Enhanced release of synaptic glutamate underlies the potentiation of oxygen–glucose deprivation-induced neuronal injury after induction of NOS-2

Experimental Neurology 190 (2004) 91 – 101 www.elsevier.com/locate/yexnr

Enhanced release of synaptic glutamate underlies the potentiation of oxygen–glucose deprivation-induced neuronal injury after induction of NOS-2 Aniruddha S. Vidwansa,b, Sandra J. Hewetta,* a

Department of Neuroscience, University of Connecticut Health Center, Farmington, CT 06030-3401, USA Department of Pediatrics, Division of Neonatology, University of Connecticut Health Center, Farmington, CT 06030-2948, USA

b

Received 24 April 2004; revised 5 June 2004; accepted 10 June 2004 Available online 2 September 2004

Abstract Reactive nitrogen oxide species (RNOS) may contribute to the progression/enhancement of ischemic injury by augmentation of glutamate release, reduction of glutamate uptake, or a combination of both. Consistent with this, induction of nitric oxide synthase (NOS-2) in murine neocortical cell cultures potentiated neuronal cell death caused by combined oxygen–glucose deprivation in association with a net increase in extracellular glutamate accumulation. However, uptake of glutamate via high affinity, sodium-dependent glutamate transporters was unimpaired by induction of NOS-2 under either aerobic or anaerobic conditions. Further, blocking possible routes of extra-synaptic glutamate release with NPPB [5-nitro-2-(3-phenylpropylamino)-benzoic acid], a volume-sensitive organic anion channel blocker, or TBOA (d,l-threoh-benzyloxyaspartate), an inhibitor of glutamate transport, exacerbated rather than ameliorated injury. Finally, treatment with riluzole or tetanus toxin attenuated the enhancement in both glutamate accumulation and oxygen–glucose deprivation-induced neuronal injury supporting the idea that increased synaptic release of glutamate underlies, at least in part, the potentiation of neuronal injury by RNOS after NOS-2 induction. D 2004 Elsevier Inc. All rights reserved. Keywords: Hypoxia/hypoglycemia; Cerebral ischemia; Mixed cortical cell cultures; Excitotoxicity; Inducible nitric oxide synthase; Neurodegeneration

Introduction While excitotoxicity is a recognized trigger of damage in the cerebral ischemic core, evidence from both animal and human studies indicates that much of the cortical neuronal Abbreviations: DETA/NO, {(z)-1-[(2-aminoethyl)-N-(2-ammonioethyl)-amino]diazen-1-ium-1,2-diolate}; EGF, epidermal growth factor; IFN-g, Interferon-g; LDH, lactate dehydrogenase; LPS, lipopolysaccharide; NPPB, [5-nitro-2-(3-phenylpropylamino)-benzoic acid]; RNOS, reactive nitrogen oxide species; SIN-1, 3-morpholinosydnonimine hydrochloride; SNAP, S-nitroso-acetyl-d,l-penicillamine; TBOA, d,l-threo-h-benzyloxyaspartate; TeNT, tetanus toxin; VSOAC, Volume-sensitive organic anion channels. * Corresponding author. Department of Neuroscience, University of Connecticut Health Center, MC-3401, 263 Farmington Avenue, Farmington, CT 06030-3401. Fax: +1 860 679 8766. E-mail address: [email protected] (S.J. Hewett). 0014-4886/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.expneurol.2004.06.018

damage resulting from stroke is secondary and outside this primary focus of damage with the process of tissue destruction lasting for hours to days (Castillo et al., 1997; Du et al., 1996; Feelisch et al., 1989; Garcia et al., 1993; Heiss et al., 1992; Knight et al., 1994). The mechanisms by which progression of neuronal injury occurs are incompletely understood. Some studies suggest that temporal changes in ischemic neuronal death may simply be due to a slow progression of excitotoxicity (Gill et al., 1992; Hossmann, 1994). Others studies demonstrate that these changes result from acute inflammatory mechanisms initiated during the post-ischemic period [for review, see (del Zoppo et al., 2000)]. Indeed, activation of the proinflammatory, inducible isoform of NOS (NOS-2) has been shown to contribute to cerebral ischemic injury progression (Iadecola et al., 1995, 1997; Parmentier-Batteur et al., 2001).

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Our previous studies indicate that these two pathways need not be mutually exclusive (Hewett et al., 1994, 1996). Using a murine in vitro co-culture system, we demonstrated that NO elaborated by NOS-2 in cytokine-activated astrocytes potentiated neuronal injury induced by the excitatory amino acid, N-methyl-d-aspartate (NMDA) (Hewett et al., 1994) or by combined oxygen–glucose deprivation (Hewett et al., 1996). Enhanced injury in the latter model was associated with a large net increase in extracellular glutamate accumulation and was prevented by NMDA receptor antagonism (Hewett et al., 1996). So, nitric oxide (NO) or related reactive nitrogen oxide species (RNOS) derived from NOS2 may contribute to the excitotoxic process after oxygen– glucose deprivation by enhancing glutamate release, reducing its cellular reuptake or a combination of both. Thus, the aim of this study was to test whether endogenously produced RNOS species derived from NOS-2 inhibit the function of glutamate transporters and/or enhance synaptic or extrasynaptic release of glutamate.

