Increased susceptibility of glutathione peroxidase-1 transgenic mice to kainic acid-related seizure activity and hippocampal neuronal cell death

Experimental Neurology 192 (2005) 203 – 214 www.elsevier.com/locate/yexnr

Increased susceptibility of glutathione peroxidase-1 transgenic mice to kainic acid-related seizure activity and hippocampal neuronal cell death R. Boonplueanga,b, G. Akopianc, F.F. Stevensona, J.F. Kuhlenkampd, S.C. Lud, J.P. Walshc, J.K. Andersena,c,* a Buck Institute for Age Research, 8001 Redwood Boulevard, Novato, CA 94945, USA Program in Molecular Biology, Department of Biological Sciences, University of Southern California, Los Angeles, CA 90089, USA c Andrus Gerontology School, University of Southern California, Los Angeles, CA 90089, USA d Division of Gastroenterology and Liver Diseases, USC Research Center for Liver Diseases, Keck School of Medicine, University of Southern California, Los Angeles, CA 90033, USA b

Received 18 June 2004; revised 15 December 2004; accepted 16 December 2004

Abstract Glutathione peroxidase (GSHPx) has been demonstrated in several in vivo studies to reduce both the risk and severity of oxidativelyinduced tissue damage. The seizure-inducing neurotoxin kainic acid (KA) has been suggested to elicit its toxic effects in part via generation of oxidative stress. In this study, we report that expression of elevated levels of murine GSHPx-1 in transgenic mice surprisingly results in increased rather than decreased KA susceptibility including increased seizure activity and neuronal hippocampal damage. Isolated transgenic primary hippocampal culture neurons also display increased susceptibility to KA treatment compared with those from wildtype animals. This could be due to alterations in the redox state of the glutathione system resulting in elevated glutathione disulfide (GSSG) levels which, in turn, may directly activate NMDA receptors or enhanced response of the NMDA receptor. D 2004 Elsevier Inc. All rights reserved. Keywords: Kainic acid; Glutathione; Glutathione disulfide; Glutathione peroxidase; Neuronal cell death; Oxidative stress; Seizure; Epilepsy; NMDA receptors; Excitotoxicity

Introduction Several selenocysteine-containing enzymes have been identified and categorized as part of the glutathione peroxidase (GSHPx) enzyme family. These include cytosolic GSHPx (GSHPx-1) (Flohe et al., 1973; Rotruck et al., 1973) gastrointestinal GSHPx (GSHPx-GI or GSHPx-2) (Chu et al., 1993), plasma GSHPx (GSHPx-P or GSHPx-3) (Takahashi et al., 1987), phospholipid hydroperoxide GSHPx (PHGSHPx or GSHPx-4) (Ursini et al., 1985), and sperm nuclei GSHPx (snGSHPx) (Pfeifer et al., 2001). The GSHPx family enzymes are found ubiquitously in mammals and are best known for their role in eliminating * Corresponding author. Buck Institute for Age Research, 8001 Redwood Boulevard, Novato, CA 94945, USA. Fax: +1 415 209 2231. E-mail address: [email protected] (J.K. Andersen). 0014-4886/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.expneurol.2004.12.017

reactive oxygen species (ROS) such as hydrogen peroxide and peroxynitrite. A number of studies have demonstrated their capacity for reducing the risk and severity of oxidatively-induced tissue damage. For example, transgenic mice expressing the human form of the GSHPx-1 enzyme displayed significantly less focal cerebral ischemia and reperfusion damage as a result of middle cerebral artery occlusion compared to wildtype littermates (WeisbrotLefkowitz et al., 1998). An elevation of GSHPx-P or GSHPx-1 levels attenuate lipopolysaccharide-induced endotoxemia in transgenic mice (Mirochnitchenko et al., 2000). Moderately elevated levels of GSHPx-1 levels (10% above wildtype) may improve the capacity of hippocampal cultures to recover after in vitro hypoxia (Furling et al., 2000). Overexpression of the human GSHPx-1 levels in transgenic mice protects the heart against oxidative stress from post-myocardial infarction (MI) left ventricle structural

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remodeling and functional failure (Shiomi et al., 2004). Recently, GSHPx-1 has also been shown to be a tumor suppressor; adenovirus-mediated overexpression of human GSHPx-1 in pancreatic MIA PaCa-2 tumor cells was shown to slow down in vitro growth compared to parental cells and of MIA PaCa-2 induced tumors in nude mice (Liu et al., 2004). Although in many studies GSHPx appears neuroprotective, other studies suggest its increased expression may increase susceptibility to various agents. For example, HIV1 infected human sup-T1 T cells overexpressing GSHPx-1 displayed more rapid HIV-1 replication and less viability than uninfected cells, possibly due to the ability of GSHPx to suppress an apoptotic response to viral infection (Sandstrom et al., 1998). Hepatocytes isolated from GSHPx-1 knockout mice were more resistant to peroxynitrite administration than those isolated from wildtype mice suggesting that GSHPx-1 activity actually promotes peroxynitrite-induced apoptosis by some unknown mechanism (Fu et al., 2001). Human GSHPx-1 transgenic mice were more susceptible to acetaminophen toxicity and to hyperthermia than wildtype possibly due to depletion of the substrate glutathione (GSH) via increased GSHPx activity (Mirochnitchenko et al., 1995, 1999). Increased GSHPx-1 activity in lens epithelial cells prepared from GSHPx-1 transgenic was demonstrated to be no more efficient in H2O2 degradation than wildtype probably due to an imbalance in the GSHPx-1/glutathione reductase (GR) activity ratio and limitations on GSH production (Spector et al., 1996). Increased expression of human GSHPx-1 in transgenic mice resulting in lower levels of H2O2 and lipid peroxides, both effective inducers of heat shock protein 70 (HSP70), leads to lowered HSP70 activation and an increased sensitivity to heat-induced hyperthermia (Mirochnitchenko et al., 1995). A plethora of data suggests that oxidative stress is likely to play a significant role in seizure-induced cellular damage in the hippocampus (Carrasco et al., 2000; Coyle and Puttfarcken, 1993; Erakovic et al., 2000; Liang et al., 2000; Patel et al., 2001). To test this hypothesis and to explore which particular species of ROS might be involved, we assessed the effects of systemic administration of the seizure-inducing agent kainic acid (KA) in mice with elevated GSHPx-1 expression to see if this would be protective against seizures and subsequent hippocampal cell death. Surprisingly, increased hippocampal levels of GSHPx-1 resulted in increased rather than decreased brain damage following KA administration.

