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Department of Physiology, School of Medicine, Xi'an Jiaotong University, Zhuque Dajie 205, Xi'an, Shannxi, People's Republic of China
Submitted 13 September 2006; accepted in final form 4 November 2006
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ABSTRACT |
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INTRODUCTION |
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first demonstrated that, in the vertebrate CNS, exogenously applied glycine potentiates the response of N-methyl-D-aspartate receptors (NMDA-Rs) to NMDA. Binding of glycine to the glycine binding site of NMDA-Rs is a prerequirement for NMDA-R activation by glutamate (Kleckner and Dingledine 1988). Thus glycine and glutamate are co-agonists for NMDA-Rs.
It is generally assumed that the glycine binding site is constantly saturated due to high concentrations of glycine in cerebrospinal fluid (CSF) (Ferraro and Hare 1985). However, glycine in the synaptic cleft is subject to a powerful uptake system (i.e., glycine transporter 1 and 2, GlyT1 and GlyT2) (Bergeron et al. 1998; Betz et al. 2006; Chen et al. 2003; Supplisson and Bergman 1997). In addition, NMDA-R isoforms exhibit different affinities to glycine (Kew et al. 1998; Kutsuwada et al. 1992). A surge of in vivo and in vitro studies have suggested that saturation of the glycine binding varies among different brain areas. Some studies have shown that the application of exogenous glycine increases NMDA-R responses (Ahmadi et al. 2003; Baptista and Varanda 2005; Berger et al. 1998; Czepita et al. 1996), whereas others have found no modulatory effect (Fletcher et al. 1989; Obrenovitch et al. 1997).
D-serine, which is a potent agonist of the glycine binding site (Kleckner and Dingledine 1988; Martineau et al. 2006; Schell et al. 1995) but is not taken up by glycine transporters (Supplisson and Bergman 1997), has been shown to potentiate NMDA synaptic currents (Baptista and Varanda 2005; Mothet et al. 2000). The ability of glycine or D-serine to potentiate NMDA-R responses clearly indicates that the glycine binding site of NMDA-Rs is not saturated.
To date, a number of studies have shown that the NMDA-Rs are present on the presynaptic membrane in addition to the postsynaptic area. Presynaptic NMDA-Rs have been found to enhance neurotransmitter release in different regions of the CNS including the spinal cord, cerebellum, entorhinal cortex, and neocortex (Bardoni et al. 2004; Berretta and Jones 1996; DeBiasi et al. 1996; Glitsch and Marty 1999; MacDermott et al. 1999; Yang et al. 2006). They may also mediate synaptic plasticity (i.e., LTP and LTD) (Casado et al. 2002; Duguid and Sjostrom 2006; Sjostrom et al. 2003). Immunocytochemical studies have demonstrated that presynaptic NR1 and NR2B-containing NMDA-Rs are present in rat visual cortex (Aoki et al. 1994). Moreover, presynaptic NMDA-Rs enhance neurotransmitter release in the visual cortical layer V neurons as proven by application of APV, a competitive antagonist of these receptors (Sjostrom et al. 2003). However, whether the glycine binding site of presynaptic NMDA-Rs may modulate glutamatergic neurotransmission in CNS is still uncertain.
In this study, we examined whether the glycine binding site of presynaptic NMDA-Rs regulates glutamate release in the rat visual cortex. Our results show that the glycine binding site on the postsynaptic NMDA-Rs is not saturated and that glycine binding sites at presynaptic NMDA-Rs regulate glutamate release in the layer II/III pyramidal neurons of the rat visual cortex.
