Ketamine-induced apoptosis in cultured rat cortical neurons

Ketamine-induced apoptosis in cultured rat cortical neurons

Toxicology and Applied Pharmacology 210 (2006) 100 – 107 Ketamine-induced apoptosis in cultured rat cortical neurons Ts...

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Toxicology and Applied Pharmacology 210 (2006) 100 – 107

Ketamine-induced apoptosis in cultured rat cortical neurons Tsuneo Takadera ⁎, Akira Ishida, Takao Ohyashiki Department of Clinical Chemistry, Faculty of Pharmaceutical Sciences, Hokuriku University, Kanazawa 920-1148, Japan Received 6 May 2005; revised 11 October 2005; accepted 14 October 2005

Abstract Recent data suggest that anesthetic drugs cause neurodegeneration during development. Ketamine is frequently used in infants and toddlers for elective surgeries. The purpose of this study is to determine whether glycogen synthase kinase-3 (GSK-3) is involved in ketamine-induced apoptosis. Ketamine increased apoptotic cell death with morphological changes which were characterized by cell shrinkage, nuclear condensation or fragmentation. In addition, insulin growth factor-1 completely blocked the ketamine-induced apoptotic cell death. Ketamine decreased Akt phosphorylation. GSK-3 is known as a downstream target of Akt. The selective inhibitors of GSK-3 prevented the ketamine-induced apoptosis. Moreover, caspase-3 activation was accompanied by the ketamine-induced cell death and inhibited by the GSK-3 inhibitors. These results suggest that activation of GSK-3 is involved in ketamine-induced apoptosis in rat cortical neurons. © 2005 Elsevier Inc. All rights reserved. Keywords: NMDA receptor; Glycogen synthase kinase-3; Ketamine; Apoptosis

Introduction Ketamine is frequently used in infants and toddlers for elective surgeries (Green and Johnson, 1990; Bergman, 1999). It is short acting and provides rapid dissociative anesthesia followed by rapid recovery. Ketamine is metabolized by cytochrome P450 (CYP3A4, CYP2B6 and CYP2CP). N-desmethylketamine, which is the main metabolite of ketamine, may contribute to the analgesic effects following ketamine administration (Shimoyama et al., 1999; Yanagihara et al., 2001; Hijazi and Boulieu, 2002). Ketamine may act as a noncompetitive blocker of Nmethyl-D-aspartate (NMDA) receptor ion channel (Anis et al., 1983; Harrison and Simmonds, 1985). The NMDA receptor, a subtype of the glutamate receptor, acts via the receptor-gated cation channel, which is permeable to Ca2+ and some monovalent cations. The NMDA receptor plays an important role as the main receptor, and in memory and learning (Collingridge and Bliss, 1995), and in differentia-

⁎ Corresponding author. Fax: +81 76 229 2781. E-mail address: [email protected] (T. Takadera). 0041-008X/$ - see front matter © 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.taap.2005.10.005

tion and development of the nervous system (Meldrum, 2000). Ikonomidou et al. (1999) have shown that ketamine induces cell death in postnatal rat brain. However, little is known about the intracellular mechanisms whereby ketamine induces neuronal cell death. Recent reports suggest that glycogen synthase kinase-3 (GSK-3) affects many fundamental cellular functions, including the cell cycle, gene transcription, cytoskeletal integrity and apoptosis (Cross et al., 1995; Grimes and Jope, 2001; Hetman et al., 2000). The phosphatidylinositol-3 kinase/Akt signaling pathway is one of the signaling systems implicated in the survival of neurons that leads to inhibition of GSK-3 by increasing Ser9 phophorylation (Cross et al., 1995; Pap and Cooper, 1998). Growth factors, such as IGF1, were reported to activate phosphatidylinositol-3 kinase, which leads to the phosphorylation and activation of Akt (Alessi et al., 1997). We reported recently that GSK-3 inhibitors blocked NMDA antagonist- and ethanol-induced apoptosis (Takadera et al., 2004; Takadera and Ohyashiki, 2004). Therefore, we examined whether anesthetic drugs such as ketamine also induce apoptotic cell death in cultured cells and whether GSK-3 is involved in the ketamineinduced cell death.