Materials and methods Materials All chemicals were purchased from Sigma-Aldrich, Saint Louis, MO, unless otherwise stated.

Transgenic animals GSHPx-1 transgenic mice were a gift from Dr. Y.S. Ho, Wayne State University, Detroit, MI). Briefly, a 5.3-kb SacI fragment containing 1.4-kb of the murine cellular GSHPx (GSHPx-1) gene isolated from a bacteriophage FIX II genomic library was microinjected into B6C3F1 fertilized eggs to create GSHPx transgenic animals (Yoshida et al., 1996). Polymerase chain reaction was used for genotyping the animals using primers against the flanking regions of adjacent transgenes. Animals used in the study were bred inhouse and were housed according to standard animal care protocols, fed ad libitum, kept on a 12-h light/dark cycle, and maintained in a pathogen-free environment in the Buck Institute Vivarium, accredited by AALAC. All animal experiments were approved by local committee review and conducted according to policies on the use of animals of The Society for Neuroscience. Glutathione peroxidase (GSHPx) activity assay A GSHPx assay kit (BIOXYTECH GPx-340, Oxis International, Portland, OR) was used to determine hippocampal GSHPx activity levels. The assay is based on a change in NADPH absorbance at 340 nm. Briefly, animals were perfused with 0.9% NaCl containing 0.16 mg/ml heparin. Hippocampi were removed and homogenized in cold 50 mM Tris–HCl, pH 7.5, 5 mM EDTA, 1 mM 2-mercaptoethanol and supernatant collected after centrifugation at 10,000  g for 10 min at 48C. Protein concentration was first estimated by using the Bradford reagent (Bio-Rad, Hercules, California). 15 Al of hippocampal lysate was then added to 150 Al of 0.05 M Tris– HCL, pH 7.6, 5 mM EDTA containing 0.24 Amol glutathione, 0.12 U glutathione reductase (GR) and 0.048 Amol h-NADPH. 75 AL of 0.007% tert-butyl hydroperoxide was added to the sample mixture. Absorbance was recorded at 340 nm for 3 min. GSHPx activity levels were determined against a standard curve. One GSSG activity unit is defined as the amount of enzyme catalyzing the oxidation of one 1 AM of NADPH per minute. Note that although this assay measures all GSHPx activities and is not specific to GSHPx1, since GSHPx-1 is the most abundant and the one with increased expression in this transgenic line, GSHPx activity is expressed as GSHPx-1 activity (Spector et al., 1996). In vivo treatment with KA, seizure assessment, and histological staining for CA3 hippocampal cell injury A dosage of 35 mg/kg body weight KA (Ocean Produce International, Nova Scotia, Canada) in PBS was injected intraperitoneally into 8–12 weeks old mice. For a period of 2 h after KA administration, seizure activities of each animal were observed and rated according to an arbitrary scale (Viswanath et al., 2000): 1—very light seizures of forepaws, 2—repeated mild seizures, rearing and loss of postural

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control, 3—whole body seizures for a short time, and 4— seizures become severe and prolonged. 24 h after KA administration, brains were collected, fresh frozen and kept at 808C. 16-Am coronal sections through the hippocampi were obtained from the fresh frozen brains using an RMC cryostat. Sections were stained by Terminal Deoxynucleotide Transferase-mediated dUTP Nick-End Labeling (TUNEL) (TdT-Fragel DNA fragmentation detection kit, Oncogene Research Products, Cambridge, MA), cresyl violet (CV) and haematoxylin and eosin (H&E) staining.

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were then extracted with 3 volumes of cold 5% metaphosphoric acid (MPA). MPA extracts were collected after a centrifugation at 1000  g for 10 min at 48C. 200 Al of 1.3 mM DTNB in 100 mM NaPO4 containing 5 mM EDTA, pH 7.5 with 5% ethanol and 3 U of GR in 100 mM NaPO4 containing 5 mM EDTA, pH 7.5 were added to 200 Al of the extract. After a 5-min incubation at room temperature and addition of 0.1 mM NADPH, the absorbance was recorded at 412 nm for 3 min. GSH and GSSG levels were determined against a standard curve. GSH/GSSG ratios were calculated from the formula (GSH-2GSSG)/GSSG.

