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Department of Anatomy and Cell Biology, Indiana University School of Medicine, Indianapolis, Indiana
Submitted 4 November 2005; accepted in final form 9 December 2005
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ABSTRACT |
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INTRODUCTION |
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; Rothman and Olney 1986). Microdialysis studies have shown that extracellular glutamate concentration dramatically increases during ischemia and rapidly returns to the baseline levels after reperfusion (Benveniste et al. 1984; Globus et al. 1988). However, changes in extracellular glutamate might not faithfully reflect the changes in the synaptic cleft (Obrenovitch et al. 2000). Both in vivo and in vitro studies have shown that excitatory synaptic transmissions in the hippocampus and the neostriatum are depressed during ischemia despite the extensive accumulation of extracellular glutamate (Gervitz et al. 2001; Sugahara et al. 2001; Tanaka et al. 2001). On the other hand, the excitatory synaptic transmission of CA1 neurons in the hippocampus is facilitated after reperfusion (Crépel et al. 1993; Gao et al. 1998b).
The neostriatum is one of the brain regions highly sensitive to cerebral ischemia. Most of the medium spiny neurons in the striatum die in 24 h after transient forebrain ischemia, whereas the large aspiny neurons in the same area survive the insult (Chesselet et al. 1990; Francis and Pulsinelli 1982). The mechanisms underlying this selective cell death remain to be elucidated. Excitatory synaptic transmission has attracted much attention due to its important roles in excitotoxicity. It has been shown in vitro that aglycemia induces depression of excitatory synaptic transmission in spiny neurons whereas oxygen and glucose deprivation (OGD) induces long-term potentiation (Calabresi et al. 1997a,b, 2002). Studies using intracellular recording in vivo indicate that the excitatory monosynaptic potentials are depressed but the excitatory polysynaptic potentials are facilitated after transient cerebral ischemia (Gajendiran et al. 2001). Interpretation and reconciliation of these results are complicated for several reasons. For in vitro preparations, the neuronal responses to OGD might differ from those to ischemia in vivo because of the dramatic difference in temperature, microenvironment, and homeostasis. In addition, the results from in vitro preparations can only reveal the changes during or shortly after OGD due to the limitations of experimental preparation. For in vivo preparations, the mixture of inhibitory and excitatory components of synaptic transmission and the difficulty in pharmacological manipulations make it very hard to clearly elucidate the changes of each component to the ischemic insults. In addition, the postsynaptic potential obtained in vivo using current-clamp recording makes it difficult to evaluate the synaptic efficacy because of the existence of somatodendritic voltage-dependent conductance (Magee and Johnston 1995; Stuart and Sakmann 1995). To circumvent these restrictions, Pang et al. (2002) used whole cell voltage-clamp recording on brain slices prepared at different intervals after ischemia in vivo and showed that excitatory synaptic currents are significantly reduced in ischemia-resistant large aspiny neurons after reperfusion, suggesting that the depression of excitatory transmission might contribute to the neuronal survival after ischemia. If this is the case, the excitatory synaptic transmission should be potentiated in ischemia-vulnerable medium spiny neurons after ischemia. To test this hypothesis, we examined the excitatory synaptic transmission in spiny neurons at 9 h after transient forebrain ischemia using whole cell voltage-clamp recording on striatal slices.
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METHODS |
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Transient forebrain ischemia
Transient forebrain ischemia was induced using the four-vessel occlusion method (Pulsinelli and Brierley 1979) with modifications (Ren et al. 1997). The animals were fasted overnight to provide uniform blood-glucose levels. For surgical preparation, the animals were anesthetized with a mixture of 1–2% halothane, 33% O2, and 66% N2 via a gas mask placed around the nose. A silicon tube loop was placed loosely around each common carotid artery to allow subsequent occlusion of these vessels. The animal was then placed on a stereotaxic frame, and the vertebral arteries were electrocauterized. A tiny temperature probe (0.025-in diam; Physitemp, Clifton, NJ) was inserted beneath the skull in the extradural space, and the brain temperature was maintained at 37°C with a heating lamp using a temperature-control system (BAT-10; Physitemp). Glass microelectrodes (5–8 µm tip diameter) filled with 2 M NaCl were used to record ischemic depolarization, which is an indication of complete ischemia (Ren et al. 1997). A burr hole was drilled at 9.5 mm anterior to the interaural line, 3.0 mm from the midline. The microelectrode was advanced 3.0 mm below dura into the neostriatum. The recordings were performed with a neuroprobe amplifier (Model 1600; A-M Systems, Carlsborg, WA). The duration of ischemic depolarization was determined by measuring the period from the beginning of the extracellular direct-current potential reaching –20 mV to the point where the potential started to repolarize after recirculation. Transient forebrain ischemia was induced by occluding both common carotid arteries to induce ischemic depolarization for 
22 min. Cerebral blood flow resumed immediately on releasing of the carotid artery clasps.
