 mV.
AMPA receptor-mediated EPSCs were evoked with a bipolar electrode placed in CA1 stratum radiatum close to the recording site. Recordings were made at –70 mV in the presence of 50 µM D-AP-5, 100 µM PiTX and 50 nM TTX to suppress polysynaptic activity. Evoked NMDA receptor-mediated EPSCs were recorded at +40 mV in the presence of 20 µM NBQX, 100 µM PiTX and 50 nM TTX.
To activate or block GluR5-containing KARs, we used the agonist (RS)-2-amino-3-(3-hydroxy-5-tertbuty-lioxazol-4-yl) propanoic acid (ATPA, 1 µM) and antagonist (3S,4aR,6S,8aR)-6-(4-carboxyphenyl)methyl-1,2,3,4,4a,5,6,7,8,8a-decahydroisoquinoline-3-carboxylic acid (LY382884, 10 µM) (Eli Lilly, Indianapolis, IN, USA). All compounds, except LY382884, were from Tocris Cookson (Bristol, UK).
Data analysis
Miniature postsynaptic currents were analysed using the MiniAnalysis Program, version 5.6.6 (Synaptosoft, Decatur, GA, USA) and Clampfit 9.2 program (Axon Instruments). The threshold for event detection was set at two times the baseline RMS noise level and the detected events were verified visually. Evoked currents were analysed offline using Clampfit 9.2. The amplitude of the evoked synaptic responses was measured as the peak relative to the average baseline level 3–8 ms before the stimulation.
Due to large differences in the baseline frequency of spontaneous events between the slice culture batches, all the pharmacological data have been normalized to the control level, obtained from same day recordings from control cultures from the same batch. All data are expressed as percentage (%) of control (100% = no change). All the pooled data are given as mean ± S.E.M. for the number of cells indicated. Student's two-tailed t test was used for statistical analysis. P < 0.05 was considered statistically significant.
Western blotting
Organotypic slices were washed three times with cooled PBS, removed from membranes and homogenized manually in lysis buffer (150 mM NaCl, 50 mM Tris, 1 mM EDTA, 1% NP-40, 0.5% deoxycholate, 0.1% SDS). Protein concentrations were measured using Bio-Rad DC protein assay (Bio-Rad Laboratories). Protein lysates were extracted in SDS-PAGE buffer (62.5 mM Tris, 1.8% SDS, 7.75% glycerol and 4.4% 2-mercaptoethanol, pH 6.8) and boiled at +95°C for 10 min. Three independent sample homogenates of both ATPA-treated and control cultures (15 mg of protein per lane) were resolved via 4–15% SDS-PAGE gels. Proteins were transferred to Hybond nitrocellulose membrane (Amersham Biosciences) by a semi-dry blotting technique. Ponceau staining of the membrane confirmed uniform protein amounts. Membranes were blocked for 1 h with 5% milk (w/v) prepared in PBS and immunoblotted by over-night incubation at +4°C with primary antibodies: anti-synaptophysin (1: 1000, rabbit polyclonal, Zymed Laboratories Inc.), anti-
-tubulin (1: 1000, mouse monoclonal, Sigma), anti-synapsin1 (1: 1000, mouse monoclonal, Synaptic Systems), anti-VGLUT1 (1: 1000, mouse monoclonal, Synaptic Systems) or anti-GluR1 (gift from K. Keinanen, 1: 1000) horseradish peroxidase (HRP)-conjugated goat anti-rabbit IgG (Bio-Rad Laboratories) and sheep anti-mouse IgG (Amersham Biosciences) were used as secondary antibodies. The antibody complexes were detected using ECL reagents (Amersham Biosciences). Developed films were scanned and the integrated optical density of protein bands was measured and background subtracted.
Immunostaining and confocal microscopy
For immunostainings, organotypic slices were washed three times with phosphate-buffered saline (PBS, pH 7.4) and fixed overnight with 4% PFA in PBS at +4°C. Fixed slices were treated with 30%, 50% and 80% methanol (in PBS) followed by Dent's fixative (80% methanol + 20% DMSO) and TBSTD (0.1% Tween + 5% DMSO in Tris-buffered saline). Permeabilized slices were blocked with 5% bovine serum albumin (BSA) and 0.4% normal goat serum (NGS) in TBSTD for 3 h. Primary antibody (anti-synaptophysin, 1: 1000, Zymed Laboratories Inc.) were added in blocking solution and slices were incubated 45–50 h with shaking at +4°C. Slices were washed for several hours with TBSTD before incubation overnight with secondary antibody (AlexaFluor 568 goat anti-rabbit IgG 1: 400, Molecular Probes).