In vitro treatment with KA and cell viability assay Primary hippocampal neuronal cultures were isolated from E18 animals. Briefly, hippocampal tissues were dissected in Ca2+–Mg2+-free Hank’s salt solution (Life Technologies, Carlsbad, California) and digested with 0.25% EDTA-trypsin. Cells were dissociated in 10% fetal bovine serum (FBS)-DMEM medium (Life Technologies, Carlsbad, California) and plated onto poly-l-lysine coated 48-well plates. One hour after plating, the medium was changed to Neurobasal medium (Life Technologies, Carlsbad, California) with B-27 supplement Minus AO (Life Technologies, Carlsbad, California) and 2 mM glutamine (Life Technologies, Carlsbad, California) to remove nonadherent cells. Cells were maintained in a humidified atmosphere of 95% air and 5% CO2 at 378C. Medium was half-changed every 4 days. Cultures were treated with 10 AM cytosine h-d-arabinofuranoside on days 3 and 6 to remove non-neuronal cells. At days 11–12 in vitro (DIV), the cultures were treated with various concentrations of KA in neurobasal medium with B-27 supplement minus AO and glutamine: 0, 10, 25, 50, 100, and 250 AM for 24 h. Cell viability assay was performed on KA-treated cultures using CellTiter-Blue cell viability assay kit (Promega, Madison, WI) by incubating cultures with 20 Al of CellTiter-Blue reagent per 100 ml of medium at 378C for 4 h. At the end of the incubation, the fluorescence is measured in a fluorescence microplate spectrofluorometer (Gemini, Molecular Devices, Sunnyvale, CA) using an excitation wavelength of 560 nm and an emission wavelength of 590 nm. GSH/GSSG ratios A GSH/GSSG assay kit (BIOXYTECH GSH/GSSG412, Oxis International, Portland, OR) was used to determine the GSH/GSSG ratio. The assay is based on the coloric development of 5,5V-dithiobis-2-nitrobenzoic acid (DTNB) reacting with GSH at 412 nm. Briefly, brains were homogenized in 1 ml of iced-cold 0.05 M Tris–HCl, pH 7.6 containing 5 mM EDTA and 1 mM mercaptoethanol with or without 33 mM 1-methyl-2-vinylpyridine (M2VP), a GSH scavenger. Supernatant was collected after a centrifugation at 10,000  g for 10 min at 48C. Protein concentrations of the lysates were estimated using the Bradford reagent (Bio-Rad, Hercules, California). Samples

High-performance liquid chromatography (HPLC) analysis of GSSG For hippocampal GSSG tissue measurements, animals were perfused with 0.9% NaCl containing 0.16 mg/ml heparin. Hippocampi were removed, weighed, frozen in liquid nitrogen and homogenized in 1 ml of 10% perchloric acid (PCA) with 1 mM bathophenanthrelinedisulfonic acid (BPDS) with or without an internal control, 1 AM GSSG. The homogenate was then sonicated at power 3 for 20 s (Sonic Dismembrator model 550, Fisher Scientific, Pittsburgh, PA) frozen and thawed. 0.5 ml of supernatant obtained after a 3-min centrifugation at 15,000  g was added to a tube containing 50 Al of 100 mM iodoacetic acid in 0.2 mM m-cresol purple solution. 0.5 ml of 2 M KOH– 2.4 M KHCO3 was then added to the tube to raise the pH of the solution up to pH 8–9. After a 10-min incubation in the dark at room temperature, 1 ml of 1% fluorodinitrobenzene was added to the solution. The solution was then mixed and stored at 48C overnight to allow derivatization to completely occur. For extracellular GSSG measurements, at 11–13 DIV primary hippocampal culture neurons isolated form E18 animals were treated with 0 or 50 AM KA for 24 h, the media collected then precipitated and derivatized using the same method as for hippocampi preparations. Hippocampal and extracellular GSSG levels were determined by the HPLC method of Fariss and Reed (1987) (LC-10ATVP pump, SCL-10AVP system control, Shimadzu) with a SPD10AVP UV detector and a SIL-10ADVP autosampler (Shimadzu) using a 3-amino propyl 5Am column (4.6  200 mm, Cel Associates, Inc., Pearland, TX). GSSG was identified by measuring absorbance at 365 nm at a sensitivity scale of 0.01. The amount of GSSG in each sample was calculated from a standard curve of GSSG prepared at the same time as the samples. The identity of GSSG peak was also confirmed by spiking the sample with GSSG standard. GR, Glucose-6-phosphate dehydrogenase (G-6-PD) and 6-phosphogluconate dehydrogenase (6-GPD) enzyme assays Brains were homogenized in cold 1 PBS containing 1 mM EDTA. After centrifugation at 8500  g for 10 min at

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48C, supernatants were collected for protein concentration estimation (Bio-Rad, Hercules, California) and enzyme activity assays. A GR assay kit (Bioxytech GR-340, Oxis International, Portland, OR) was used to determine brain GR activity levels according to manufacturer’s instructions. The assay is based on the oxidation of NADPH to NADP+ catalyzed by GR. GR activity levels were determined by a change in NADPH absorbance at 340 nm. One GR activity unit is defined as the amount of GR catalyzing the reduction of 1 AM of GSSG per minute. A G-6-PD/6-GPD assay kit (Bioxytech G6PD/6PGD-340, Oxis International, Portland, OR) was used to determine brain G-6PD and 6-GPD levels, according to manufacturer’s instructions. The assay is based on the increase of NADPH absorbance at 340 nm catalyzed by G-6-PD and 6-GPD. One unit of either enzyme activity is defined as the amount of enzyme producing 1 AM of NADPH per minute. Western blot analysis of NMDA receptor subunits Hippocampi (n = 3 for each group, HM, HT, WT) were dissected 24 h after 0.9% NaCl administration and homogenized in RIPA buffer (1  PBS, 1% Igepal CA630, 0.5% sodium deoxycholate, 0.1% SDS containing 10 mg/ml PMSF and 30 Ag/ml aprotinin). 100 Ag protein per lane of samples were electrophoresed on 10% SDS/PAGE gels. Samples were then transferred to poly-vinylidene difluoride (PVDF) membranes. After incubation in 5% non-fat milk in TBS for 1 h at room temperature, membranes were incubated overnight at 48C with rabbit polyclonal antibodies against either glutamate receptor subunits NMDAR1 (NR1) (1:1000), NMDAR2A (NR2A) (1:200) or NMDAR2B (NR2B) (1:200). A horseradish peroxidase-conjugated anti-rabbit antibody (1:2000– 1:3000, Amersham Pharmacia Biotech, Piscataway, NJ) was used as secondary antibody. A chemiluminescence substrate system (ECL, Amersham Pharmacia Biotech, Piscataway, NJ) was used to detect antibody binding. h-actin antibody (1:5000) binding was used to normalize optical densities of protein bands. The relative optical densities of protein bands were quantitated using a ChemiImager 5500 gel documentation system (Alpha Innotech Corporation, San Leandro, California).

