Opposing roles for caspase and calpain death proteases in l -glutamate-induced oxidative neurotoxicity

Toxicology and Applied Pharmacology 232 (2008) 258–267

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Toxicology and Applied Pharmacology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / y t a a p

Opposing roles for caspase and calpain death proteases in L-glutamate-induced oxidative neurotoxicity Lucy M. Elphick a,1, Mohammad Hawat a, Nick J. Toms b, Annika Meinander c,d, Andrey Mikhailov c,d, John E. Eriksson c,d, George E.N. Kass a,⁎ a

Division of Biochemical Sciences, Faculty of Health and Medical Sciences, University of Surrey, Guildford, GU2 7XH, UK Peninsula Medical School, St Luke's Campus, Heavitree Road, Exeter, EX1 2LU, UK Turku Centre for Biotechnology, P.O.B. 123, FIN-20521 Turku, Finland d Department of Biochemistry and Pharmacy, Åbo Akademi University, FIN-20521 Turku, Finland b c

a r t i c l e

i n f o

Article history: Received 20 December 2007 Revised 30 June 2008 Accepted 1 July 2008 Available online 19 July 2008 Keywords: Apoptosis Necrosis Caspases Calcium Calpains Stroke Ischemic injury

a b s t r a c t Oxidative glutamate toxicity in HT22 murine hippocampal cells is a model for neuronal death by oxidative stress. We have investigated the role of proteases in HT22 cell oxidative glutamate toxicity. L-glutamateinduced toxicity was characterized by cell and nuclear shrinkage and chromatin condensation, yet occurred in the absence of either DNA fragmentation or mitochondrial cytochrome c release. Pretreatment with the selective caspase inhibitors either benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (pan-caspase), Nacetyl-Leu-Glu-His-Asp-aldehyde (caspase 9) or N-acetyl-Ile-Glu-Thr-Asp-aldehyde (caspase 8), significantly increased L-glutamate-induced cell death with a corresponding increase in observed nuclear shrinkage and chromatin condensation. This enhancement of glutamate toxicity correlated with an increase in L-glutamatedependent production of reactive oxygen species (ROS) as a result of caspase inhibition. Pretreating the cells with N-acetyl-L-cysteine prevented ROS production, cell shrinkage and cell death from L-glutamate as well as that associated with the presence of the pan-caspase inhibitor. In contrast, the caspase-3/-7 inhibitor Nacetyl-Asp-Glu-Val-Asp aldehyde was without significant effect. However, pretreating the cells with the calpain inhibitor N-acetyl-Leu-Leu-Nle-CHO, but not the cathepsin B inhibitor CA-074, prevented cell death. The cytotoxic role of calpains was confirmed further by: 1) cytotoxic dependency on intracellular Ca2+ increase, 2) increased cleavage of the calpain substrate Suc-Leu-Leu-Val-Tyr-AMC and 3) immunoblot detection of the calpain-selective 145 kDa α-fodrin cleavage fragment. We conclude that oxidative Lglutamate toxicity in HT22 cells is mediated via calpain activation, whereas inhibition of caspases-8 and -9 may exacerbate L-glutamate-induced oxidative neuronal damage through increased oxidative stress. © 2008 Elsevier Inc. All rights reserved.

Introduction Oxidative stress plays a major role in the induction of neuronal cell death in a number of disease states (Simonian and Coyle, 1996). The Lglutamate-induced model for oxidative stress in the murine hippocampal HT22 cell line has been shown to be an established model of oxidative neuronal cell death (Tan et al., 1998b; Satoh et al., 2000; Dargusch and Schubert, 2002; Rossler et al., 2004). In this model, increased levels of extracellular L-glutamate reduce cystine uptake by limiting or even reversing a cystine/L-glutamate exchanger, the socalled xc− transporter (Murphy et al., 1989). Under physiological conditions, this antiporter, which consists of the two subunits xCT and 4F2, mediates the Na+-independent transport of extracellular cystine into the cell, in exchange for intracellular L-glutamate (Bannai,

⁎ Corresponding author. Fax: +44 1483 686401. E-mail address: [email protected] (G.E.N. Kass). 1 Present address: Department of Cell and Molecular Biology, Imperial College London, South Kensington, London, SW7 2AZ, UK. 0041-008X/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.taap.2008.07.008

1986; Sato et al., 1999). Intracellular cystine is reduced to cysteine which may then be incorporated into the key intracellular antioxidant molecule, glutathione (Halliwell and Gutteridge, 1999). However following excessive L-glutamate exposure, the exchanger is either blocked or even reversed to deplete intracellular cysteine levels. The consequent reduction in intracellular glutathione levels may then render the cell susceptible to reactive oxygen species (ROS)-induced damage, ultimately leading to oxidative cell death (Tan et al., 1998a). The xc− transporter is specific for glutamate (Sato et al., 1999) and, at least in PC12 cells, oxidative glutamate toxicity was not mimicked by either aspartate or glycine (Schubert et al., 1992). Caspases represent a ubiquitous family of aspartate-specific cysteine proteases known to play a prominent role in apoptotic cell death (Riedl and Shi, 2004; Kumar, 2007). They are present in cells as inactive zymogens, organized in a hierarchical manner involving upstream initiator caspases and downstream execution caspases. Upon activation of plasma membrane death receptors (e.g. CD95) or following selective mitochondrial cytochrome c release, the apical caspases-8 and -9, respectively become activated and in turn pro-