Blind-coded immunostained slices were analysed with confocal microscope (Zeiss LSM 5 Pascal with Axioplan 2 microscope, using a C-Apochromat 40x, N.A 1.2 water objective at 2x). The gain was optimized in the beginning and held constant through the group of sister control and ATPA-treated slices that were processed simultaneously. Digitized images were captured from the CA1 stratum radiatum close to the stratum pyramidale.
Images were analysed using ImagePro Plus 6.0 software (MediaCybernetics). The threshold function was set to distinguish the stained area from the background and the same threshold was maintained for all images obtained from the same staining. The total area of the objects above the threshold was assessed and used to estimate the number of synaptophysin-positive puncta. The average diameter of synapses was assessed using automatic particle detection analysis. Each staining included at least three (usually 5–7) control and ATPA-treated slices. After imaging and image analysis the blind code was broken and the values obtained from the ATPA-treated slices were normalized against the sister control slices from the same staining. Student's two-tailed t test was used for statistical analysis.
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Results |
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The expression pattern and function of GluR5 in hippocampal slice cultures has not been characterized previously; thus, we first examined the acute effect of ATPA, a selective agonist for the GluR5 subunit-containing KARs (Clarke et al. 1997), on glutamatergic transmission in area CA1 of the slice cultures. Slices were prepared from P9–P10 animals and whole-cell voltage clamp recordings made from the CA1 pyramidal cells at 13–15 days in vitro (DIV). Application of 1 µM ATPA caused a reversible decrease in the frequency of action potential-independent, spontaneous AMPA receptor-mediated miniature EPSCs (mEPSCs) in CA1 pyramidal cells (45 ± 7% of control, n = 7, P < 0.005) (Fig. 1A). Neither the amplitude (Fig. 1C) nor the kinetics of mEPSCs (data not shown) were altered in the presence of ATPA. ATPA had no significant effect on the holding current of these cells (–3 ± 4 pA, P = 0.15), indicating that at the concentration used (1 µM), ATPA did not activate postsynaptic ionotropic currents in the CA1 pyramidal cells. These findings show that ATPA-sensitive KARs depress action potential-independent glutamate release onto CA1 pyramidal cells in hippocampal slice cultures.
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Long-term activation of GluR5 subunit-containing KARs regulates glutamatergic input to CA1 pyramidal neurons
Since presynaptic KARs are expressed but not endo-genously activated at the CA1 region of the slice cultures (13–15 DIV), these cultures can be used as a model to study the consequences of long-term pharmacological activation of KARs on synaptic connectivity. GluR5-containing receptors were activated by ATPA (1 µM) for 16–20 h under culture conditions. We first ensured that long-term ATPA treatment was a non-toxic procedure. Consistent with previous reports (Kristensen et al. 2001), treatment of the slices with 1 µM ATPA did not cause any detectable excitotoxic cell death, assessed with propodium iodide staining (1 µg ml–1 PI applied for 30 min) (not shown).
To study the effects of long-term activation of GluR5-containing receptors on glutamatergic transmission, AMPA receptor-mediated mEPSCs were recorded from CA1 pyramidal cells from ATPA-treated and control slices (Fig. 2A). ATPA was washed out before the recordings were started. The frequency of mEPSCs in ATPA-incubated slices was significantly higher compared with control slices (281 ± 42% of control, ATPA n = 24, control n = 25, P < 0.005) (Fig. 2B). Except for the increase in frequency, mEPSCs were indistinguishable from those recorded in control slices with respect to amplitude (100 ± 12% of control) or kinetics (decay time 104 ± 8%, rise time 103 ± 5% of control) (Fig. 2Bc). To confirm that the effect on mEPSC frequency was selectively mediated by GluR5-containing KARs, slices were incubated in the presence of LY382884 (10 µM) together with ATPA. LY382884 totally blocked the ATPA-induced increase in mEPSC frequency (120 ± 20% of control, ATPA + LY382884 n = 13, control n = 11) (Fig. 2C). LY382884 alone had no significant effect on mEPSC frequency on 20 h time scale (123 ± 26% of control, LY382884 n = 15, control n = 14). Further, 16–20 h treatment with 0.3 µM AMPA had no significant effect on mEPSCs (frequency 144 ± 25% of control, amplitude 107 ± 5% of control, AMPA n = 20, control n = 22) (Fig. 2C).