(Camden Instruments, Loughborough, UK). The slices were then bathed in aerated (95% O2, 5% CO2 normal ACSF (124 mM NaCl, 1.3 mM MgSO4, 3 mM KCl, 1.25 mM NaH2PO4, 25 mM NaHCO3, 2.4 mM CaCl2, and 10 mM glucose) for at least 1 h prior to electrophysiological recordings. In some experiments, MgSO4 was omitted to create Mg-free conditions. Electrophysiological recordings Whole-cell recordings were obtained from neurons in brain slices bathed in normal ACSF using a fixed stage microscope (Axioscop, Zeiss, Germany) and water immersion lenses. Patch electrodes used for whole-cell recording were pulled on a Flaming/Brown p-87 micropipette puller (Sutter Instrument, Novato, California) and filled with internal solutions either (i) 120 mM Cs-gluconate, 2 mM MgCl2, 0.5 mM EGTA, 10 mM HEPES, 10 mM TEA, 3 mM QX-314, 3 mM Na-ATP, pH 7.2, 270–280 mOsm for voltage clamp readings or (ii) 120 mM K-gluconate, 10 mM HEPES, 30 mM KCl, 0.2 mM EGTA, 3 mM Na ATP, 2 mM MgCl2, pH 7.2, 270–280 mOsm for current-clamp readings. The electrodes were positioned using a 3-axis MP-285 motorized micromanipulator (Sutter Instrument, Novato, California) for fine positioning and a mechanical manipulator, Newport MX 110 (Newport, Irvine, California) for coarse positioning. pCLAMP data acquisition software (Axon Instruments, Union City, California) and an Axopatch-1D patch clamp amplifier (Axon Instruments, Union City, California) were used to monitor electrode resistance in voltage clamp mode. Series of resistance were monitored throughout the experiment by measuring the instantaneous current response to a 10 mV voltage step from 70 mV. A gravity-fed array of inflow tubes of 0.58 mm inner diameter and an outflow tube attached to a vacuum reservoir provide solution flow which allowed the slice to be continuously perfused with oxygenated solution during the search for neurons, rapid solution changes, and a small volume when the flow was stopped during recording for exposure to calcium channel antagonists. The ground electrode was consisted of a salt bridge constructed from glass electrode filled with agar. Drug application

Brain slice preparation GSHPx-1 transgenic and WT animals (1–2 months old) were anesthetized with halothane vapors and decapitated. Brains were quickly removed and placed in cold lowsodium sucrose substituted saline artificial cerebral spinal fluid [sucrose ACSF: 90 mM NaCl, 105 mM sucrose, 1.3 mM MgSO4, 3 mM KCl, 1.25 mM NaH2PO4, 25 mM NaHCO3, 2.4 mM CaCl2, and 10 mM glucose, modified from Aghajanian and Rasmussen’s methods (Aghajanian and Rasmussen, 1989)]. 200–400 AM coronal sections of the brains through hippocampi were cut with a vibroslicer

100 AM NMDA were applied for 1 second to brain slices by using SF-77B Perfusion Fast Step (Warner Instrument, Hamden, CT). Control responses were evoked for 6 min to ensure the stability of the response to NMDA and then 5 mM DTT was perfused through the recording chamber. Responses were recorded for 6 min after the addition of DTT. DTT-mediated changes in the response to NMDA application were plotted over the time of DTT exposure as a percentage of the control NMDA response amplitude. For GSSG pressure application, 20 mM GSSG were applied by Pneumatic PicoPump (PV-820, World Precision Instru-

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ments, Saratosa, FL), at holding membrane potentials of 60, or 75 with constant current or a holding membrane potential of 80 mV with small hyperpolarizing pulses of current (200 ms). For GSSG bath application, 20 mM GSSG was applied at a holding membrane potential of 70 mV. For GSSG or sucrose perfusion application, 20 mM GSSG or 20 mM sucrose was applied by Perfuse Fast Step System (Warner Instruments, Hamden, CT) at a holding membrane potential of 55 mV. Voltage-clamp ramps were used to examine the voltage-dependency of NMDA and GSSG responses. Cells were held at a holding potential of 70 mV and the membrane potential was slowly ramped over a one second period to a final peak potential of +40 mV. The membrane potential was then repolarized to 70 mV using a 200 ms ramp. Net NMDA and GSSG induced currents were obtained by subtracting currents obtained from ramps delivered before agonist application from currents obtained after agonist application. Acute isolation of CA3 neurons Neurons were isolated using a procedure described earlier (Schumacher et al., 1998). The isolation procedure is as follows: Hippocampal slices were prepared in iced cold solution (in mM) NaCl 90; KCl 3; CaCl2 2; MgSO4 2; Na-pyruvate 1; HEPES 10; glucose 10; sucrose 105 (pH 7.4, 100% O2) and transferred to the ASCF. Isolation of viable neurons has been possible up to 10 h after the preparation of slices. After an equilibration period of 30–45 min the individual slice were transferred to a polystyrene tube with 5 ml of saline containing (in mM): NaCl 126; KCl 2,5; CaCl2 2; MgCl2 2; NaH2 PO4 1.25; PIPES 26; glucose 10 (pH 7.3, 100% O2). Pronase, 1.5 mg/ml (protease type XIV, Sigma) was added to the oxygenated medium. After an incubation period of 25–30 min at 328C, the slice was washed in the same ice-cold, enzyme-free saline. The CA3 region was dissected and triturated in 2 ml of ice-cold saline through fire polished Pasteur pipettes of decreasing apertures. The cells were then transferred to one compartment of two-compartment recording chamber and allowed to settle for 5–10 min. Neurons with pyramidal shaped soma were patched, lifted and transferred to the second small-volume (300–400 Al) compartment of recording chamber where they were perfused with the desired experimental solution.