Material and methods Materials Antibodies directed against VAMP-2 (synaptobrevin-2) and NOS-2 were purchased from Synaptic Systems (Gfttingen, Germany) and Upstate Cell Signaling Solutions (Lake Placid, NY), respectively. 2,4-Pyrrolidone dicarboxylic acid (2,4-PDC) and 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) were purchased from Biomol Research Laboratories (Plymouth Meeting, PA). Tris, leupeptin, diaphorase, and aprotinin were obtained from Roche Diagnostics Corp (Indianapolis, IN). Lipopolysaccharide (LPS) was purchased from DIFCO laboratories, (Detroit, MI). Recombinant mouse Interferon-g (IFNg) was obtained from R&D systems (Minneapolis, MN). DETA/NO {(z)-1diazen-1-ium-1,2-diolate}, SIN-1 (3-morpholinosydnonimine hydrochloride) and SNAP (S-nitroso-N-acetyl-d,lpenicillamine) were purchased from Alexis Biochemicals (San Diego, Ca). Tetanus toxin (clostridium tetani) was purchased from Calbiochem (La Jolla, CA). Riluzole and d,l-threo-h-benzyloxyaspartate (TBOA) were obtained from Tocris, (Ballwin, MO). Pyruvate, glutamate, h-NADH, cytosine h-d-arabinofuranoside (Ara-C), h-NAD+, glutamate dehydrogenase (GDH), Nonidet P-40 (NP40), deoxycholate, p-iodonitrotetrazolium violet (INT) and MEM amino acids (50) were purchased from Sigma, (St. Louis, MO). Iodoacetamide and triethanolamine chloride were obtained from Acros Organics (Morris Plains, NJ). Phenylmethanesulfonyl fluoride (PMSF) was obtained from Boehringer Mannheim (Mannheim, Germany). Epidermal growth factor (EGF) and antibiotics were purchased from Gibco/BRL, (Life Technologies, Grand Island, NY). Modified Eagle’s medium (MEM, Earle’s salts) was obtained from Mediatech (Herndon, VA). All sera were obtained

from Hyclone (Logan, UT). 3H-d-aspartic acid was purchased from NEN Life Science Products (Boston, MA). Cell culture Primary murine mixed cortical cell cultures containing both astrocytes and neurons were prepared from either CD1 or Swiss Webster mice (Charles River Laboratories, Wilmington, MA) as described in detail previously (Trackey et al., 2001; Vidwans et al., 1999). Briefly, cortical astrocytes were first prepared from postnatal (1–3 days) animals. Following an aseptic dissection of cerebral cortices, dissociated cells were plated (1–1.5 hemispheres/ 10 ml/plate) in 15 mm 24-well plates (Falcon Primaria; Becton Dickinson, Lincoln Park, NJ) in a media stock (MS; see below) containing 10% fetal bovine serum (FBS), 10% iron-supplemented calf serum (CS), 50 I.U./ml penicillin, 50 Ag/ml streptomycin and 10 ng/ml EGF. Once confluent and thus contact inhibited, astrocyte cultures were treated with 8 AM Ara-C for 48–72 h to inhibit the growth of any rapidly dividing contaminating cells such as microglia. Hereafter, cultures were maintained in a maintenance medium (MS plus 10% CS and antibiotics). Cortical neurons were cultured from embryonic (day 15) animals using an identical dissection protocol. Dissociated cells were placed into MS supplemented with 5% FBS, 5% CS and antibiotics, and plated (3–3.75 hemispheres/10 ml/plate) on an established bed of astrocytes. After 6–7 days in vitro, mixed cultures were exposed to 8 AM Ara-C for 48 h. Cells were subsequently shifted into maintenance medium. The medium was changed twice weekly. All cultures were kept at 378C in a humidified 6% CO2-containing atmosphere. These cultures contain an approximate 50:50 neuron to astrocyte ratio and a negligible amount of microglia (Hewett et al., 2000). Experiments were performed on cortical cultures after 14–15 days in vitro. MS consisted of glutamine-free EMEM (Earle’ salt) supplemented with lglutamine, glucose, and sodium bicarbonate to a final concentration of 2.0, 25.7, and 28.2 mM respectively. NOS-2 induction To induce NOS-2, cultures were exposed to LPS (2 Ag/ ml) and IFN-g (1 ng/ml) in MS containing 2% FBS at 378C in a humidified 6% CO2-containing atmosphere for 24–36 h before other experimental manipulations. Both LPS and INFg are required to maximally induce NOS-2 (Hewett et al., 1993b; Vidwans et al., 2001), which we previously determined to occur in glia but not neurons (Hewett et al., 1993a, 1994), without causing direct damage to either mixed cortical cell cultures or astrocytic cultures over a 48-h time period (Hewett et al., 1994). Induction was confirmed herein in some experiments via assessment of NOS-2 protein via Western Blot analysis and in all experiments by measurement of nitrite, a stable oxidation breakdown product of NO (Green et al., 1982).