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In addition to its effects on glutamatergic transmission in area CA1, ATPA depolarizes CA1 interneurons and depresses GABA release (Clarke et al. 1997; Cossart et al. 1998; Rodriguez-Moreno et al. 2000; Clarke & Collingridge, 2004; Maingret et al. 2005). To assess if long-term application of ATPA has any effects on GABAergic transmission, we next recorded spontaneous GABAA receptor-mediated miniature IPSCs (mIPSCs) after the ATPA treatment. We found no difference in the frequency (101 ± 16% of control, ATPA n = 12, control n = 12), amplitude (106 ± 4%) or kinetics (decay time 105 ± 5%, rise time 102 ± 2% of control) of mIPSCs between control and ATPA-treated slices (Fig. 2D), suggesting that GluR5 is specifically involved in the regulation of glutamatergic synaptic connectivity.
Endogenous KAR activity regulates development of glutamatergic connectivity to CA1
The GluR5 antagonist LY382884 was used to study the involvement of endogenously activated KARs in the regulation of glutamatergic connectivity. LY382884 (10 µM) had no effect on mEPSCs after 20 h application at 13–15 DIV (Fig. 2C); however, at this developmental stage endogenous synaptogenesis in the slice cultures is relatively slow and LY382884-sensitive KARs are not physiologically activated. To block GluR5-containing receptors during the entire in vitro development, LY382884 (10 µM) was added into the cultures after slice preparation and fresh drug solution was applied every other day until recording. In the slices where GluR5 was chronically blocked by LY382884, mEPSC frequency was strikingly lower as compared with control slices (54 ± 17% of control, LY382884 n = 26, control n = 25, P < 0.05), while there was no change in mEPSC amplitude (97 ± 4% of control) or kinetics (decay time 106 ± 3%, rise time 107 ± 4% of control) (Fig. 3A–C). These data show that endogenous GluR5 subunit-containing KARs are involved in the development of glutamatergic synaptic connectivity in area CA1.
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The higher frequency of mEPSCs in ATPA-treated slices could be due to changes in glutamate release probability (Pr) or insertion of AMPA receptors to ‘silent’ synapses lacking functional AMPA receptors at the postsynaptic membrane. To examine whether Pr is increased after ATPA treatment, we analysed the amplitude ratio of evoked AMPA-R-mediated EPSCs in response to paired-pulse stimulation. There was no significant difference in the paired-pulse facilitation ratio (PPR) between slices treated with ATPA as compared with control slice cultures (PPR with interpulse interval 50 ms; control 1.55 ± 0.15, n = 15; ATPA 1.39 ± 0.09, n = 15) (Fig. 4A). Further, we compared the progressive decline of evoked NMDA receptor-mediated EPSCs in the presence of an open-channel blocker (MK-801, 40 µM) in control and ATPA-treated slices, a method that has been used to assess alterations in Pr after induction of LTP and prolonged pharmacological treatments (Weisskopf & Nicoll, 1995; Lüthi et al. 2001). However, no difference in the effect of MK-801 on NMDA responses was seen between the ATPA-treated versus control slices (n = 6 for both groups) (Fig. 4B). Together, these results suggest that long-term GluR5 activation does not affect Pr.
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Receptor internalization has been shown to occur after prolonged treatment by agonists (e.g. Tsao & von Zastrow, 2000; Fairfax et al. 2004). As presynaptic GluR5-containing KARs depress glutamate release onto CA1 pyramidal cells, higher mEPSC frequency after long-term ATPA treatment could simply be due to receptor internalization or desensitization. To examine this possibility, we tested whether re-application of ATPA in slices after long-term ATPA treatment affects glutamatergic transmission. Application of 1 µM ATPA in ATPA-treated cultures caused a reversible decrease in the frequency of mEPSCs (53 ± 10% of control, n = 4, P < 0.005) (Fig. 4D), an effect that was not significantly different from that seen in control slices (P = 0.3). Thus, loss or desensitization of presynaptic KARs does not explain the ATPA-induced increase in mEPSC frequency.