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METHODS |
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Visual cortical slices were prepared from Sprague–Dawley rats aged 13–15 days. All animals were housed in a standard environment on a 12/12-h light/dark cycle with light on at 07:00, and they were allowed ad libitum access to water and food. The use and care of animals in this study follow the guidelines of the Xi'an Jiaotong University Animal Research Advisory Committee. Rats were initially anesthetized with ether and then immersed in ice-cold water, with the nose exposed, for 3 min to reduce brain temperature. Immediately after decapitation, the brain was placed in an ice-cold artificial cerebrospinal fluid (ACSF) bubbled with 95% O2-5% CO2 (pH 7.4). The ACSF consisted of (in mM) 124 NaCl, 5 KCl, 1.2 KH2PO4, 1.3 MgSO4, 2.4 CaCl2, 26 NaHCO3, and 10 glucose, and it had an osmolality of 305–310 mosM/kg H2O. A block of tissue containing the primary visual cortex was cut into 350-µm-thick slices with a vibratome (Campden Instruments, London, UK). Slices were transferred to an incubating chamber containing ACSF equilibrated with carbogen (95% O2-5% CO2) and incubated for 
1.5 h at room temperature (20°C) prior to electrophysiological recording.
Recordings
For recording, slices were individually transferred to a recording chamber where they were perfused (2.5–3 ml/min) with oxygenated Mg-free ACSF at 31 ± 0.5°C. The temperature of the recording chamber was continuously monitored and controlled by a custom-made temperature controller. The slices were placed on an upright infra-red video microscope with differential interference contrast (DIC) optics (OLYMPUS BX51WI), which was mounted on a Gibraltar X–Y table. Slices were observed through a x40 water-immersion objective using an infrared-sensitive camera (DAGE-MTI, IR-1000). Layer II/III pyramidal neurons of the visual cortex were visually selected (Mason and Larkman 1990). Patch-clamp recordings were performed in the whole cell configuration. Unpolished and uncoated patch pipettes (1.5 mm/1.1 mm; Sutter Instruments, Novato, CA) with a resistance of 4–6 M
were pulled using a horizontal puller (model P-97, Sutter Instruments, Novato, CA). The pipette solution contained (in mM) 130 cesium methanesulfonate, 5 NaCl, 5 QX-314, 2 MgCl2, 0.1 CaCl2, 1 EGTA, 10 HEPES, 2 Na2ATP, and 0.25 Na3GTP (pH 7.3–7.4, adjusted with CsOH, 280–290 mosM/kg H2O). To block postsynaptic NMDA-Rs, the following pipette solution was used (in mM): 124 cesium methanesulfonate, 5 QX-314, 2 MgCl2, 10 BAPTA, 1 (+)-MK-801, 10 HEPES, 2 Na2ATP, and 0.25 Na3GTP (pH 7.3–7.4, adjusted with CsOH, 280–290 mosM/kg H2O). To facilitate the blockade of the open channel blocker MK-801, neurons were depolarized to –10 mV for 10 s at intervals during a 10-min period after breakthrough to whole cell access, and this interval might be enough for MK-801 to block all the channels according to Yang's work (Yang et al. 2006). Currents were recorded at a holding potential of –60 mV using an Axopatch 700B amplifier (Axon Instruments, USA) and digitized using a data-acquisition board (Digidata 1322A) operated by pCLAMP 9.2 software (Axon Instruments). They were filtered at 2 kHz and digitized at 10 kHz. Compensation for the series resistance was not employed. Statistical comparisons were performed only when series resistance was <20 M and did not change by >10%.
Glycine, D-serine, strychnine, picrotoxin, 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), D-2-amino-5-phosphonovalerate (D-APV), (+)-MK-801, QX-314, BAPTA, and 7-chloro-kynurenic acid (7-Cl KYNA) were purchased from Sigma. Tetrodotoxin (TTX) was obtained from Hebei Province Marine Science and Technolog, China. TTX (0.5 µM), strychnine (1 µM) and picrotoxin (100 µM) were added to Mg-free ACSF to suppress the sodium currents, glycinergic and GABAergic transmission, respectively. Mg-free ACSF was obtained by omitting MgSO4 (1.3 mM) from the ACSF without compensation for the loss of osmolarity or for the amount of divalent ions. All drugs were applied at known concentrations by changing the perfusion line.