Slice preparation and whole cell voltage-clamped recording
Brain slices were prepared from animals before ischemia and at 9 h after reperfusion as described previously (Pang et al. 2002). The animals were anesthetized with ketamine-HCl (80 mg/kg ip) and decapitated. The brains were quickly removed and immersed in ice-cold artificial cerebrospinal fluid (ACSF), which was composed of (in mM) 130 NaCl, 3 KCl, 2 CaCl2, 2 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, and 10 glucose. For Ca2+-free medium, Ca2+ was replaced by Mg2+, and 3 mM EGTA was included. Transverse striatal slices of 300 µm thickness were cut using a vibratome (VT 1000S; Leica, Nussloch, Germany) and incubated in ACSF for 
1 h at 35°C before being transferred to the recording chamber. The slice was submerged beneath the fluid surface and superfused continuously with oxygenated ACSF. The flow rate was adjusted to 2–3 ml/min. Recordings were performed at 33–35°C.
For whole cell recording, patch electrodes were prepared from borosilicate glass (Warner Instruments, Hamden, CT) using a horizontal electrode puller (P-97; Flaming/Brown; Sutter, Novato, CA) to produce tip openings of 1–2 µm (3–5 M
). Electrodes were filled with an intracellular solution containing (in mM) 43 CsCl, 92 CsMeSO4, 5 TEA, 2 EGTA, 1 MgCl2, 10 HEPES, and 4 ATP (Sigma, St. Louis, MO). Neurobiotin (2%; Vector Laboratories, Burlingame, CA) was included in some experiments to verify the neurons recorded. Neurons were visualized with an infrared-differential interference contrast (DIC) microscope (BX 50 WI; Olympus Optical, Tokyo) and a CCD camera. Only those with medium somata (10–15 µm) were selected for recording. Positive pressure was applied to the recording pipette as it was lowered into the medium and approached the cell membrane. Constant negative pressure was applied to form the seal (>1 G) when the recording pipette attached to the membrane. A sharp pulse of negative pressure was applied to rupture the cell membrane and access whole cell configuration. The series resistance of the pipette was 10 M. Voltage-clamped recording was performed with an Axopatch 200 B amplifier (Axon Instruments, Foster City, CA). Signals were filtered at 2 kHz and digitized at a sampling rate of 5 kHz using a data-acquisition program (Axograph 4.9; Axon Instruments). Intrastriatal stimulation was delivered every 10 s using a bipolar tungsten electrode (Micro Probe, Potomac, MD), and 0.1-ms current pulses were used to evoke the excitatory responses. One to five times of threshold stimulus intensity (1-5T) was used in the present experiments. For paired-pulse test, two stimuli with the same intensity (2T) and duration (100 µs) were applied at an interval of 50 ms. The Axograph 4.9 Program was used to do miniature analysis. Individual synaptic events with fast onset and exponential decay kinetics were captured with template detectors in Axograph 4.9 software. The detection parameters for analyzing synaptic events in each cell in the absence or presence of drugs were the same. The values were presented as means ± SE. Paired or unpaired Student’s t-test was used to detect statistical difference in most of the results and Kolmogorov-Smirnov test was used to compare the miniature EPSC (mEPSC) amplitude before and after ischemia (Statview 5.0, SAS Institute). Changes were considered significant if P < 0.05.
In some experiments, neurobiotin was iontophoresed into the cell by passing depolarizing pulses after recording (Deng et al. 2005). The slice was then fixed in 4% paraformaldehyde overnight and incubated in 0.1% horseradish peroxidase-conjugated avidin D (Vector Laboratories) in 0.01 M potassium PBS, pH 7.4, with 0.5% Triton X-100 for 24 h at room temperature. After detection of peroxidase activity with 3,3'-diaminobenzidine, slices were examined in potassium PBS. Slices containing labeled neurons were mounted on gelatin-coated slides and processed for light microscopy.