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Materials and methods Materials. Ketamine, alsteropaullone, anti-glial fibrillary acidic protein (GFAP), anti-actin and protease inhibitor cocktail were purchased from Sigma (St. Louis, USA). 5-Iodoindirubin-3′-monoxime, 3-(2-Chloro-3-indolylmethylene)-1,3-dihydroindol-2-one (Cdk1 inhibitor) and N4-(6-Aminopyrimidin-4yl)-sulfanilamide (Cdk2/5 inhibitor) were purchased from Merck (Tokyo, Japan). Hoechst 33258 (bis-benzimide) was purchased from Molecular Probes (Oregon, USA), Ac-Asp-Glu-Val-Asp-4-methyl-coumaryl-7-amide (AcDEVD-MCA) and 7-amino-4-methyl-coumarin (AMC) were acquired from Peptide Institute (Osaka, Japan). Rabbit anti-phospho-Akt (Ser473) affinitypurified polyclonal antibody was purchased from Biosource International (Camarillo, USA). Anti-neuronal nuclei (NeuN) monoclonal antibody was purchased from Chemicon International (Temecula, USA). Western blot kit BCIP/NBT system was purchased from KPL (Gaithersburg, USA). Bicinchoninic protein assay reagent was purchased from Pierce (Rockford, USA). Cycloheximide and other chemicals were purchased from Wako Pure Industries (Osaka, Japan). Cell culture. Cerebral cortical cells were obtained and cultured essentially as described by Dichter (1978) from fetal rats (Wistar) after 18–19 days of gestation. The dissociated cortical cells were cultured on poly-D-lysine-coated 35 mm dishes (Falcon 3001) (2 × 106 cells/dish) or coverglass in Dulbecco's modified Eagles medium (DMEM) containing 10% fetal calf serum. Cells were treated with cytosine-β-D-arabinofuranoside (10 μM) for 1 day (DIV5–6). The cells were maintained for 10 days prior to drug treatment. We examined the relative proportion of neurons to other cell types by staining the cells with antineuronal nuclei (NeuN) monoclonal antibody and anti-glial fibrillary protein (GFAP) visualized with an immunohistochemical staining kit, Vecstastain ABC kit (Vector). The relative proportion of neurons (NeuN+) to glia cells (GFAP+) was about 80%. Cell treatment and cell viability. The cells were washed twice with Trisbuffered salt solution containing (in mM): NaCl 120, KCl 5.4, CaCl2 1.8, MgCl2 0.8, Tris–HCl 25 and glucose 15 at pH 6.5 (washing buffer) and then replaced


with 2 ml of DMEM. To protect the cells from glutamate toxicity induced by serum containing glutamate, the serum was removed. Therefore, control cells also underwent up to 25% apoptosis depending on the experimental conditions by serum withdrawal. We also used washing buffer at pH 6.5 to avoid glutamate neurotoxicity which depends on extracellular pH. Cell treatment with various reagents was carried out for 48 h at 37 °C. Morphological cell changes were observed by phase-contrast microscopy during the treatment. Cell viability was checked by staining the cells with trypan blue dye. We used the washing buffer at pH 6.5 to block the glutamate neurotoxicity during the treatment. Quantitation of apoptosis by nuclear morphological changes. Apoptotic cell death was determined by staining the cells with Hoechst dye H33258. The cells were fixed with a 10% formalin neutral phosphate buffer solution (pH 7.4) for 5 min at room temperature. After washing the cells with distilled water, they were stained with 8 μg/ml of H33258 for 5 min. The nuclear morphology was observed under a fluorescent microscope (Olympus IX70 model). Apoptosis was quantitated by scoring the percentage of cells with apoptotic nuclear morphology at the single cell level. Condensed or fragmented nuclei were scored as apoptotic. A total of 5–7 randomly selected fields were captured using Polaroid PDMC II software. At least 200 cells were counted per condition, and each experiment was repeated in at least 3 different cultures. Analysis of DNA fragmentation. Low molecular weight DNA was isolated from 6 × 106 cells. The samples were treated with RNase (0.4 mg/ml) and proteinase K (0.625 mg/ml) for 1 h at 37 °C. Agarose (1.2%) gel electrophoresis of DNA was performed in a 40 mM Tris–HCl buffer (pH 8.1) containing 2 mM of EDTA. After electrophoresis, the gels were stained with ethidium bromide (0.5 μg/ml) for 15 min at room temperature. Western blotting. Primary cultured cells were scraped off the dish and collected by centrifugation (400 × g for 5 min) followed by homogenization in ice-cold buffer (50 mM Tris–HCl buffer containing 50 mM NaCl, 10 mM EGTA, 5 mM EDTA, 2 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM benzamide and 10% protease inhibitor cocktail, pH 7.4). The cell suspension was placed on ice for 30 min and centrifuged at 18,000 × g for 30 min. The supernatant was diluted with an equal volume of