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Nitrite levels were determined by mixing 100 Al portions of culture media with 100 Al of Greiss reagent (one part 1% sulfanilamide in 60% acetic acid plus one part 0.1% naphthylenediamine dihydrochloride in distilled water) and absorbance at 540 nm was determined on a microtiter plate reader (Thermolabs, Chantilly, VA). Only cultures that demonstrated accumulated nitrite at least four times over levels determined in medium alone (negligible) were used for further experimentation. Exposure to RNOS compounds All compounds were prepared as 10 mM stock solutions in MS immediately before experimentation and diluted into the desired medium to their final concentration. Cells were exposed for varying periods of time at 378C in a humidified 6% CO2-containing incubator and subsequently viewed under phase contrast microscopy to confirm lack of gross neuronal damage before any further experimentation. DETA/NO releases NO
upon decomposition (Maragos et al., 1991). SNAP is an S-nitrosothiol and therefore an NO+ equivalent (Arnelle and Stamler, 1995). SIN-1 provides a continuous source of NO
and O2
(Feelisch et al., 1989) producing OONO under these experimental conditions (Trackey et al., 2001). The reported t1/2 for these compounds at 378C (pH 7.4) are as follows: DETA/NO c20 h (Keefer et al., 1996); SNAP c6–8 h (Feelisch and Stamler, 1996); and SIN-1 c8 h (Manzoni et al., 1992). Combined oxygen–glucose deprivation Cultures and experimental solutions were placed in a Forma Scientific Anaerobic Station (Marietta, OH) that utilizes an anaerobic gas mixture of 80% N2, 5% CO2 and 15% H2. This chamber has an interchange intake system that, using a vacuum system, cycles with 100% nitrogen (2) followed by the anaerobic gas mixture (1) for a total cycling time of three min, thus removing any room air. As such, transfer of cells and solutions to and from the workstation occurs without contaminating the work chamber with atmospheric gas. In addition, the work chamber contains a palladium catalyst wafer that will remove any trace O2 by coupling it with H+. A dessicant wafer then absorbs any H20 that might be made during this procedure. An oxygen electrode (OM-4; Microelectrodes, Inc; Bedford, NH) calibrated to room air remains in the chamber so that we can monitor chamber oxygen levels at all times. Once inside the chamber, all solutions were thoroughly deoxygenated by bubbling with the anaerobic gas mixture for 10 min. This duration of bubbling effectively reduces the oxygen content of the solution to 0 ppm (Fogal and Hewett; unpublished observations). Cell culture media were then replaced by thorough exchange (3  750 Al) with this deoxygenated, glucose-free balanced salt solution (BSS0; see below) with or without drugs as described and then placed in a 378C humidified incubator within the chamber

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for 25 or 40–45 min, as specified. Deoxygenated glucosecontaining BSS (20 mM, BSS20) served as a control since the presence of glucose prevents neuronal injury even in the absence of oxygen. Cultures were subsequently removed from the incubator and 100 Al of cell culture supernatant collected to assess early cell death, should it have occurred, using the lactate dehydrogenase (LDH) assay (described below). Exposure medium was then exchanged with oxygenated MS and cells returned to a normoxic (21% O2) incubator at 378C. Twenty–24 h later, an additional 100 Al of cell culture supernatant is collected to assess delayed neuronal death. LDH values determined immediately after and 20–24 h following oxygen–glucose deprivation were summed to determine the total neuronal cell death. The balanced salt solution contained (mM) 116 NaCl, 5.4 KCl, 0.8 MgSO4, 1.0 NaH2PO4, 26.2 NaHCO3, 1.8 CaCl2, 1 MEM amino acids, and 0.01 glycine. Assessment of neuronal cell death Neuronal cell death was estimated by examination of cultures under phase-contrast microscopy and quantified by measurement of LDH released by damaged or destroyed cells into the bathing medium as described in detail (Uliasz and Hewett, 2000). If present, the small amount of LDH in the medium of sister cultures subjected to sham-wash was subtracted from the levels in experimental conditions to yield the LDH signal specific to experimental injury. Data are expressed as the percentage of total neuronal LDH (=100%) that was determined for each experiment by assaying the supernatant of sister cultures after 24 h exposure to 300 AM NMDA, which produces a neuron selective injury as cortical astrocytes lack NMDA receptors (Backus et al., 1989; Chan et al., 1990; Janssens and Lesage, 2001) and are not injured following NMDA exposure (Choi et al., 1987). Of note, under conditions of combined oxygen–glucose deprivation alone with or without NOS-2 induction, we see no evidence of astrocyte injury immediately following or 20–24 h later as assessed by phase-contrast microscopy and confirmed by trypan blue exclusion. However, the addition of certain drugs did potentiate the injury to a great enough extent that astrocytic injury occurred. Thus, when LDH values greater than 100% are reported, this indicates that all the neurons and some of the astrocytes were destroyed. Quantitation of extracellular glutamate Samples of the extracellular medium, collected before the termination of oxygen–glucose deprivation, were assayed for glutamate using a colorimetric endpoint assay. To generate samples, cultures were washed into a phenol redfree deoxygenated BSS solution (essentially as described above save for amino acids). After 25 min, culture supernatant (300 Al) was collected and stored at 208C until analyzed. This time point was chosen for while neurons

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were swollen, they appeared intact under phase contrast microscopy and no LDH activity was detected in the cell culture medium. After confirming absence of LDH leakage in all samples, 200 Al was boiled (95–1008C; 5 min) to inactivate any enzymes present in the cell culture supernatant that use NAD+ or NADH as cofactors and as such might interfere with the assay. Samples were allowed to cool to room temperature following which 100 Al of a reaction cocktail, prepared by combining in a 3:1:1 ratio an aqueous solution of 5 mM K2HPO4, 0.25% Triton X-100, and 160 mM triethanolamine chloride (pH 8.6), an aqueous solution of 30 mM NAD+ and 1.6 U/ml diaphorase (made fresh daily), and an aqueous solution of 1.2 mM p-INT, was added. Glutamate dehydrogenase was then added to this mixture to a final concentration of 30 U/ml. After incubation for 45 min at room temperature, absorbance at 490 nm was spectrophotometrically measured using a MRX microtiter plate reader (Thermolabs). Measurement of glutamate transport 3