Drug application
(–)-Bicuculline methiodide (BIC), (–)-2-amino-5-phosphonopentanoic acid (APV), 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), and pyridoxal-phosphate-6-azophenyl-2',4'-disulfonic acid (PPADS) were purchased from Sigma. Antagonists were applied via bath superfusion. BIC was used to block GABAA receptors at a concentration of 30 µM, and 50 µM APV was used for N-methyl-D-aspartate (NMDA) receptor (NMDAR) blockade. Exogenous glutamate was applied through a Y-tube system (Pang et al. 2002). The tip of the Y-tube had a diameter between 100 and 150 µm and was placed close to the recorded neuron.
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RESULTS |
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Whole cell voltage-clamp recordings were performed on the striatal neurons with medium-sized somata in brain slices. To ensure the recorded neurons being medium spiny neurons, we labeled the neurons randomly with neurobiotin after recording. As shown in Fig. 1A, the labeled neuron had a medium-sized cell body (15 µm) and the large numbers of spines on their dendrites. Intrastriatal stimulation evoked both EPSCs and inhibitory postsynaptic currents (IPSCs) from medium spiny neurons (Kita 1996). To isolate EPSCs, bicuculline (30 µM) was added to the bath solution to block GABAA receptors. Both AMPA receptor (AMPAR)- and NMDAR-mediated components contribute to EPSCs in spiny neurons. Inward EPSCs were evoked at a holding potential of –70 mV. Bath application of APV (50 µM) slightly decreased the peak amplitude of EPSCs, whereas CNQX (30 µM) completely blocked the rest part in a reversible manner (Fig. 1B), indicating that eEPSCs in spiny neurons in our experimental condition (2 mM MgCl2 in external solution and holding potential of –70 mV) were mediated largely by AMPA receptors. Therefore the present study focused on changes in AMPAR-mediated EPSCs in spiny neurons after ischemia. APV (50 µM) and bicuculline (30 µM) were applied throughout the experiments unless otherwise noted.
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Presynaptic mechanism is involved in the postischemic potentiation of EPSCs
Enhancement of evoked synaptic transmission after ischemia could be due to the alterations in pre- or/and postsynaptic function. To examine the locus of ischemia-induced enhancement, we analyzed mEPSCs in the presence of 1 µM TTX. mEPSCs in the spiny neurons were measured under voltage clamp at a holding potential of –70 mV. The mEPSCs can be completely blocked by bath application of CNQX (30 µM), confirming them to be AMPA/kainite receptor-mediated events. As shown in Fig. 2, in control neurons (n = 16), mEPSCs had a mean frequency of 5.82 ± 0.66 Hz and a mean amplitude of 11.51 ± 0.57 pA. In the spiny neurons subjected to ischemia (n = 45), the frequency and amplitude of mEPSCs was increased to 8.83 ± 0.63 Hz, and 14.46 ± 0.31 pA, respectively (P < 0.05). Analysis of kinetics of mEPSCs before and after ischemia showed that there were no changes in rising time (1.02 ± 0.06 ms in control; 0.98 ± 0.07 ms after ischemia; P = 0.73) and decay time constants (3.31 ± 0.18 ms in control; 3.57 ± 0.29 ms after ischemia; P = 0.54). No detectable changes in spontaneous EPSCs frequency were observed before and after TTX (1 µM) application in either control or postischemic spiny neurons (data not shown), suggesting no spontaneous action potential mediated event in spiny neurons.
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To further rule out the possible contribution of postsynaptic mechanism to the potentiation of eEPSCs after ischemia, postsynaptic responses were examined by focal application of exogenous glutamate in the presence of 50 µM APV. Application of glutamate induced an inward current at a holding potential of –70 mV. The AMPA receptors rapidly desensitize (Trussell et al. 1988), preventing us from accurately measuring the peak amplitude in slices. Therefore we measured the steady-state currents in control and postischemic neurons. To eliminate the impact of cell size, currents were normalized by cell capacitance and expressed as current densities. Glutamate (100 µM) induced the inward currents of 19.4 ± 5.0 and 23.7 ± 4.0 pA/pF in control (n = 7) and postischemic neurons (n = 7), respectively (P = 0.51. Fig. 3, C and D). Likewise, no significant differences in current densities were found between control (128.6 ± 16.6 pA/pF, n = 7) and ischemia groups (151.4 ± 18.2 pA/pF, n = 7) in response to 500 µM glutamate (P = 0.36. Fig. 3, C and D).