Data analysis
Miniature excitatory postsynaptic currents (mEPSCs) were detected and analyzed with MiniAnalysis software (Version 6.0, Synaptosoft) using a threshold-crossing criterion. Although the threshold level varied from neuron to neuron, it was always the same before and after drug application (Bradaia and Trouslard 2002). Detection criteria also included rise time <3 ms (Sjostrom et al. 2003), which may eliminate the purely NMDA-R-mediated mEPSCs because the average 10–90% rise time of purely NMDA-R-mediated mEPSC events is 8 ± 2 ms (O'Brien et al. 1997). This means that our data do not include the silent synapses. Multiple peak events were discarded. Averaged mEPSCs were obtained by aligning events on the rising phase. Events occurring during 120 s were averaged and analyzed in every group. To quantify the NMDA component of mEPSCs, we integrated the average mEPSCs waveforms occurring over a 90-ms time period, which began 10 ms after the start of the mEPSCs (Berger et al. 1998). This integral was used as an index of NMDA-R-mediated charge transfer. Comparison of mEPSC distributions was performed using the Kolmogorov-Smirnov test, and values of P < 0.01 were accepted as significant (Berretta and Jones 1996). Group means were compared using a paired Student's t-test, and P < 0.05 was considered significant. All data are expressed as means ± SE.
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RESULTS |
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Whole cell mEPSCs recordings were performed on 58 pyramidal neurons in layer II/III of the visual cortical slices. Spontaneous mEPSCs in the presence of TTX (0.5 µM), strychnine (1 µM), and picrotoxin (100 µM) were abolished by application of D-APV (50 µM) and CNQX (10 µM) in the Mg-free ACSF (data not shown). This indicates that the mEPSCs were composed of both NMDA and non-NMDA glutamatergic components (Berger et al. 1998; O'Brien et al. 1997). Application of exogenous glycine (100 µM) enhanced the NMDA-R-mediated mEPSCs, and this effect was completely blocked by D-APV (50 µM; Fig. 1, A and B). This enhancement of NMDA-R-mediated mEPSCs or increased NMDA charge transfer (fC), is shown in Fig. 1C. Charge transfers under control conditions and in response to glycine treatment were 622 ± 111 and 795 ± 104 fC, respectively (28% increase, P < 0.007, n = 8). These results indicate that the glycine binding site on synaptically active NMDA-Rs is not saturated. However, application of glycine (100 µM) did not alter the frequency or the peak amplitude of the mEPSCs (Fig. 1, D and E). Application of D-APV (50 µM) decreased the frequency (control, 2.82 ± 0.5 Hz vs. treated, 1.50 ± 0.5 Hz, P < 0.001, n = 8) but not the peak amplitude of the mEPSCs (control, 46.9 ± 4.4 pA vs. treated, 46.0 ± 4.6 pA, P > 0.18, n = 8). This result suggests that glycine binding site on presynaptic NMDA-Rs is saturated and the NMDA-Rs may tonically regulate glutamate release.
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). Therefore we examined the effect of D-serine on NMDA-mediated mEPSCs to verify the results obtained with glycine. Application of exogenous D-serine (100 µM) also increased the NMDA-R-mediated component of mEPSCs, which was abolished by D-APV (50 µM; Fig. 2, A and B). NMDA-mediated charge transfer increased from 631 ± 83 fC in the control to 960 ± 186 fC in D-serine-treated neurons (52% increase, P < 0.005, n = 9, Fig. 2C). Thus D-serine had a greater effect on the glycine binding site of NMDA-Rs than glycine. However, D-serine did not alter the frequency or the peak amplitude of the mEPSCs (Fig. 2, D and E). Application of D-APV reduced the frequency (control, 2.27 ± 0.6 Hz vs. treated, 0.98 ± 0.3, P < 0.001, n = 9) but not the peak amplitude (control, 40.3 ± 3.6 pA vs. treated, 39.4 ± 3.5 pA, P > 0.21, n = 9) of the mEPSCs. These results verified that D-serine has the same effect as glycine.