Fig. 1. Ketamine-induced apoptosis. (A) Phase-contrast (a and d), trypan blue staining (b and e) and H33258 fluorescence (c, f) microscopy of the cortical cells after exposure to ketamine. Cells were incubated in the absence (a–c) or presence (d–f) of 100 μM ketamine for 48 h at 37 °C as described in Materials and methods. Arrowheads indicate healthy neurons. Arrows indicate neurons with apoptotic morphology. Scale bars = 10 μm. (B) DNA fragmentation after exposure to ketamine. Cortical cells were incubated in the absence (Cont) or presence (Ket) of ketamine (100 μM) for 24 h at 37 °C. Low-molecular weight DNA was isolated from the cells, and DNA laddering was detected by agarose electrophoresis as described in Materials and methods. Size marker of 100 bp DNA ladder was used.


T. Takadera et al. / Toxicology and Applied Pharmacology 210 (2006) 100–107 sample buffer containing 62.5 mM of Tris–HCl (pH 6.8), 2% sodium dodecyl sulfate (SDS), 10% glycerol, 50 mM of dithiothreitol and 0.1% bromphenol blue, heated at 95 °C for 5 min and stored at −20 °C. Protein concentration was determined by the bicinchoninic acid assay (Smith et al., 1985). Each sample (20 μg/lane) was loaded and separated using 7.5% SDS-polyacrylamide gel electrophoresis. Proteins were transferred on a nitrocellulose membrane and blocked with Tris-buffered saline containing 5% skim milk for 1 h at room temperature and then incubated with an anti-phospho-Akt (Ser473) polyclonal antibody in Tris-buffered saline overnight at 4 °C. After washing for 5 min with three changes of Tris-buffered saline, the membrane was incubated with a phosphatase-conjugated goat anti-rabbit antibody for 1 h at room temperature in Tris-buffered saline. After washing for 5 min with three changes of Trisbuffered saline, immunoreactive bands were visualized with a Western blot detection kit BCIP/NBT system. Equal amounts of protein extracts were also analyzed by Western blot analysis with anti-actin antibody. Caspase-3 activity. The caspase-3 activity was measured as described previously (Takadera et al., 1999). Briefly, the cells were washed with phosphate-buffered saline and suspended in 50 mM of Tris–HCl buffer (pH 7.4) containing 1 mM of EDTA and 10 mM of EGTA. The cells were treated with 10 μM of digitonin for 10 min. The lysates were obtained by centrifugation at 10,000 × g for 5 min, and the cleared lysates containing 50–100 μg protein were incubated with 50 μM of enzyme substrate Ac-DEVD-MCA for 1 h at 37 °C. The reaction was terminated by addition of monoiodoacetic acid (5 mM). AMC levels were measured by using a spectrofluorometer (Hitachi 850, Japan) with excitation at 380 nm and emission at 460 nm, and the activity was expressed as nmol of AMC released/min/mg protein. Statistics. Treatment effects were statistically analyzed by one-way ANOVA followed by post hoc Scheffe's comparisons.

Results Fig. 2. Time dependency of ketamine-induced neuronal apoptosis. (A) Decrease in the number of neurons after ketamine treatment. The cells were incubated in the presence (right columns) or absence (left columns) of ketamine (100 μM) for 0–48 h at 37 °C. Neuronal cells were detected as described in Materials and methods. Data are shown as the mean ± SEM. n = 4 per group. *P b 0.05 (vs. untreated cells at 0 time). (B) Ketamine-induced apoptosis. The cells were incubated in the presence (right columns) or absence (left columns) of ketamine (100 μM) for 48 h at 37 °C. Apoptosis was detected as described in Materials and methods. Data are shown as the mean ± SEM. n = 4 per group. *P b 0.05 (vs. untreated cells at 12 h).