Postischemic increase in mEPSCs amplitude do not result from the increased mEPSCs frequency
It is worthwhile to point out that despite the lack of involvement of postsynaptic mechanism, the amplitude of mean mEPSCs was increased after ischemia. This is an unconventional phenomenon since mEPSCs amplitude reflects the postsynaptic responses to single vesicle release. Interestingly, several previous studies have also shown the similar results and suggested that the increase in mEPSCs amplitude may arise from a random superimposition of the more frequent mEPSCs (Sharma and Vijayaraghavan 2003; Shigetomi and Kato 2004). If the increase in mEPSCs amplitude was caused by random superimposition of mEPSCs, more frequent mEPSCs should create more large-amplitude events, which in turn increase the mean amplitude of mEPSCs. Consequently, we should observe a positive correlation between the increase in frequency and mean amplitude of mEPSCs (Shigetomi and Kato 2004). However, there was no correlation between the relative frequency and mean amplitude in postischemic spiny neurons (Fig. 4A). Examination of rise time of the mEPSCs after ischemia and their relation to amplitude also suggests that the increase in mEPSCs amplitude did not arise from random superimposition of mEPSCs. If the increase in mEPSCs amplitude was caused by the spatial summation of mEPSCs at distinct sites, the rise time of large-amplitude events would be longer than that of small-amplitude events because of the weak synchronization of release (Sharma and Vijayaraghavan 2003; Shigetomi and Kato 2004), and there should be a positive correlation between the rise time and amplitude of mEPSCs after ischemia as well. However, as shown in Fig. 4B, there was no such a correlation. These results suggest that the increase in mEPSCs amplitude do not arise from random superimposition of events.
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Calcium plays important roles in neurotransmitter release. To investigate the calcium-dependence of enhanced presynaptic release after ischemia, we compared the frequency of mEPSCs in ischemic neurons with and without calcium. After 5-min baseline recording of mEPSCs in postischemic neurons, we switched the standard ACSF to Ca2+-free ACSF by removing the calcium from the external medium and adding EGTA (3 mM) in the presence of TTX (1 µM). Treatment of Ca2+-free ACSF for 5–10 min reversibly decreased the mEPSCs frequency by 44.5 ± 5.9% (P < 0.05, n = 8, Fig. 5). Interestingly, Ca2+-free medium only slightly reduced the mEPSCs frequency in control neurons (86.5 ± 8.1% of pre-Ca2+ removal, P = 0.15, n = 6). To reveal the involvement of voltage-gated calcium channels (VGCCs) in this process, we bath applied CdCl2 (100 µM) in postischemic slices to block the VGCCs and did not find any significant changes in the frequency of mEPSCs (96.6 ± 3.6% of pre-Cd2+ application, P = 0.48, n = 5, Fig. 5C). These results suggest that the increase of glutamate release after ischemia depends on external Ca2+, and VGCCs do not participate in this event.
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The preceding data indicate that the excitatory synaptic transmission onto spiny neurons is presynaptically potentiated after ischemia, resulting from Ca2+ influx into presynaptic terminals through the channels other than VGCCs. Previous studies have shown that extracellular ATP concentration is increased after brain ischemia through multiple mechanisms (Braun et al. 1998; Dutta et al. 2004), and the increased ATP can facilitate glutamate release in CNS through activation of presynaptic P2X receptors (Khakh et al. 2003; Shigetomi and Kato 2004). To examine whether activation of P2X receptors is responsible for the increased glutamate release after ischemia, we applied P2X receptor antagonist PPADS (50 µM) after 5-min stable recording of mEPSCs in either control or ischemic spiny neurons. Bath application of PPADS did not change the holding current and input resistance, suggesting its deficiency in postsynaptic function. As shown in Fig. 6, PPADS reliably reduced the mEPSCs frequency of ischemic neurons in a reversible manner (63.8 ± 4.7% of pre-PPADS application, P < 0.05, n = 9), but it had no detectable effects in control neurons (99.6 ± 3.2% of pre-PPADS application, P = 0.73, n = 7, Fig. 6B). Furthermore, we tested the effects of PPADS (50 µM) in Ca2+-free external solution. Removal of extracellular Ca2+ consistently reduced the mEPSCs frequency in ischemic neurons. However, PPADS failed to further change the mEPSCs frequency in postischemic spiny neurons in the absence of external Ca2+ (96.5 ± 3.8% of pre-PPADS application, P = 0.26, n = 7). Theses results suggest that activation of P2X type of ATP receptor resulted in enhanced glutamate release in ischemic neurons through a Ca2+-dependent manner.