H-d-aspartate was used as the substrate for the high affinity glutamate transporters as it is not metabolized by astrocytes, nor is it a substrate of the synaptic vesicle uptake system (Drejer et al., 1983; Tabb and Ueda, 1991). Initial experiments determined that 3H-d-aspartate uptake was sodium-dependent, linear over the experimental time points chosen in both the aerobic and anaerobic environments (Fig. 5), and blocked in a concentration-dependent manner by the glutamate transporter inhibitors 2,4-PDC (data not shown) and TBOA (Fig. 6 inset). To measure 3Hd-aspartate uptake under aerobic conditions (i.e., in the presence of both oxygen and glucose), cells were washed into a HEPES-buffered salt solution (HBSS) containing (mM) 120 NaCl; 5.4 KCl; 0.8 MgCl2; 1.8 CaCl2; 20 HEPES; 15 glucose, and 0.01 glycine (pH 7.4) and allowed to equilibrate while shaking for 10 min at room temperature. 3H-d-aspartate (250 Al), adjusted to either 0.1 or 0.25 ACi/ml with 1 AM non-radioactive d-aspartate, was then added to culture wells. To measure 3H-d-aspartate uptake under anaerobic conditions, cells were moved into the anaerobic chamber and washed into either BSS0 (deprived of both oxygen and glucose) or BSS20 buffers (deprived of oxygen only as solution contains 20 mM glucose). Forty min later, 250AL of buffer containing 3Hd-aspartate (0.25 ACi/ml) and 1 AM non-radioactive daspartate was added to culture wells. Uptake was terminated in both experimental paradigms after 5 min by washing (3  750 Al) with ice-cold sodium-free choline stop buffer containing (mM) 116 choline chloride, 0.8 MgSO4, 1 KH2PO4, 10 HEPES, 5 KOH, 10 glucose, 0.9 CaCl2, and 5 non-radioactive d-aspartate. Culture wells were subsequently aspirated dry and cells lysed by addition of 400 Al of warm 0.2% sodium dodecyl sulfate (SDS). The amount of accumulated radioactivity was estimated in an aliquot of cell lysate (200 Al) using a

Packard Tricarb 4000 scintillation counter. Data are expressed as either uptake/mg protein or as the percentage of uptake in untreated cultures (set to 100%), which was determined in each experiment. Western blot analysis Cells were washed twice with ice-cold PBS followed by addition of 100 Al lysis buffer to each culture well. Lysis buffer contained 1% NP40, 5 mM EDTA, 10 Ag/ml each of leupeptin and aprotinin, 5 mM iodoacetamide and 1 mM PMSF. Cultures were placed on ice while rocking for 30 min and then cell lysates were collected following centrifugation 10,000  g; 5 min. Protein lysates (20 Ag as determined with BCA Assay; Pierce Chemical, Rockford, IL) were separated by SDS-PAGE (8% or 15% polyacrylamide gel for NOS-2 and synaptobrevin, respectively), and electrophoretically transferred to nitrocellulose. Membranes were incubated with primary antibodies (anti-NOS-2, 1:1000; anti-synaptobrevin-2, 1:2500) overnight at 48C. Blots were further processed using the Western Breeze Chemiluminescent Immunodetection protocol per manufacturer’s instructions (Invitrogen, Carlsbad, CA) and visualized on x-ray film (Hyperfilm, Amersham, Arlington Heights, IL).

Results We previously demonstrated that mixed cortical cell cultures deprived of oxygen and glucose for 40–50 min develop a moderate level of neuronal cell death by the next day, that this neuronal injury is markedly enhanced by prior induction of NOS-2, and occurs in association with a corresponding increase in [glutamate]e. Under conditions of combined oxygen–glucose deprivation alone with or without NOS-2 induction, we see no evidence of astrocyte injury immediately following or 20–24 h later as assessed by phase-contrast microscopy and confirmed by trypan blue exclusion (data not shown). A good correlation between NO formation, extracellular glutamate accumulation, and neuronal death exists such that NOS inhibition attenuates NO formation, glutamate accumulation, and potentiated neuronal injury in parallel (Hewett et al., 1996). To assess whether synaptic glutamate release contributed to the enhancement of [glutamate]e demonstrated following NOS-2 induction, cultures were treated with the sodium channel blocker, riluzole (10AM) during the oxygen-deprivation period only. Riluzole ameliorated oxygen–glucose deprivation-mediated neuronal injury in both control cultures and in cultures induced to express NOS-2 (Fig. 1A) with extracellular glutamate levels reduced in parallel (Fig. 1B). Importantly, this concentration of riluzole did not prevent neuronal injury induced by exposure to NMDA (100 AM; 5 min) directly (data not shown). The NOS-2-mediated enhancement of extracellular

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ured following induction of NOS-2 might result from a combination of glutamate release (suggested by the data above) coupled with a simultaneous diminution in glutamate uptake. Thus, we assessed transporter function by measurement of 3H-d-aspartate uptake. Interestingly, overall function of the transporters in our mixed cortical cell culture system was not decreased at any time point tested by prior induction of NOS-2 when measured under either aerobic (Table 1) or anaerobic conditions (Table 1; Fig. 4). Concentrations of non-radioactive d-aspartate in our experiments (1 AM) were not rate limiting as experiments performed using 10, 50, and 100 AM of unlabeled daspartate, concentrations which span the K m of the neuronal and astrocytic glutamate transporters, gave identical results (Fig. 5); data not shown). A concentration