Results Hippocampal GSHPx activity in HT (heterozygous, containing 1 copy of the murine GSHPx-1 transgene) mice demonstrated a four-fold increased in hippocampal GSHPx activity and HM (homozygous, containing 2 copies of the transgene) a six-fold increase compared to wildtype (WT) mice hippocampi (26.38 F 4.51 mU/mg protein) (Fig. 1). Higher levels of hippocampal GSHPx-1 in transgenic animals were hypothesized to be protective against KA

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Fig. 1. Hippocampal GSHPx activity in HM, HT and WT animals. GSHPx, glutathione peroxidase; HM, homozygous; HT, heterozygous; WT, wildtype. n = 4–5, *P = 0.01 HM vs. HT, **P = 0.01 HM and HT vs. WT.

treatment, yet surprisingly, more frequent and severe seizures as well as more neuronal cell death was observed in the hippocampi of HT and HM animals compared to WT (Figs. 2a–b). 8–12 weeks old animals were intraperitoneally injected with KA (35 mg/kg body weight) and for a period of 2 h immediately after KA administration, seizure activities were observed, scored and graphed. HM and HT mice showed a clear increase in the frequency and severity of seizures compared to WT animals (n = 3 of each genotype) (Fig. 2a). At 24 h after KA injection, brains were removed, fresh-frozen, sectioned, and stained with haematoxylin and eosin (H&E), cresyl violet (CV), and terminal deoxynucleotide transferase-mediated dUTP nick-end labeling (TUNEL) to assess extent of neuronal damage. Coronal brain sections demonstrated increased damage in area CA3 in HM and HT compared to WT mice, which has been shown to be the most vulnerable brain area to seizureinduced cell death (Ben-Ari, 1985) (Fig. 2b). Some cell damage was also observed in the CA2 in some cases (Fig. 2b). Overall, 63.41% of KA-treated HMs (26 out of 41), 48.84% of KA-treated HTs (21 out of 43) and 35.71% of KA-treated WTs (15 out of 42) displayed severe, prolonged seizures (grade 4) (Fig. 2c). Almost half of all KA-treated HMs (47.62%, 10 out of 21), 28% of KA-treated HTs (7 out of 25) and 19.05% of KA-treated WTs (4 out of 21) also demonstrated damage to the CA3 region as assessed by TUNEL staining (Fig. 2c). No significant differences in seizure activities or hippocampal damage were observed between male and female mice (data not shown). To study whether or not GSHPx-dependent susceptibility to KA toxicity also occurs in vitro, the effects of KA on dispersed hippocampal primary neuronal cultures from HMs and WTs were studied. Cultures were treated with various concentrations of KA (0–250 AM) and cell viability assayed 24 h following KA addition. At lower concentrations of KA (10, 25 and 50 AM), no significant differences between HM and WT were observed (Fig. 2d). However, at higher concentrations of KA (100 and 250 AM), a significantly reduced level of cell survival was observed in the HM cultures. In contrast, higher concentrations of KA did not increase the amount of cell death in WT cultures (Fig. 2d).

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Fig. 2. Increased GSHPx activity results in increased susceptibility to KA both in vivo and in vitro. (a) Representative seizure severity and frequency in HT and HM animals vs. WT (n = 3 each) over time following systemic KA administration in vivo. 0, no seizures; 1, very light seizures of forepaws; 2, repeated mild seizures, rearing and loss of postural control; 3, whole body seizures for a short time; 4, seizures become severe and prolonged. (b) Assessment of cell injury and/or loss in representative coronal sections from HT, HM, and WT hippocampi 24 h following systemic KA treatment. CV, cresyl violet; H&E, haematoxylin and eosin; TUNEL, terminal deoxynucleotide transferase-mediated dUTP nick-end labeling. (c) Percentages of KA-treated animals displaying severe, prolonged (grade 4) seizures and CA3 cell injury (based on TUNEL+ CA3 cell counts). White bars, HM; Grey bars, HT; Black bars, WT. P b 0.05 for HM and HT vs. WT in both cases. (d) Cell death in primary hippocampal neuronal cultures from HM vs. WT mice 24 h following KA treatment at dosages of 0, 10, 25, 50, 100 and 250 AM. Open bars, WT; filled bars, HM, n = 3–6 each, *P = 0.01 HM vs. WT.

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Interestingly, basal brain GSH/GSSG ratios were found to be significant reduced in HM and HT GSHPx-1 transgenics compared to WT prior to KA administration (Fig. 3b). This is likely due to increased conversion of GSH to GSSG via elevated hippocampal GSHPx levels (Mirochnitchenko et al., 1999). HPLC measurements of cellular GSSG levels were subsequently performed on hippocampal tissue isolated from HM vs. WT mice; levels of GSSG were found to be significantly elevated in the transgenics (Fig. 3f ). To investigate whether or not other enzymes involving in GSH conversion were altered in HM and HT animals vs.