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DISCUSSION |
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Accumulating evidence has shown that excitatory synaptic transmission in ischemia-vulnerable and ischemia-resistant neurons are differentially disturbed by ischemia. In the hippocampus, in vivo intracellular recording showed that evoked excitatory postsynaptic potentials (eEPSPs) in CA1 neurons are potentiated after ischemia (Gao and Xu 1996; Gao et al. 1998b; Tsubokawa et al. 1994). In contrast, the slope and amplitude of EPSPs are reduced or unchanged in CA3 neurons and dental granule cells subjected to the same insult (Gao et al. 1998a). In the striatal cholinergic interneurons, which are resistant to ischemia, excitatory synaptic transmission is depressed after ischemia because of activation of presynaptic adenosine A1 receptors and inhibited glutamate release (Pang et al. 2002). Studies using in vitro ischemia model demonstrated that ischemia is capable of inducing long-term synaptic potentiation (LTP) in spiny neurons but not in large aspiny interneurons (Calabresi et al. 2002). It is hypothesized that ischemia-induced LTP may, at least in part, account for the cell type-specific vulnerability after ischemia. The animal model of ischemia in the present study results in >80% cell death in the striatum 24 h after reperfusion (Pulsinelli et al. 1982; Ren et al. 1997). Therefore the potentiation of EPSCs at 9 h after ischemia as shown in the present study is probably associated with the cause rather than the result of ischemic cell death.
Both pre- and postsynaptic mechanisms affect the synaptic strength. Based on the studies in the hippocampus, the potentiation of excitatory neurotransmission results from the enhancement in postsynaptic responsiveness (Tsubokawa et al. 1995). For instance, exogenous application of glutamate receptor agonist evokes larger current and higher intracellular Ca2+ in postischemic brain slices than those in normal slices, suggesting that postsynaptic mechanism is involved (Mitani et al. 1998). More detailed studies have shown the upregulation of postsynaptic glutamate receptor function by protein kinase phosphorylation in CA1 pyramidal neurons after transient global ischemia. For example, ischemia induces tyrosine phosphorylation of NR2B subunit by Src-family kinases, and serine phosphorylation of NMDA NR2A subunit by cyclin-dependent kinase 5, leading to the enhancement of NMDAR currents (Takagi et al. 2003; Wang et al. 2003). Blockade of NMDAR phosphorylation, in turn, protects the neurons from ischemic injury (Wang et al. 2003). Increased phosphorylation of GluR1 subunits of AMPA receptors by calcium/calmodulin-dependent kinase II were also observed in the hippocampus (Takagi et al. 2003). In addition, ischemia is also capable of inducing the changes in subunits composition of AMPA receptor, particularly, downregulation of GluR2 submits that limits Ca2+ entry (Gorter et al. 1997; Pellegrini-Giampietro et al. 1992). Taken together, postsynaptic mechanism has been widely accepted as a major factor enhancing excitatory synaptic transmission in ischemic neuronal injury.