Fig. 1. Inhibition of glutamate release by riluzole reduces the NOS-2mediated potentiation of oxygen-glucose deprivation-induced neuronal injury. (A) Following overnight incubation with media alone ( Induction) or medium containing LPS (2 Ag/ml) plus IFN-g (1 ng/ml) to induce NOS-2 (+ Induction), cultures were deprived of oxygen and glucose (45 min) in the absence or presence of riluzole (10 AM). The amount of cell injury assessed via measurement of LDH released into the cell culture supernatant immediately after combined oxygen–glucose deprivation, as well as 20– 24 h later was summed to reflect total neuronal cell death, which is expressed as a percentage of total neuronal LDH (mean + SEM; n = 6 from three different experiments). (B) In a parallel set of cultures, samples for the measurement of glutamate accumulation in the bathing medium were collected 25 min following the initiation of oxygen–glucose deprivation (25 min). Data are expressed as mean + SEM (n = 6 from 3 separate experiments). An asterisk (*) denotes a significant NOS-mediated increase in cell death and glutamate accumulation when compared to uninduced cultures (between group comparison) while a pound sign (#) represents a significant riluzole-mediated diminution (within group comparison), as determined by a two-way ANOVA followed by Bonferroni’s t test for multiple comparisons. Significance was assessed at P b 0.05.

glutamate levels, as well as neuronal cell death was also prevented by tetanus toxin (TeNT), a clostridium toxin that cleaves synaptobrevin-2 and thus prevents synaptic vesicle exocytosis (Schiavo et al., 1992) (Fig. 2). We have determined previously that this form of tetanus toxin effectively cleaves neuronal but not astrocytic synaptobrevin-2 (Fig. 3A) (Taylor and Hewett, 2002). Additionally, it did not alter glial NOS-2 protein induction or the release of NO (Figs. 3B and C). Thus, these observations indicate that synaptic release of glutamate does indeed contribute to the potentiation of neuronal injury by RNOS after NOS-2 induction. As extracellular glutamate levels are normally efficiently controlled by glutamate transporters expressed predominantly by astrocytes but also in neurons, the resultant increase in extracellular glutamate levels meas-

Fig. 2. Inhibition of glutamate release by tetanus toxin reduces the NOS-2mediated potentiation of oxygen–glucose deprivation-induced neuronal injury. (A) Mixed cortical cell cultures were treated overnight with medium alone, or medium containing tetanus toxin (0.3 Ag/ml) alone, LPS (2 Ag/ml) plus IFN-g (1 ng/ml), or all three compounds, then deprived of oxygen and glucose for 40 min. The amount of cell injury assessed via LDH measured immediately after oxygen–glucose deprivation, as well as 20–24 h later was summed to reflect the total amount of neuronal cell death. Data are expressed relative to the LDH signal corresponding to complete neuronal cell death (=100%, measured after exposure of sister cultures to 300 AM NMDA for 20–24 h) (n = 6 from 3 different experiments) (B). In a parallel set of cultures, samples for the measurement of glutamate accumulation in the bathing medium were collected 25 min following the initiation of oxygen–glucose deprivation. Data are expressed as mean + SEM (n = 6 from 3 different experiments). An asterisk (*) denotes a significant NOSmediated increase in cell death and glutamate accumulation when compared to uninduced cultures (between group comparison) while a pound sign (#) represents a significant reduction as a result of TeNT exposure. Data were analyzed by two-way ANOVA followed by Bonferroni’s t test for multiple comparisons. Significance was assessed at P b 0.05.

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A.S. Vidwans, S.J. Hewett / Experimental Neurology 190 (2004) 91–101 Table 1 Effect of RNOS on 3H-d-aspartate uptake in mixed cortical cell cultures 3

H-d-aspartate uptake (% of control)

+ [Oxygen and glucose] Pre-treatment LPS +IFNg DETA/NO (0.3 mM) SNAP (0.3 mM) SIN-1 (1 mM)

Fig. 3. Tetanus toxin effectively cleaves synaptobrevin-2 but does not interfere with NOS-2 induction or NO synthesis: Mixed cortical cell cultures were treated with medium alone, or medium containing tetanus toxin (0.3 Ag/ml), LPS (2 Ag/ml) plus IFN-g (1 ng/ml), or all three. (C) Thirty hours later, cell culture supernatants were collected and nitrite accumulation was determined as a measurement of NOS-2 catalytic activity (n = 6 from 3 different experiments). An asterisk (*) indicates values that are significantly greater than basal levels as determined by one-way ANOVA followed by Dunnett’s t test for comparison to control. Significance was assessed at P b 0.05. (A, B) Total cellular protein from the same cells as used in panel C was isolated and 20 Ag separated by SDSPAGE. Western blot analysis was performed using antibodies specific for Synaptobrevin-2 (VAMP-2) (A) or NOS-2 (B). Results are representative of three independent experiments.

of 1 mM unlabeled d-aspartate did effectively diminish measurable 3H-d-aspartate uptake in both the control and induced cultures, thus indicating that mM concentrations of glutamate would be necessary to compete with our tracer for the transporters (data not shown). The inability of endogenous NOS-2-derived RNOS to inhibit transporter function in either environment was confirmed by the use of various compounds that release alternate redox forms of NO, namely OONO (SIN-1; 0.03–1 mM), NO+ (SNAP; 0.03–0.3 mM) or NO
(DETA-NO; 0.03–0.3 mM): none inhibited 3H-d-aspartate uptake at any concentration tested (Table 1). Glutamate transporter function could be inhibited, however, by exogenous addition of arachidonic acid (c25% inhibition at 10 AM) or H202 (c25% inhibition at 3 mM), as was reported previously (Barbour et al., 1989; Volterra et al., 1992, 1994b), indicating that our assay was sensitive enough to detect a change should it have occurred (data not shown). While the lack of effect of RNOS on 3H-d-aspartate uptake suggested that glutamate transporter dysfunction did not contribute to the enhancement of glutamate accumulation that followed combined oxygen–glucose deprivation, there remained the possibility that transporters could contribute by reversing, thereby releasing glutamate into the extracellular space (Li et al., 1999; Obrenovitch, 1996; Phillis et al., 2000). However, exposure of mixed cortical cell cultures to TBOA, which