Fig. 3. Elevation in intracellular GSSG levels in untreated hippocampi from GSHPx-1 transgenic vs. WT mice and in the extracellular media from primary transgenic vs. WT hippocampal cultures following KA administration. (a) Schematic of the GSH metabolic pathway. G-6-P, glucose-6-phosphate; Ru-5-P, ribulose-5-phosphate; G6PD, glucose-6phosphate dehydrogenase; 6PGD, 6-phosphogluconate dehydrogenase; GR, glutathione reductase. (b) GSH/GSSG ratios, n = 3–5, *P = 0.05 HM vs. HT; **P = 0.01 HM and HT vs. WT. (c) Glutathione reductase (GR) activity, n = 3–6, *P = 0.05 HM vs. HT; **P = 0.01 HM and HT vs. WT. (d) Glucose-6-phosphate dehydrogenase (G6PD) activity, n = 3–6, *P = 0.01 HM vs. WT. (e) 6-phosphogluconate dehydrogenase (6PGD) activity, n = 3–6, * P = 0.05 HT vs. WT, **P = 0.01 HM vs. WT. (f) HPLC measurement of intracellular GSSG levels in HM, HT and WT hippocampi, n = 3, *P = 0.05. (g) HPLC measurement of extracellular GSSG levels in the media of primary cultures derived from embryonic HM and WT hippocampi at 24 h after 0 or 50 AM KA treatment. Open bars, WT; filled bars, HM, n = 3–5, *P = 0.01.

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WT (see Fig. 3a for schematic of the glutathione metabolic pathway), hippocampal activity levels of glutathione reductase (GR), glucose-6-phosphate dehydrogenase (G6PD) and 6-phosphogluconate dehydrogenase (6PGD) were determined. GR activity levels in HM and HT were slightly but significantly higher than in WT (35.66% and 21.88% increase, respectively). However this is unlikely to compensate for the 400–600% increase in GSHPx activity in HT and HM transgenics, respectively (Fig. 1 vs. Fig. 3c). An approximately 10% decrease in G6PD activity levels was observed in HM compared to WT mice. There was no significant difference between HT and WT animals (Fig. 3d). Unlike G6PD, activity levels of 6PGD were decreased both in HM and HT compared to WT (Fig. 3e). Decreased G6PD and 6PGD could conceivably act to limit NADPH required for conversion of GSSG to GSH despite the small compensatory GR elevation. Application of KA or the glutamate receptor agonist (NMDA) have both been reported to result in increased extrusion of GSSG from cultured cells and acute hippocampal rat brain slices (Wallin et al., 1999). We therefore measured GSSG levels via HPLC in the extracellular media from primary cultures derived from GSHPx-1 vs. WT hippocampi following addition of KA. We found GSSG levels to be significantly increased in the media from GSHPx-1 cells vs. WT 24 h following 50 vs. 0 AM KA administration (Fig. 3g). Based on our data, increased GSHPx activity levels in the transgenic animals appears to be responsible for an elevation in extracellular GSSG levels following KA application. Elevated extracellular GSSG could make the mice more prone to seizures and hippocampal cell death (Bellissimo et al., 2001; Erden-Inal et al., 2002; Inal et al., 2001). We assessed possible direct interaction of extracellular GSSG with the NMDA receptor (Hiramatsu and Mori, 1981). Previously, it has been reported that exogenous application of GSSG and to a lesser extent GSH can stimulate calcium (Ca2+) entry in dissociated neurons which is inhibited by the NMDA receptor agonist MK801 (Leslie et al., 1992). GSSG has also been demonstrated to be capable of directly binding to NMDA receptors although the electrophysiological effects of this binding on receptor activity were not directly tested in these studies (Hermann et al., 2002; Janaky et al., 2000). GSHPx-mediated elevation of intracellular GSSG levels coupled with KA-induced extrusion of GSSG may result in an increase in GSSG available to bind and directly activate surface NMDA receptors. An electrophysiological study, performed on acute hippocampal slices from HM vs. WT animals, demonstrated that the NMDA-evoked response in HM slices was approximately 3 times higher than in WT slices (Fig. 4a, min 2, 4, 6 following NMDA application, HM = 2451.00 F 457.07 pA, 2496.64 F 458.82 pA, 2582.59 F 449.22 pA; WT = 845.10 F 278.70 pA, 844.15 F 239.18 pA, 799.22 F 250.53 pA, respectively). The NMDA receptor has been reported to contain a redox modulatory site which can

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Fig. 4. NMDA-evoked responses produced in CA3 neurons from WT and HM hippocampal slices. (a) Peak amplitude measurements of responses to NMDA from voltage clamped CA3 neurons. (b) DTT-induced changes in NMDA-evoked response in CA3 neurons from WT and HM mice. 100 AM NMDA was applied using the Perfusion Fast Step system. At minute 6, 5 mM DTT was added via Fast Step. Responses were recorded for 6 min after the addition of DTT, n = 3. Representative current traces from WT and HM CA3 neurons are shown.