Together, these data suggest that the increase in mEPSC frequency after long-term GluR5 activation is due to an increase in the number of functional synaptic contacts.
Long-term activation of GluR5-containing KARs increases synaptic density in area CA1
To assess the influence of prolonged GluR5 activation on synapse number, we immunoblotted synaptic marker proteins from ATPA-treated and control slice cultures (Calhoun et al. 1996; Lauri et al. 2003). Western blots from slice extracts showed significantly higher levels of several synaptic proteins in ATPA-treated slices as compared with controls (Fig. 5A). The most pronounced increase was observed in the levels of synaptophysin (141 ± 9% of control, P < 0.05) (Fig. 5A), while modest significant increases were observed in the expression of VGLUT1, GluR1 and synapsin1 (Fig. 5Ab). In the same samples, no significant changes in the levels of -tubulin were detected.
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GluR5 regulates synaptic density independently of the network activity and via a PKC-dependent mechanism
Endogenous GluR5-containing kainate receptors strongly regulate the characteristic synchronous network activity in the neonatal hippocampus (Lauri et al. 2005). Thus, long-term GluR5 activation might increase synapse number indirectly by inducing homeostatic mechanisms to compensate for diminished network activity in the slice cultures (e.g. Burrone & Murthy, 2003). We next examined this possibility by incubating the slices simultaneously with ATPA and tetrodotoxin (TTX, 1 µM), which blocks all the action potential-dependent neuronal activity. However, inclusion of TTX had no significant effect on the ATPA-induced increase in the frequency of mEPSCs (210 ± 29% of TTX, ATPA + TTX n = 11, TTX n = 15, P < 0.05) (Fig. 6A), indicating that GluR5 regulates synapse number independently of the network activity. As reported previously (Huupponen et al. 2007), 20 h incubation with TTX alone had no effect on the frequency (102 ± 22% of control, TTX n = 15, control n = 13) or amplitude (110 ± 28% of control) of mEPSCs.
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Presynaptic KARs regulating glutamatergic transmission in the area CA1 have been shown to signal through a G-protein-coupled mechanism (Frerking et al. 2001; Lauri et al. 2006). In the neonate hippocampus, effects of presynaptic KAR on glutamatergic transmission are also dependent on protein kinase C (PKC) (Lauri et al. 2005; Sallert et al. 2007). Therefore, we next studied whether the effect of GluR5 activation on synaptic density is dependent on PKC. The slice cultures were incubated with a selective PKC inhibitor, bisindolylmaleimide VII acetate (BIS, 0.5 µM), for 16–20 h with and without ATPA. BIS treatment alone did not significantly change mEPSC frequency as compared with control slices (135 ± 21% of control, BIS n = 9, control n = 12, P = 0.22). When ATPA was co-applied with BIS, the mEPSC frequency did not differ from BIS-treated or control slices (101 ± 2% of BIS-treated, ATPA + BIS n = 12, BIS n = 9, P = 0.96) (Fig. 6C). These data suggest that the effects of GluR5-containing KARs on synapse number are dependent on PKC activation.
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Discussion |
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Here we show that signalling via GluR5 subunit-containing KARs is critical in regulating the number of functional glutamatergic synapses to CA1 pyramidal neurons in hippocampal slice cultures. Thus, long-term (16–20 h) pharmacological activation of GluR5 subunit-containing KARs lead to dramatic increase in the frequency of action potential-independent spontaneous EPSCs (mEPSCs) in CA1 pyramidal neurons, without affecting their amplitude or kinetics. This effect persisted for at least 24 h and was specific for glutamatergic synapses since ATPA treatment had no effect on GABAA receptor-mediated mIPSCs. Further, we were unable to detect significant changes in presynaptic function after prolonged ATPA treatment showing that the observed increase in the mEPSC frequency was most likely due to an increase in the number of functional glutamatergic synapses. In line with this, we found that immunostaining against synaptic marker proteins in area CA1 was significantly higher in ATPA-treated slices as compared with controls.