[Oxygen and glucose]

4h n.d. 98.1 F 0.3

24 h 102.8 F 1.1 114.8 F 0.8

24 h 94.3 F 2.2 125.2 F 2

113.7 F 4 100.9 F 0.2

108.5 F 2.3 101.9 F 0.6

100.1 F 7.4 95.2 F 8.5

NOS-2 was induced in mixed cortical cell cultures (LPS + IFNg; 24 h exposure) or cultures were exposed to RNOS generating compounds for either 4 or 24 h as indicated. 3H-d-aspartate uptake (5 min) was initiated under normoxic [+ oxygen and glucose] or hypoxic/hypoglycemic [ oxygen and glucose] conditions as described in methods. Concentration– response curves for each RNOS generating compound were performed (0.03–0.3 mM for DETA/NO and SNAP; 0.03–1 mM for SIN-1). As there was no significant decrease in uptake at any concentration tested as determined by ANOVA results for only the highest concentration of each compound are shown. Values represent the mean F SEM (n = 4 – 8 culture wells from two separate experiments) scaled to control (untreated = 100%). n.d. = not determined.

inhibits both uptake (Shimamoto et al., 1998) (Fig. 6 inset), as well as reversal of glutamate transporters (Waagepetersen et al., 2001), did not prevent but actually potentiated oxygen–glucose deprivation-induced neuronal injury in both control and induced cultures alike (data not shown). In fact, exposure of cultures to TBOA for as little as 35 min under normal conditions of oxygen and glucose, directly resulted in neuronal injury that was more pronounced in cultures that were induced to express NOS-2 (Fig. 6). Because this TBOA-mediated toxicity in

Fig. 4. NOS-2 induction does not alter 3H-d-aspartate uptake into mixed cortical cell cultures. NOS-2 was induced in mixed cortical cell cultures by exposure to LPS (2 Ag/ml) plus IFN-g (1 ng/ml). After confirmation of NOS-2 induction, cultures were deprived of oxygen and glucose for varying periods of times as indicated. During the last 10 min of deprivation, 3H-daspartate was spiked into the culture wells, uptake was then terminated and the amount of 3H-d-aspartate that accumulated into cells measured via scintillation counting. Data are expressed as mean F SEM cpm/mg protein (n = 4 cultures pooled from two separate experiments). There were no statistically significant between or within group differences in uptake as determined by two-way ANOVA.

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both control and induced cultures was prevented by either pretreatment with tetanus toxin or by simultaneous application of MK-801, an NMDA receptor antagonist (Wong et al., 1986), we conclude that that block of reuptake of synaptically released glutamate and subsequent over-stimulation of NMDA receptors underlies TBOA toxicity (Fig. 6). Volume-sensitive organic anion channels (VSOACs) have been shown to be portals of glutamate release into extracellular space under various pathological conditions (Basarsky et al., 1999; Kimelberg et al., 1990; Rutledge et al., 1998). Thus, if glutamate release through VSOACs contributes to the NOS-2-mediated enhancement of extracellular glutamate levels under the condition of combined oxygen–glucose deprivation, then the VSOAC antagonist NPPB (Crepel et al., 1998) should ameliorate the potentiation of neuronal injury demonstrated under these conditions. However, treatment of our cultures with NPPB (10–300 AM), like TBOA, actually enhanced rather than inhibited oxygen–glucose deprivation-mediated injury in both unin-

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Fig. 6. Inhibition of glutamate transport function unmasks a NOS-2mediated neurotoxicity. Following 28–36 h incubation with medium alone [ Induction], or medium containing tetanus toxin (0.3 Ag/ml) alone, LPS (2 Ag/ml) plus IFN-g (1 ng/ml) [+ Induction], or all three compounds, cultures were exposed to increasing concentrations of TBOA with or without MK-801 (5 AM) for 35 min. The amount of cell injury, assessed via measurement of LDH released into the cell culture supernatant immediately after TBOA exposure, as well as, 20–24 h later was summed to reflect total neuronal cell death. Death is expressed as percentage of total neuronal LDH (=100) measured after exposure to 300AM NMDA for 24 h. An asterisk (*) indicates values significantly different from basal (0 mM), (within group comparison), while a pound sign (#) indicates significant enhancement of injury in NOS-2-induced conditions compared to cultures that were uninduced (between groups comparison), which was determined via twoway ANOVA followed by Bonferroni’s t test for multiple comparisons (mean + SEM; n = 6 cultures per condition pooled from three different experiments). Significance was assessed at P b 0.05. INSET; TBOA inhibits 3H-d-aspartate uptake in a concentration-dependent manner. 3H-daspartate uptake (5 min) into mixed cortical cell cultures was measured in the absence or presence of increasing concentration of TBOA. Values represent the mean F SEM; n = 9–12 cultures pooled from three separate experiments scaled to untreated control [0 mM = 100%]. An asterisk (*) denotes a significant TBOA-induced diminution of 3H-d-aspartate uptake as compared to control, as determined via one-way ANOVA followed by a Dunnett’s t test. Significance was assessed at P b 0.05.

duced (31 F 10% vs. 115 F 9% cell death; 10 AM; P b 0.05) and induced cultures (87 F 10% vs. 124 F 10%; 10 AM; P b 0.05).