Ketamine-induced apoptosis We examined the effect of ketamine, one of NMDA receptor antagonists, on rat cultured cortical cells. After treatment of cortical neurons with 100 μM of ketamine for 48 h, cells were stained with trypan blue, indicating loss of cell viability, and showed apoptotic morphology, including shrunken cell bodies, fragmented processes and condensed or fragmented nuclei (Fig. 1A). In addition, ketamine

Fig. 3. Blockade of ketamine- and MK801-induced apoptosis by NMDA. The cells were treated with ketamine (A, 100 μM) or MK801 (B, 100 nM) in the presence or absence of NMDA (25 μM) for 24 h at 37 °C. Data are shown as the mean ± SEM. n = 6 per group. *P b 0.05 (vs. ketamine- or MK801-only-treated cells).

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IGF-1 prevented ketamine-induced apoptosis IGF-1 has been reported to play a role in differentiation and survival in CNS (Stewart and Rotwein, 1996) and to activate the phosphatidylinositol-3 kinase/Akt (also known as protein kinase B) signaling pathway (Alessi et al., 1997). Whether IGF-1 protected the cells from ketamine-induced apoptosis was checked. As shown in Fig. 4, IGF-1 (5 and 50 ng/ml) attenuated ketamine-induced apoptosis in a dose-dependent manner. Ketamine decreased Akt phosphorylation levels Fig. 4. Protective effect of IGF-1 on ketamine-induced apoptosis. The cells were incubated with 100 μM of ketamine in the presence of IGF-1 (5 and 50 ng/ml) for 48 h at 37 °C. Data are shown as the mean ± SEM. n = 4 per group. *P b 0.01 (vs. ketamine-only-treated cells).

treatment increased nucleosomal size DNA fragmentation (Fig. 1B). Treatment of cells with 100 μM of ketamine resulted in a time-dependent decrease in the number of neurons and increased apoptotic cell death (Fig. 2). Up to 45% of neurons showed apoptotic cell death by treatment of cells with 100 μM of ketamine for 48 h. To confirm that the effect of ketamine on cell viability was related to the NMDA receptor, we examined the ability of NMDA, an agonist of NMDA receptor, to block ketamine-induced cell death. As shown in Fig. 3A, NMDA blocked ketamine-induced apoptosis, the same as NMDA antagonist MK801-induced apoptosis (Fig. 3B).

Akt is phosphorylated at two sites that are associated with activation of enzyme activity; Thr308 in the catalytic domain and Ser473 in the cytoplasmic domain. Phosphorylation of both sites is critically dependent upon PI3-K activity (Alessi et al., 1997). Akt phosphorylation at Ser473 was determined by immunoblot analysis utilizing a phospho-Akt-(Ser473)-specific antibody. Incubation of cortical cultures with 50 μM of NMDA for 1 h enhanced phospho-Akt levels (Fig. 5A). The NMDA receptor antagonist ketamine blocked the NMDA-enhanced phospho-Akt levels. Ketamine also reduced phospho-Akt levels in the absence of NMDA in a dose-dependent manner (Fig. 5B). Inhibitors of GSK-3 prevented ketamine-induced apoptosis Glycogen synthase kinase-3 is a principal physiological substrate of Akt, and the activity of GSK-3 is inhibited

Fig. 5. Ketamine decreased Akt phosphorylation levels. (A) Ketamine inhibited NMDA-stimulated Akt phosphorylation levels. The cells were incubated with or without 50 μM NMDA in the presence or absence of 100 μM ketamine for 1 h at 37 °C. Equal amounts of protein extracts were analyzed by Western blot analysis with phospho-Akt (Ser473). The optical densities for pAkt bands were compared to that for actin. Data are expressed as a percentage of optical density value for control. Data are shown as the mean ± SEM. n = 3 per group. *P b 0.01 (vs. NMDA-only-treated cells). (B) Ketamine decreased Akt phosphorylation levels of cells. The cells were incubated in the presence or absence of 1–100 μM ketamine for 1 h at 37 °C. Data are shown as the mean ± SEM. n = 5 per group. *P b 0.05 (vs. untreated cells).