effects its activity (Aizenman et al., 1990; Gilbert et al., 1991). To investigate if a change in redox modulation was responsible for the increase in NMDA-evoked activity observed in the GSHPx-1 transgenic hippocampus, 5 mM dithiothreitol (DTT) was applied to the slice cultures after minute 6. No difference in relative DTT-induced NMDAevoked activity was observed between WT and HM cultures suggesting that the GSHPx-mediated effects is independent of the NMDA receptor redox state (Fig. 4b). In addition, results from immunoblotting assays demonstrated no increase in numbers of NMDA receptor subunits expressed in the hippocampus including NMDAR1 (NR1), NMDAR2A (NR2A) nor NMDAR2B (NR2B) (data not shown) suggesting that the NMDA-evoked response is also separate from NMDA receptor numbers. To investigate the possibility that elevation in levels of extracellular GSSG could directly activate CA3 neurons, GSSG was directly applied to individual neurons in acute hippocampal brain slices from WT animals. Application of 10–20 mM GSSG via pressure pipette and recording of electrophysiological responses by whole-cell current clamp revealed large GSSG-evoked depolarizations generating a barrage of action potentials reminiscent of the bparoxysmal depolarizing shiftQ seen in models of epilepsy (Bradford, 1995; Johnston and Brown, 1984) (Fig. 5a). To eliminate the possibility of depolarization being generated by pressure pipette application, GSSG was also applied via bath perfusion. Similar results were obtained, that is, a slow depolarization associated with a series of action potentials (Fig. 5b). We tested the conductance change associated with the GSSG-mediated depolarization by passing constant current pulses through the recording pipette. We found the GSSG depolarization was associated with an increase in membrane resistance (the size of the voltage transient produced by the current pulses increased); this change is indicative of channel closures during the hyperpolarization (Mayer and Westbrook, 1985) (Fig. 5c). The effects of GSSG application could not be explained via osmotic effects as equi-molar sucrose application showed no response (Fig. 5d). While GSSG responses were blocked by the NMDA receptor

antagonist AP-5, they were not blocked by the AMPA receptor antagonist CNQX (Figs. 6A, B). The bursting behavior induced by GSSG application and the increase in resistance seen with hyperpolarizing pulses suggested GSSG could directly activate NMDA receptors. To test this hypothesis, we compared voltage-dependent and pharmacological properties of GSSG responses with those obtained for responses to direct application of NMDA in CA3 neurons. Individual CA3 neurons were voltageclamped at a holding potential of 70 mV and exposed to a 1 second depolarizing ramp which peaked at a membrane potential of +40 mV and then returned to 70 mV during a 200-ms repolarizing ramp. The neurons were then exposed to GSSG and the ramp was re-delivered. Subtraction of the control ramp from the ramp obtained after GSSG exposure revealed a voltage-dependent GSSG-induced current which looked similar to that induced by NMDA (Figs. 7a, b). Application of 100 AM AP-5 was able to block subsequent GSSG-evoked responses. Similarly, 5-aminophosphonovaleric acid (AP-5) blocked responses to NMDA application in the same cell (Figs. 7a, b). We also found that bathing neurons in an Mg2+-free solution eliminated the voltagedependence of the response to GSSG, again suggesting GSSG is activating NMDA receptors normally regulated by extracellular Mg2+. It is possible that GSSG could cause a release of glutamate and thus not be directly activating NMDA receptors. To eliminate this possibility, we examined the response of CA3 neurons to GSSG application in neurons acutely isolated from surrounding hippocampal tissue. This method allows recording from single CA3 neurons (cell body and small primary dendrite) in total isolation from tissue (other cells axons, synapses, dendrites, cell bodies). We found GSSG also evoked an inward current in these isolated neurons (Fig. 7c).

Discussion The glutathione (GSH/GSSG) redox status is important for maintaining a reducing environment in cells and tissues.

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Fig. 5. GSSG-evoked electrophysiological responses in WT CA3 neurons. (A) Response of CA3 neuron to pressure application of 20 mM GSSG (Vm = 75 mV). (B) Response of CA3 neuron to slow bath application of 20 mM GSSG. GSSG (20 mM) produced a depolarization and an associated barrage of action potentials (top trace, Vm = 60 mV). (C) Example of response to pressure application of GSSG (20 mM) in the same CA3 neuron held at membrane potentials of 60 mV (a) and 80 mV (b) using constant current injection. Small hyperpolarizing pulses of current (200 ms) were delivered at 80 mV to monitor changes in membrane resistance associated with the GSSG-induced depolarization. The GSSG response was associated with an increase in membrane resistance (the size of the voltage transients induced by the current pulses increased). (D) Application of 20 mM GSSG generates a depolarization and action potentials (a), while application of equimolar sucrose (b) did not produce a response (Vm = 55 mV).

Any alterations in the ratio of oxidized GSSG to reduced GSH could cause a shift in the cellular redox environment (Filomeni et al., 2002a,b; Schafer and Buettner, 2001) A growing body of evidence suggests that alterations in this redox state can have adverse effects. This has been suggested to be specifically due to either GSH depletion or increases in GSSG levels. For example, depletion of GSH and GSSG via either the GSH synthesizing inhibitor buthionine sulfoxamine (BSO) or diethyl maleate (DEM) in Bcl-2 overexpressing HL60 cells leads to increased ROS production and selective cell death (Armstrong and Jones, 2002). Similar reductions in GSH via 2-cyclohexen-1-one (CHX) has also been reported to enhance activator protein-1 (AP-1) DNA binding in the murine hippocampus and to result in more severe KA-induced seizures (Ogita et al., 2001). We examined whether increasing GSHPx activity would help to protect against damage induced by kainite

injections and surprisingly found the opposite effect, GSHPx-1 Tg mice showed greater seizures and associated brain damage following exposure to KA. During the course of our experiments, we discovered that exposure of GSHPx-1 Tg neurons to KA resulted in an increase in extracellular levels of GSSG and we hypothesized neuronal vulnerability to kainate could be linked to increased GSSG (Fig. 3). Elevated GSSG has also been reported to induce detrimental effects. Exogenous GSSG application, for example, has been shown to result in apoptosis in a promonocytic cell U937 by altering intracellular redox environment resulting in activation of the p38 mitogen protein pathway (Filomeni et al., 2002a,b). Biroccio et al. (2002) demonstrated that down-regulation of c-myc also induces program cell death in melanoma cells via a accumulation of intracellular GSSG. Cereser et al. (2001) demonstrated that thiram, a dithiocarbamate fungicide,