On the other hand, studies have shown that abnormal synaptic transmissions after ischemia are ascribed to the presynaptic intervention. In vitro studies have demonstrated that excitatory synaptic transmissions are presynaptically depressed during ischemia/hypoxia, conferred by unchanged response to exogenous application of glutamate (Calabresi et al. 1997b; Hershkowitz et al. 1993) and increase in paired-pulse facilitation (Calabresi et al. 1997a; Tanaka 2001). These effects can be antagonized by pretreatment with adenosine A1 receptors antagonist 8-CPT, suggesting that presynaptic A1 receptors are activated during ischemia and subsequently inhibit synaptic glutamate release (Calabresi et al. 1997a; Fowler et al. 2003; Tanaka 2001). Coincided with these observations, presynaptic depression of excitatory neurotransmission through adenosine A1 receptors has also been reported in large cholinergic interneurons in the striatum after ischemia in vivo (Pang et al. 2002). Presynaptic mechanism has also been implicated in the potentiation of excitatory synaptic transmission in CA1 pyramidal neurons 
48 h after reperfusion (Gao et al. 1998b). To further explore the role of presynaptic mechanism in ischemic injury, Abdel-Hamid and Tymianski (1997) enhanced neuronal Ca2+ buffering with BAPTA-AM to protect neurons against oxygen glucose deprivation. The protection by Ca2+ buffering appears to be presynaptic because BAPTA-AM is ineffective when endogenous transmitter release is bypassed by directly applying NMDA to the cultures (Abdel-Hamid and Tymianski 1997). Similarly, Monyer and co-workers (1992) found that tetanus toxin, presumably acting as a presynaptic blocker, attenuated neuronal injury after combined oxygen and glucose deprivation. In consistent with the preceding observations, the present provides direct evidence indicating that facilitated presynaptic release strengthens the excitatory neurotransmission in spiny neurons after reperfusion. These results offer new insights into the mechanisms of enhancement of excitatory neurotransmission in ischemia-vulnerable neurons after ischemia.
We have shown that blockade of P2X receptors almost completely reverses the increase of mEPSC frequency induced by ischemia, suggesting that the enhancement of P2X receptor function is responsible to the increased glutamate release from presynaptic terminals after ischemia. In the CNS, ATP is not only an energy source of the cell but also a potent extracellular signaling molecule eliciting a wide variety of physiological effects through activating ionotropic P2X and metabotropic P2Y receptors (Volonte et al. 2003). In physiological condition, ATP is present at a low concentration in extracellular space (Dubyak and el-Moatassim 1993). However, mounting evidence indicates that extracellular ATP concentration is increased in the pathophysiological conditions, such as ischemia, tissue injury and hypoxia (Braun et al. 1998; Lutz and Kabler 1997; Volonte et al. 2003). ATP may be released from swollen cells through ATP-conductive anion channels in response to ischemia or hypoxia (Dutta et al. 2004; Nieber et al. 1999), or co-released with glutamate from presynaptic terminals (Mori et al. 2001). Extracellular ATP is rapidly hydrolyzed to ADP and adenosine by a surface-located enzyme chain. Loss of ectoATPase activity in the pathological conditions provides additional evidence for the accumulation of extracellular ATP (Robson et al. 1997; Zimmermann 1994). P2X receptors are widely expressed in the CNS as indicated by immunocytochemistry and in situ hybridization (Deuchars et al. 2001; Kanjhan et al. 1999). Electron microscopic studies have revealed that the upregulation of P2X receptors after ischemia is located on the presynaptic appositions of synapses (Franke et al. 2004). Activation of presynaptic P2X receptors depolarizes presynaptic terminals and activates voltage-gated calcium channels to enhance neurotransmitter release (Khakh and Henderson 1998). Calcium could enter the terminal through presynaptic P2X receptors directly and facilitate transmitter release (Shigetomi and Kato 2004). Our data suggest that the enhancement of presynaptic release of glutamate in the striatum after ischemia is probably due to the calcium influx directly through the P2X receptor rather than the voltage-gated calcium channels.
The present study indicates that activation of ATP P2X receptor potentiates excitatory synaptic transmission in medium spiny neurons while activation of adenosine A1 receptor depresses excitatory synaptic transmission in large aspiny neurons after ischemia (Pang et al. 2002). Given the facts that hydrolyzing extracellular ATP can form adenosine (Zimmermann 1994), we speculate that such differential changes in excitatory input to medium spiny and large aspiny neurons presumably result from the distinct activities of ectoATPase after ischemia. It has been well established that high level of ATP is neurotoxic (Amadio et al. 2002; Neary et al. 1996; Ryu et al. 2002). Application of ATP receptor antagonists has the ability to prevent excitotoxicity (Cavaliere et al. 2001; Ralevic 2002). The detrimental mechanisms of ATP are not fully understood. Our results demonstrated a novel putative mechanism mediating neurotoxic effects of ATP.
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GRANTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: Z. C. Xu, Dept. of Anatomy and Cell Biology, Indiana University School of Medicine, 635 Barnhill Dr., MS 507, Indianapolis, IN 46202 (E-mail: zxu{at}anatomy.iupui.edu)
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