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To determine whether the glycine binding site of the NMDA-Rs may be involved in the modulation of glutamate release, we employed 7-Cl KYNA, an antagonist of the glycine binding site of the NMDA-Rs. As shown in Fig. 3D, inclusion of 7-Cl KYNA (20 µM) in the extracellular solution blocked the postsynaptic NMDA component and reduced the frequency of the mEPSCs (control, 1.9 ± 0.7 Hz vs. treated, 1.1 ± 0.4 Hz, P < 0.001, n = 10). Combined treatment with glycine (100 µM) and 7-Cl KYNA reversed this effect (control, 1.9 ± 0.7 Hz vs. treated, 1.8 ± 0.6 Hz, P > 0.08, n = 10) and recovered the postsynaptic NMDA component (Fig. 3, B and D). As shown in Fig. 3C, the peak amplitude of the mEPSCs was not influenced by 7-Cl KYNA or glycine (control, 41.0 ± 3.3 pA; 7-Cl KYNA, 39.6 ± 4.0 pA; 7-Cl KYNA + glycine, 40.4 ± 3.9 pA, n = 10).
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It is possible that 7-Cl KYNA reduced the frequency of the mEPSCs by acting on the glycine binding site of presynaptic NMDA-Rs or postsynaptic NMDA-Rs. In the latter case, presynaptic neurotransmitter release could be affected through retrograde messengers. To test these possibilities, we included the open-channel NMDA-R blocker, dizocilpine maleate (MK-801; 1 mM), in the recording pipette as described by Berretta and Jones (1996). Blockade of the postsynaptic NMDA-Rs with MK-801 completely abolished the NMDA component of mEPSCs (Fig. 5). Inclusion of 7-Cl KYNA (20 µM) under these conditions reduced the frequency of the mEPSCs. Moreover, this effect could be reversed by the addition of glycine (100 µM; control, 2.6 ± 0.5 Hz; 7-Cl KYNA, 1.7 ± 0.3 Hz; 7-Cl KYNA + glycine, 2.6 ± 0.6 Hz; P < 0.001, n = 10; Fig. 5C). As before, the peak amplitude of the mEPSCs was not affected by treatment with 7-Cl KYNA and glycine. (control, 34.3 ± 1.5 pA; 7-Cl KYNA, 34.4 ± 1.2 pA; 7-Cl KYNA + glycine, 34.5 ± 1.7 pA; n = 10).
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DISCUSSION |
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As described in the preceding text, application of exogenous glycine potentiated NMDA-R-mediated mEPSCs. Glycine may achieve this effect through at least three mechanisms. First, glycine might act through presynaptic, strychnine-sensitive glycine receptors to facilitate the release of glutamate, an action that has been demonstrated in the brain stem (Turecek and Trussell 2001). In this study, the contribution of such as mechanism is probably limited because mEPSCs were recorded in the presence of strychnine (1 µM) and picrotoxin (100 µM). Second, glycine might act on the NMDA-Rs containing NR3A or NR3B subunits (Chatterton et al. 2002), which are activated by glycine in the absence of glutamate. The finding that D-serine, which blocks NMDA-Rs containing NR3A or NR3B subunits, potentiates NMDA-R–mediated mEPSCs argues against this possibility. Third, glycine might act postsynaptically by activating the glycine binding site of NMDA-Rs. To test this hypothesis, we employed D-serine, another NMDA-R glycine binding site agonist. The ability of D-serine to increase the NMDA component provides evidence that glycine potentiates NMDA-R–mediated mEPSCs via the NMDA-R glycine binding site. In addition, the non-NMDA component was not influenced by glycine or D-serine, and the application of APV or 7-Cl KYNA abolished the NMDA component of the mEPSCs. Taken together, these results suggest that the glycine binding site on the postsynaptic NMDA-Rs is not saturated as has been shown in cat visual cortex (Czepita et al. 1996).