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CNQX. These data suggest that elevated GSSG may be involved in the initiation of seizures through direct activation of NMDA receptors. Elevated intracellular GSSG has been correlated with an increased risk of seizure generation in elderly patients vs. those in younger age groups (Erden-Inal et al., 2002; Inal et al., 2001). In addition, systemic pilocarpineinduced seizures in rats has been associated with increased GSHPx activity prior to status epilepticus although it is

Fig. 6. GSSG-induced response is not due to glutamate contamination. (A) Response to application of GSSG alone or after application of 10 AM CNQX and (B) the effects of GGSG alone vs. following addition of 50 AM of AP-5.

induces increased GSSG levels and lowers cell viability in human fibroblast cultures. It was hypothesized that the elevated intracellular GSSG could be transported in response to oxidative stress into the extracellular media and there act as a powerful pro-oxidizing agent (Filomeni et al., 2002a,b). In the current study, increased GSHPx activity was found to induce increased extracellular GSSG levels in response to KA administration in vitro. In addition, KA administration results in increased seizure activity and hippocampal neuronal cell death in vivo. GSSG itself was demonstrated to have direct electrophysiological effects via activation of NMDA receptors. Several groups reported that GSSG reduced NMDA responses via oxidation of the NMDA receptor (Gilbert et al., 1991; Janaky et al., 1993; Sucher and Lipton, 1991; Varga et al., 1997). We hypothesized increases in basal extracellular GSSG might modify the redox state of NMDA receptors in GSHPx-1 Tg’s, however, we found the thiol reducing agent DTT had an equal effect in Wt and GSHPx-1 Tg mice (Fig. 4). Prior work has demonstrated that GSSG binds to NMDA receptors and it increases intracellular Ca2+ via NMDA receptor activation (Hermann et al., 2002; Janaky et al., 2000; Leslie et al., 1992). We found GSSG elicited bursts of neuronal activity, reminiscent of that seen in electrophysiological models of epilepsy. Much like first described for NMDA-evoked activation of NMDA receptors by Mayer and Westbrook (1985), the voltage-dependent block of GSSG responses produced by Mg2+ acting on NMDA receptors could explain the apparent increase in membrane resistance produced by GSSG and the ability of GSSG to support repetitive firing. We also found removal of Mg2+ from the extracellular media eliminated the voltage-dependence of responses to GSSG and NMDA in the same cell and GSSG responses were blocked by the selective NMDA receptor antagonist AP-5 but not the AMPA receptor antagonist

Fig. 7. GSSG-evoked response acts via direct NMDA receptor activation in CA3 neurons from WT mice. Response to application of (a) 100 AM NMDA alone (NMDA) or 100 AM NMDA after application of 100 AM AP5 (AP-5). (b) Response to 20 mM GSSG alone (GSSG) or 20 mM GSSG after an application of 100 AM AP-5 (AP-5) (Perfusion Fast Step, holding potential = 70 mV, 1-s depolarizing ramp, peaked at +40 mV and returned to 70 mV during a 200-ms repolarizing ramp). (c) Response of an acutely isolated CA3 neuron to application of 20 mM GSSG (holding potential = 40 mV).

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unclear whether this was causative or compensatory (Bellissimo et al., 2001). In another study, epilepsy-prone mice were reported to have decreased cerebellar GSH/GSSG ratio both before and during seizure (Hiramatsu and Mori, 1981). Our study suggests that alterations in GSH/GSSG ratios via increased GSHPx-1 expression results in elevated extracellular GSSG levels which may directly interact with and activate NMDA receptors in turn resulting in increased seizure activity and CA3 hippocampal neuronal cell death associated with kainic acid administration. The increased seizure response in the animals may alternatively be due to an increase in NMDA receptor response separate from the observed increase in extracellular GSSG levels. We cannot totally exclude the possibility that our GSSG sample was contaminated by glutamate and that the electrophysiological responses we observed following GSSG application were due to this contamination. Previous TLC analysis of the compound by the vendor (Sigma) did not demonstrate any glutamate contamination; however, sensitivity of the assay may preclude detection of small levels of the compound. The lack of an electrophysiological AMPA response following GSSG application could also be attributed to a trace glutamate contamination as NMDA receptors are 100 times more sensitive to glutamate than AMPA receptors. In addition, the levels of GSSG used in our electrophysiological studies was likely higher than that found physiologically in the extracellular space although microenvironments near the membrane could exist following this extrusion at which this concentration is attained. However, Wallin et al. (1999) have demonstrated that levels of glutamate remain unchanged following hypoxia–ischemia so disintegration of GSSG to glutamate does not appear to occur. In conclusion, elevation of GSHPx-1 expression surprisingly results in increased rather than decreased KA susceptibility which correlates with elevated extracellular GSSG levels which in turn may directly activate the NMDA receptors. This may have major implications for the use of antioxidants as treatment for ischemia.

Acknowledgments This work was supported by NIH grants AG12141 (Andersen, J.K.) and DK45334 (Lu, S.C.) as well as the Cell Biology Core of the USC Research Center for Liver Diseases (P30 DK48522) for HPLC measurements. We would like to thank Dr. Y.S. Ho, Wayne State University, Detroit, Michigan for the gift of the transgenic animals.

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