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by Akt-mediated phosphorylation in response to trophic stimulation. To investigate directly the role of the activity of endogenous GSK-3 in the cell death in response to ketamine treatment, we assayed the effects of selective inhibitors, alsteropaullone and 5-iodoindirubin-3′-monoxime, on ketamine-induced apoptosis. The GSK-3 inhibitors showed a dose-dependent protective effect against the ketamine-induced apoptosis (Figs. 6A and B). Alsteropaullone and 5-iodoindirubin-3′-monoxime are also known to inhibit cyclin-dependent protein kinase 1 or 5 (Leclerc et al., 2001; Leost et al., 2000). However, cyclin-dependent protein kinase 1 and 2/5 selective inhibitors such as 3-(2Chloro-3-indolylmethylene)-1,3-dihydroindol-2-one (Cdk1 inhibitor) and N 4 -(6-Aminopyrimidin-4-yl)-sulfanilamide (Cdk2/5 inhibitor) did not prevent the ketamine-induced apoptosis (Fig. 6C), suggesting that alsteropaullone and 5iodoindirubin-3′-monoxime prevented apoptosis by inhibiting GSK-3 activity.

Inhibitors of GSK-3 prevented ketamine-induced caspase-3 activation The key apoptosis effectors in mammals are a family of aspartate-specific proteinases called caspases (Nicholson et al., 1995; Alnemri et al., 1996). Caspase-3 was shown to play a critical role during normal brain development (Kuida et al., 1996). Therefore, we measured caspase-3 activity after treatment of the cells with 100 μM of ketamine using AcDEVD-MCA as a peptide substrate. Ketamine increased the caspase-3 activity. In addition, GSK-3 inhibitors such as alsteropaullone and 5-iodoindirubin-3′-monoxime reduced the ketamine-increased caspase-3 activity (Fig. 7). Discussion We showed in this report that GSK-3 inhibitors protected cortical neurons from the ketamine-induced apoptosis,

Fig. 6. Protective effect of GSK-3 inhibitors on ketamine-induced apoptosis. (A) Effect of alsteropaullone on ketamine-induced apoptosis. The cells were incubated with 100 μM of ketamine in the presence or absence of alsteropaullone (0.1 and 1 μM) for 48 h at 37 °C. Each value represents the mean ± SEM. n = 4 per group. *P b 0.05 (vs. ketamine only treated cells). Ket, ketamine; Als, alsteropaullone. (B) Effect of 5-iodoindirubin-3′-monoxime on ketamine-induced apoptosis. The cells were incubated with 100 μM of ketamine in the presence or absence of 5-iodoindirubin-3′-monoxime (1 and 5 μM) for 48 h at 37 °C. Each value represents the mean ± SEM. n = 4 per group. *P b 0.05 (vs. ketamine-only-treated cells). Iodo, 5-iodoindirubin-3′-monoxime. (C) Effects of CDK1 and CDK-2/5 inhibitors on ketamine-induced apoptosis. The cells were incubated with 100 μM of ketamine in the presence or absence of 3-(2-Chloro-3-indolylmethylene)-1,3-dihydroindol-2one (Cdk1 inhibitor) and N4-(6-Aminopyrimidin-4-yl)-sulfanilamide (Cdk2/5 inhibitor) for 48 h at 37 °C. Each value represents the mean ± SEM. n = 4 per group. *P b 0.05 (vs. ketamine only treated cells). Cdk1, cdk1 inhibitor; Cdk2/5, cdk2/5 inhibitor.

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Fig. 7. Effects of GSK-3 inhibitors on ketamine-stimulated caspase-3 activation. The cells were incubated with 100 μM ketamine in the presence of alsteropaullone (0.1 and 1 μM) or 5-iodoindirubin-3′-monoxime (1 and 5 μM) for 4 h at 37 °C. The caspase-3 activity of the lysates was measured as described in Materials and methods. Each value represents the mean ± SEM. n = 3 per group. *P b 0.01 (vs. ketamine-only-treated cells).