Discussion Fig. 5. Time course of 3H-d-aspartate uptake under oxygen–glucose deprivation. NOS-2 was induced in mixed cortical cell cultures by exposure to LPS (2 Ag/ml) plus IFN-g (1 ng/ml). After confirmation of NOS-2 induction, cultures were deprived of oxygen and glucose for 50 min. At the specified time points before termination of oxygen–glucose deprivation, 3 H-d-aspartate with either 1 AM (A) or 50 AM (B) non-radioactive aspartate was spiked into the culture wells. Uptake was terminated at 50 min and the amount of 3H-d-aspartate that accumulated into cells estimated via scintillation counting. Data are expressed as mean F SEM cpm/mg protein (n = 4 from two different experiments). No statistically significant differences were found between the induced and uninduced cultures as analyzed via two-way ANOVA.

Glutamate is the major neurotransmitter of the mammalian CNS (Fonnum, 1984). Maintenance of extracellular glutamate concentrations within a narrow physiological range involves control of its release as well as its uptake and is necessary to protect neurons from excitotoxic injury. We have previously shown that neuronal injury resulting from oxygen–glucose deprivation is exacerbated in the presence of NOS-2 induction and that this is accompanied by an increase in extracellular glutamate levels (Hewett et al., 1996). Here, we show that enhanced synaptic release

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of glutamate and not an alteration in its uptake or its release from extra-synaptic sources appears to underlie the potentiating effects of NOS-2 induction. It should be noted that the data interpretation in this murine neuronal/ astrocyte co-culture system is not complicated by the presence of NOS-1 (i.e., neuronal NOS) since we reported previously that these cultures express very little neuronal NOS (Hewett et al., 1993a). In addition, death that followed oxygen–glucose deprivation or direct NMDA exposure alone was not prevented by NOS inhibition, while the potentiated neuronal death that followed NOS-2 induction was indeed sensitive to inhibitors of NOS (Hewett et al., 1993a, 1996). Rapid buffering and subsequent uptake of synaptically released glutamate is achieved by glutamate transporters, thus maintaining a high signal to noise ratio (Diamond and Jahr, 1997; Maragakis and Rothstein, 2001; Nicholls and Attwell, 1990). Given the sensitivity of neurons to excitotoxicity, a defect in the system for clearing glutamate would be expected to have disastrous consequences due to a pathological rise in glutamate levels. In this study, we demonstrated no change in 3H-d-aspartate clearance under conditions of combined oxygen and glucose deprivation with or without induction of NOS-2. While most investigators agree that a lack of metabolites necessary for energy formation would eventually result in a diminution or loss of glutamate uptake, the results are mixed with respect to the time frame in which this occurs. For instance, Huang et al. (1993) measure a slight decrease following 30 min of oxygen–glucose deprivation, in pure astrocyte cultures, that progressively declines over the next 2 h. However, Stanimirovic et al. (1997), report a transient increase in uptake that is then followed by a decrease after 4 h of oxygen–glucose deprivation. Finally, others utilize chemicals to induce hypoxia and while this does result in a fairly rapid loss of glutamate uptake, it is difficult to directly compare the results obtained by essentially paralyzing a cellular system vs. allowing the natural progression of cellular events to occur after removal of energetic substrates (Jabaudon et al., 2000; Swanson, 1992). Thus, overall, it is likely that the lack of a measurable effect of oxygen–glucose deprivation alone on transporter function in our experimental paradigm is simply due to the short duration of the insult. In addition to potential alterations in uptake caused by oxygen–glucose deprivation alone, it should be noted that the function of the three major glutamate transporters, GLAST, GLT-1, and EAAC-1, can be alternately enhanced and inhibited by treatment with reducing and oxidizing reagents, respectively, indicating the presence of SH-based redox modulatory sites with the potential to be modulated by NO
or other RNOS (Trotti et al., 1997). In fact, there are reports that exposure to cytokines known to induce NOS-2 in astrocytes results in a suppression of glutamate transport, an effect blocked by NOS inhibition (Hu et al., 2000; Ye and Sontheimer, 1996). However, as mentioned, 3H-d-aspartate uptake was unaffected by NOS-2 induction in our exper-

imental system. This finding is in agreement with others demonstrating a lack of effect of RNOS Species (NO
and NO+) on astrocytic glutamate transport (Piani et al., 1993; Sorg et al., 1997; Volterra et al., 1994a). While glutamate uptake into synaptosomes was reported to be inhibited by NO
(Pogun et al., 1994), it is likely that this occurred secondary to inhibition of the H+-ATPase found in synaptic vesicles and not to a direct effect on the glutamate transporters themselves (Wolosker et al., 1996). Evidence does suggest that peroxynitrite (OONO ), formed by the near diffusion-limited reaction of NO
with superoxide (O
2 ) (Beckman et al., 1990; Blough and Zafiriou, 1985), can impair glutamate transporter function. However, we saw no evidence of inhibition when our cultures were exposed to SIN-1, a compound that we have determined to produce peroxynitrite under our experimental conditions (Trackey et al., 2001). While the reasons for the lack of effect of RNOS species on 3H-d-aspartate uptake in this study are not clear, one important difference between our study and others is that we used mixed cortical cell cultures and not just pure astrocyte cultures or reconstituted liposomes. The presence of neurons is known to alter patterns of expression and functional characteristics of glutamate transporters. Specifically, astrocytes cultured alone express only the GLAST transporter, while those cultured with neurons express both GLAST and GLT-1 (Swanson et al., 1997). Interestingly, numerous studies suggest a dominant role for GLT-1 in synaptic glutamate uptake (Raghavendra Rao et al., 2000; Rothstein et al., 1996; Tanaka et al., 1997). Thus, it is intriguing to speculate that GLAST is more susceptible to oxidative/nitrosative damage than GLT-1. Another possible reason could be the age of astrocytes in our co-cultures (5–6 weeks since initial plating). Indeed, Ye and Sontheimer (1996) observed in their study that the inhibitory effect of cytokines and RNOS species was not evident in older cultures. Whatever the reason, the lack of inhibitory effect on functional glutamate uptake indicates that glutamate transporter dysfunction was not a contributing factor to the increase in [glutamate]e levels found in this study after NOS-2 induction. Volume-sensitive organic anion channels (VSOACs) have been shown to act as portals for astrocytic glutamate efflux during spreading depression (Basarsky et al., 1999) and following exposure to hypotonic environment (Kimelberg et al., 1990) or high-potassium (Rutledge and Kimelberg, 1996). However, the exacerbation of oxygen– glucose deprivation-mediated neuronal injury following block of these channels in induced and control (i.e., oxygen-deprived only) cultures coupled with the initiation of astrocyte injury is inconsistent with this hypothesis. It should be noted that the concentration of NPPB used herein was not toxic to cultures maintained in an oxygen and glucose-containing environment. Thus, it would appear that VSOACs function in cell volume regulation/homeostasis plays a more important role in the recovery of cells rather than contributing to the injury process. Very recently, Ye et