The increase in mEPSC frequency was selectively mediated by GluR5-containing KARs since the effect was completely blocked when ATPA was co-applied with the GluR5 antagonist LY382884. LY382884 alone had no effect on a 20 h time scale in mature cultures. However, blocking the physiological activation of GluR5 by LY382884 throughout the slice cultivation period (14 days) led to a dramatic decrease in the mEPSC frequency as compared with sister control cultures, implying a role for endogenous GluR5 activation in the development of glutamatergic synapses. We have previously shown that presynaptic GluR5 subunit-containing receptors are tonically activated in the neonate but not in the adult hippocampus (Lauri et al. 2005, 2006). Thus the physiological activation of these receptors corresponds to the time of maturation of the glutamatergic circuitry.
Long-term alterations in the network activity have been shown to induce a variety of synaptic mechanisms to compensate for the changes in the overall activity levels (reviewed by Burrone & Murthy, 2003). Such indirect homeostatic mechanisms are unlikely to contribute here since the effect of ATPA on synaptic density was independent of the network activity and persisted when ATPA was co-applied with tetrodotoxin (TTX). TTX alone at the same time scale (20 h) did not induce changes in mEPSCs, in line with the previous findings suggesting that the homeostatic plasticity response to compensate for changes in overall levels of network activity in mature hippocampal cultures requires longer periods (> 48 h) of activity deprivation (Huupponen et al. 2007).
Which KAR population is responsible for the effect of ATPA on synaptic density? In addition to the presynaptic receptors at the glutamatergic terminals, ATPA-sensitive KARs are expressed in CA1 interneurons (Cossart et al. 1998; Rodriguez-Moreno et al. 2000; Maingret et al. 2005). These receptors act to depolarize the cell soma via an ionotropic action, while another population of KARs depress evoked IPSCs and GABA release via a G-protein-dependent mechanism (Clarke et al. 1997; Mulle et al. 2000; Rodriguez-Moreno et al. 2000; Christensen et al. 2004; Maingret et al. 2005). The effect of ATPA on synapse density was independent of action potential firing, independent of GABA receptor activation and dependent on PKC, indicating that the effect is not due to altered GABAergic transmission in response to activation of KARs at interneurons. We cannot completely rule out the possibility that chronic treatment with ATPA induces release of a transmitter or growth factor other than GABA that indirectly mediates the effect of ATPA on glutamatergic connectivity. However, the simpliest explanation for the data is that the regulation of synaptic density is mediated via activation of metabotropic GluR5 subunit-containing KARs at the glutamatergic terminals.
Previously, KARs have been shown to have a bidirectional concentration-dependent effect on filopodial motility at mossy fibres (Tashiro et al. 2003). It was proposed that at this synapse, KAR activation initially promotes contact formation between pre- and postsynaptic cells via increasing filopodial motility due to an apparently ionotropic mechanism that involves voltage-gated calcium channels. Later on, KARs stabilize the contacts by inhibiting the motility via a G-protein-dependent mechanism. The properties of presynaptic KARs in CA1 synapses are different from those in mossy fibres in several respects; for example, the ionotropic KAR mechanism that facilitates glutamate release at the mossy fibres has not been detected in CA1 (Frerking & Nicoll, 2000; Kullmann, 2001; Lerma, 2003). However, it is possible that similar G-protein-dependent signalling mechanisms underlie the KAR-dependent synaptic stabilization both in areas CA3 and CA1. Even though the time of intense synaptogenesis in hippocampal cultures is restricted to the first 2 weeks in vitro, morphological development in slice cultures continues until 21 DIV (De Simoni et al. 2003). Therefore, an increase in the glutamatergic connectivity in response to GluR5 activation could be a consequence of stabilization of immature synapses, without the requirement of an active role of GluR5 in synapse induction.
In conclusion, our results suggest that the activity of presynaptic GluR5-containing receptors is involved in the formation and/or stabilization of functional glutamatergic synapses in area CA1. These receptors are endogenously activated in the developing but not adult hippocampus (Lauri et al. 2006), implying that their physiological role might be specifically linked to the activity-dependent maturation of the circuitry.
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