The levels of synaptic glycine are known to be tightly controlled by glycine transporters. There are two types of glycine transporters, GlyT1 and GlyT2. GlyT1, which is localized to glia and neurons, is closely associated with the NMDA-Rs (Cubelos et al. 2005); whereas, GlyT2 is colocalized with strychnine-sensitive glycine receptors (Gomeza et al. 2003). As has been described in the hippocampus (Watanabe et al. 1992), D-serine was capable of potentiating NMDA synaptic currents. Here, application of exogenous D-serine increased the NMDA component by 
52%, and glycine increased the NMDA component of the mEPSCs by 28%. The greater effect of D-serine, which is not taken up by glycine transporters (Supplisson and Bergman 1997), suggests that levels of synaptic glycine are relatively low and are tightly controlled by glycine transporters.
Our results also show that APV or 7-Cl KYNA reduce the frequency but not the peak amplitude of mEPSCs, regardless of the presence of MK-801 in the recording pipette. These findings support the existence of presynaptic NMDA-Rs and are consistent with studies showing that presynaptic NMDA-Rs tonically facilitate glutamate release as a result of increased calcium influx into the terminals (Berretta and Jones 1996; Sjostrom et al. 2003; Yang et al. 2006).
It was reported that mEPSCs events rely on presynaptic Ca2+ transient either in hippocamcal area CA1 or in layer II neurons of the rat barrel cortex (Rusakov 2006; Simkus and Stricker 2002). Presynaptic Ca2+ transient could produce Ca2+ release from stores, named Ca2+-induced Ca2+ release (CICR), and then the postsynaptic EPSCs. The amplitude of CICR from internal Ca2+ stores depends on the amount of Ca2+ influx and probably on the cytosolic Ca2+ concentration (Rusakov 2006). Activation of presynaptic NMDA-Rs could induce the Ca2+ influx and then improve the CICR and finally facilitate the neurotransmitter release.
The difference in saturation of the glycine site between pre- and postsynaptic NMDA-Rs seems to have different affinities for glycine (Kew et al. 1998; Kutsuwada et al. 1992). There is strong evidence that at P14 the postsynaptic NMDA receptors are mostly NR1-NR2A, whereas the presynaptic ones are mostly NR1-NR2B (Sjostrom et al. 2003; Woodhall et al. 2001; Yang et al. 2006). The EC50s of glycine and of D-serine for NR1-NR2A receptors are known to be higher than those for NR1-NR2B. Thus a concentration of glycine will be saturating for the (presynaptic) NR2B-containing receptors and not saturating for the (postsynaptic) NR2A-containing receptors. Also consistent with our findings, presynaptic NR1- and NR2B-containing NMDA-Rs are known to exist (Aoki et al. 1994) and enhance neurotransmitter release in the visual cortex (Sjostrom et al. 2003).
The ability of glycine or D-serine to reverse the effects of 7-Cl KYNA when MK-801 was present in the recording pipette argues against the possibility that glycine acts on postsynaptic receptors to facilitate glutamate release via retrograde messengers. Additionally, the finding that D-serine is able to reverse the effects of 7-Cl KYNA, combined with the fact that strychnine was present in all experiments, makes it unlikely that presynaptic, strychnine-sensitive glycine receptors facilitated glutamate release (Turecek and Trussell 2001).
In summary, our data show that glycine can act on glycine binding sites at presynaptic and postsynaptic NMDA-Rs to enhance the NMDA-R function, which could be indicative of a general role for the glycine binding site of presynaptic NMDA-Rs in regulating glutamate release in the CNS. Moreover, these findings may be clinically relevant to schizophrenia, where enhancing NMDA-R function is considered to be a promising strategy for treatment of the disease (Coyle et al. 2002; Duncan et al. 1999).
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GRANTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: T.-Z. Han, Dept. of Physiology, School of Medicine, Xi'an Jiaotong University, Zhuque Dajie 205, Xi'an, Shaanxi 710061, PR China (E-mail: htzhen{at}mail.xjtu.edu.cn)
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ABSTRACT
INTRODUCTION