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al. (2003) reported that glutamate could be released from astrocytes via astrocytic hemichannels and NPPB has also been shown to cause inhibition of these channels expressed in Xenopus oocytes (Eskandari et al., 2002). While we did not examine this mechanism directly, Ye et al. (2003) demonstrated that opening of hemichannels was accompanied by a greatly reduced glutamate uptake due to degradation of ionic gradients. Since there was no change in glutamate uptake into mixed cortical cell cultures in our experimental protocol, it seems unlikely that these channels contributed to the observed increase in [glutamate]e or to the neurotoxicity. Further, if this was to be the mechanism of action of NPPB in our experimental paradigm then NPPB should have had been protective rather than having an injury-enhancing effect as was demonstrated. Finally, prior in vitro and in vivo evidence indicates that NO donor compounds can increase neurotransmitter release (Kano et al., 1998; Lonart et al., 1992) perhaps via stimulating both Ca2+-dependent and -independent synaptic vesicle release (Meffert et al., 1994, 1996). Indeed, this process is hypothesized to be responsible for the initiation and maintenance of long-term potentiation (Arancio et al., 1996; O’Dell et al., 1991). While this is an important and necessary physiological event, it would appear that under pathological conditions, such as hypoxia–ischemia, astrocyte-derived NO could lead to an inappropriate enhancement of synaptic neurotransmitter release. Our data with TBOA, demonstrating significantly greater neuronal cell death in NOS-2-induced cultures along with the prevention of this toxicity with MK-801 and tetanus toxin, would suggest that this is indeed the case. It should be noted, however, that a recent report indicates that TBOA may be transported by astrocytic glutamate transporters, albeit slowly and after prolonged exposure, resulting in heteroexchange with glutamate (Anderson et al., 2001). This raises the possibility that the initial inciting event that follows TBOA exposure could be glutamate efflux due to heteroexchange causing further release of synaptic glutamate due to depolarization of neurons. Thus, tetanus toxin could be protective in this instance by preventing the secondary release of glutamate. Nevertheless, the ability of riluzole and tetanus toxin to prevent the NOS-2-mediated increased accumulation of [glutamate]e and inhibit the enhancement of oxygen–glucose deprivation-induced neuronal injury is consistent with the conclusion that increased synaptic release of glutamate is the predominant mechanism by which RNOS after NOS-2 induction potentiate neuronal injury. There has been a suggestion in the literature that in addition to it being a good sodium channel blocker (Benoit and Escande, 1991; Hebert et al., 1994), riluzole can inhibit NMDA receptors directly (Debono et al., 1993). We have confirmed in our hands that concentrations of riluzole from 0.1 to 1 mM do not prevent NMDA-induced neuronal injury directly. The ability of tetanus toxin to slightly but significantly protect against oxygen–glucose deprivationinduced neuronal injury has been reported previously

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(Monyer et al., 1992). However, in the study herein, the ability of tetanus toxin pretreatment to prevent the enhancement of neuronal injury that occurs after NOS-2 induction is much more dramatic. This may be because there is a tonic increase in synaptic glutamate release following NOS-2 induction. This is consistent with a study by Smith and Otis (2003) demonstrating that NO donors or NO generated from parallel fiber activity cause a long-term increase in the spontaneous firing rate of Purkinje neurons. However, since both astrocytes and neurons are capable of releasing glutamate in a Ca2+-dependent manner (Rubin, 1970; Vesce et al., 1999) and a recent study reported that NO caused a rapid, calcium-dependent release of glutamate from cultured rat astrocytes (Bal-Price et al., 2002), it could be argued that the source of glutamate is astrocytic and not neuronal. But, we used the holotoxin form of tetanus toxin, which cleaves the neuronal but not the astrocytic synaptobrevin-2 (Parpura et al., 1995), indicating that the glutamate is released solely from neurons. In summary, the present study demonstrates the enhanced astrocytic RNOS production can potentiate combined oxygen–glucose deprivation-induced neuronal injury, in part, by enhancing synaptic excitatory amino acid release. These results suggest that NO-mediated augmented excitatory dynamics during times when metabolic states are already compromised could contribute to the development and progression of cerebral ischemic neuronal injury.

Acknowledgments The authors wish to thank Ms. Tracy F. Uliasz for her excellent technical assistance. This work was supported by Grant NS36812 from NINDS, National Institutes of Health and by a grant from the American Heart Association. SJH is an Established Investigator of the American Heart Association.

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