suggesting that GSK-3 activity is critical for neuronal cell death induced by blocking the trophic effect of NMDA receptor with ketamine. Glutamate plays an important role in neuronal survival in the developing brain largely through facilitating the entry of Ca2+ (Balazs et al., 1988; Yan et al., 1994). Activation of NMDA receptors induces Akt activation through an NMDA receptor-phosphatidylinositol-3 kinase pathway (Zhang et al., 1998; Sutton and Chandler, 2002) or calcium/calmodulindependent kinase (Yano et al., 1998). Ketamine inhibited the NMDA-induced Akt phosphorylation, suggesting that ketamine induced apoptosis by disturbing the phosphatidylinositol-3 kinase or calcium/calmodulin-dependent kinase/Akt signaling pathway. Survival of neuronal cells appears critically dependent upon an optimal intracellular ‘calcium set point’ (Johnson and Deckwerth, 1993). Excitotoxic neuronal necrosis is triggered by overstimulation of NMDA receptors. An inadequate Ca2+ level may overcome the trophic effect of calcium and activate calcium-dependent enzymes including calpain which induces proteolysis of neuronal cytoskeleton (Yokota et al., 2003; Bano et al., 2005). On the other hand, NMDA antagonists induce apoptosis to cortical neurons at times when synapses are forming, suggesting that spontaneous NMDA receptor activity supports survival of neurons in development. The spontaneous NMDA receptor activity may maintain basal p-AKT levels, and ketamine induces apoptosis by blocking the NMDA receptor activity and decreasing p-AKT levels. Glycogen synthase kinase-3 activity is known to be suppressed when it becomes phosphorylated on serine 9 by activation of Akt. Therefore, spontaneous NMDA activity may contribute to the survival of cortical cells by activating Akt followed by inhibiting GSK-3. However, the downstream substrates of GSK-3beta that ultimately induce neuronal death are not clear. Recently, Linseman et al. (2004) reported that GSK-3 phosphorylates Bax, a pro-apoptotic Bcl-2 family member that stimulates the intrinsic (mitochondrial) death pathway by eliciting cytochrome c release from mitochondria, and promotes its mitochondrial localization.

The apoptosis induced by ketamine was accompanied by activation of caspase-3. Similarly, we have previously reported that NMDA antagonists such as MK801 induce apoptotic cell death accompanied by caspase-3 activation in rat cortical cells (Takadera et al., 1999). We do not rule out the possibility that caspase-7 also contributes to the ketamine-induced apoptosis because this caspase also cleaves the peptide substrate (Thornberry et al., 1997). In the hippocampus and cerebral cortex, the expression of NR1 and NR2B subunit is predominant at times when synapses are forming, whereas the expression of NR2A subunit is low and then increases to plateau levels later in development (Li et al., 1998; Watanabe et al., 1992). The rat cultured cortical neurons are considered developing neurons because NR2A and NR2B subunit proteins have developmental profiles in cultured cortical neurons similar to those seen in vivo. NR1 and NR2B subunits display high levels of expression within the first week. In contrast, the NR2A subunit is barely detectable at 7 days in vitro (DIV) and then gradually increased to mature levels at DIV21 (Li et al., 1998). The window of vulnerability to NMDA antagonist-induced apoptosis coincides with the period of synaptogenesis, also known as the brain growth spurn period. This period in rats is largely confined to the postnatal period; it begins 1 day before birth and terminates at approximately 14 days after birth, whereas in the human it spans the last 3 months of pregnancy and extends into the first several postnatal years (Dobbing and Sands, 1979). Blockade of NMDA receptors triggers apoptotic neurodegeneration in developing rat brain, suggesting that glutamate acts as a physiological regulator of programmed cell death during normal development. On the other hand, application of a single dose of NMDA or other excitotoxins produces a massive necrotic neuronal death, suggesting that glutamate neurotoxicity is involved in the pathogenesis such as ischemia. Blood levels of ketamine are 5 μM at anesthetic levels and 0.5 mM at toxic levels. Ketamine does not induce neuronal death when blood levels are close to an anesthetic level in human. However, intake of a repeated dose of ketamine might be quite harmful in fetuses or infants (Farber and Olney, 2003; Scallet et al., 2004).


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We showed in this report for the first time that GSK-3 inhibitors protected cortical neurons from the ketamine-induced apoptosis induced possibly by blocking the trophic effect of NMDA receptor. Therefore, GSK-3 inhibitors may deserve to be evaluated as therapeutic agents in anesthetic drug-induced neurodegenerative disorders.

Acknowledgment This study was supported in part by a special in-house research grant from Hokuriku University.

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