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teolytically activate the execution caspases-3 and -7 (Riedl and Shi, 2004; Kumar, 2007). Previous studies employing caspase inhibitors have provided conflicting results on the possible caspase role in L-glutamate-induced oxidative neuronal cell death. For example, caspase inhibition was found to prevent L-glutamate-induced HT22 in several studies (Tan et al., 1998a; Tan et al., 1998b; Dargusch and Schubert, 2002; Stanciu and DeFranco, 2002) but this was not reproduced in other studies (van Leyen et al., 2005; Zhang and Bhavnani, 2006). Likewise, Ha and Park (2006) reported an increase in caspase-1 and caspases-3/-7 activity following treatment of HT22 cells with L-glutamate (Ha and Park, 2006) whereas no proteolytic caspase3 activation was detected in another study (van Leyen et al., 2005). Some of these discrepancies may be explained by experimental differences and biochemical endpoints used to assess cytotoxicity. Furthermore, caspases are not the exclusive protease family which may mediate cell death. Similar to caspases, calpains also require proteolytic activation and possess a similar target specificity, exemplified by αII-spectrin (fodrin), actin and poly(ADP-ribose)polymerase which are cleaved by calpains and caspase-3 (Wang, 2000; Friedrich, 2004). Calpain activation has been associated with many neurological disorders including excitotoxicity (Van den Bosch et al., 2002; Das et al., 2005; Takano et al., 2005), traumatic brain injury (Pike et al., 1998), spinal cord injury (Momeni and Kanje, 2005; Arataki et al., 2005) and ischemia (Rami, 2003; Cho et al., 2004; Kawamura et al., 2005). Indeed, their in vivo inhibition has been reported to produce successful cytoand functional protection (Markgraf et al., 1998; Arataki et al., 2005). In the case of L-glutamate-induced HT22 cell death, the role of calpains has also been controversial, with two studies (Tan et al., 1998b; van Leyen et al., 2005) reporting that calpain inhibitors had no effect on the cytotoxic effect of L-glutamate, whereas a third study reported a dependency of L-glutamate-induced cell death on calpain activity (Zhang and Bhavnani, 2006). In this study we have employed murine hippocampal HT22 cells to re-investigate the role of proteases in L-glutamate-induced oxidative toxicity. We report here that L-glutamate-induced oxidative cell death shares several morphological features of apoptosis, including cell and chromatin condensation. However, no execution caspase activation was observed and cell death occurred in the absence of cytochrome c release from mitochondria and nuclear DNA fragmentation. The mechanism of oxidative cell death was ROS- and Ca2+-dependent and was mediated by calpains. Unexpectedly, caspase inhibition exacerbated L-glutamate-induced oxidative HT22 cell death through enhanced ROS production. Materials and methods Chemicals. L -Glutamate, Triton X-100, dimethylsulfoxide, dithiothreitol, EGTA, propidium iodide, RNase A, N-acetyl-L-cysteine (NAC), cycloheximide and Hoechst 33258 were purchased from SigmaAldrich Ltd, (Poole, UK). Benzyloxycarbonyl-Val-Ala-Aspfluoromethylketone (Z-VAD-FMK), N-acetyl-Leu-Glu-His-Asp-aldehyde (Ac-LEHD-CHO), Ac-Ile-Glu-Thr-Asp-aldehyde (Ac-IETD-CHO), Nacetyl-Asp-Glu-Val-Asp aldehyde (Ac-DEVD-CHO) and Ac-Asp-GluVal-Asp-7-amido-4-trifluoromethylcoumarin (Ac-DEVD-AFC) were bought from Bachem (Bubendorf, Switzerland). [L-3-trans(propylcarbamoyl)oxirane-2-carbonyl]-L-isoleucyl-L-proline methyl ester (CA-074-Me) and N-acetyl-Leu-Leu-Nle-CHO (ALLN) were purchased from Calbiochem (Nottingham, UK). Suc-Leu-Leu-ValTyr-7-amino-4-methylcoumarin (Suc-LLVY-AMC) was purchased from Alexis Biochemicals, Birmingham UK. 5-(and -6)Chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate acetyl ester (CM-H2DCFDA), BAPTA-AM and all cell culture media and reagents were obtained from InVitrogen (Paisley, UK). FuGENE transfection reagent, lactate dehydrogenase (LDH) detection kit and bovine serum albumin were obtained from Roche (Lewes, UK). Monoclonal anticytochrome c antibody (clone 6H2.B4) and anti-CD95 antibody (Jo2)

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were purchased from BD Pharmingen (Oxford, UK). Anti-α-fodrin monoclonal antibody was obtained from Affiniti Research Products (Matford Court, UK) Secondary HRP- and FITC-conjugated antibodies were obtained from DAKO Ltd. (Cambridge, UK). Cell culture and apoptosis assays. HT22 cells were cultured in DMEM containing 10% FBS, 2 mM L-glutamine and 50 U/ml penicillin and 50 μg/ml streptomycin under a humidified atmosphere (5% CO2, 37 °C). Unless indicated otherwise, 8 × 103 cells were seeded into 24well plates and incubated overnight, prior to media renewal and treatment of the cells with the compounds of interest in DMEM containing 2% FBS. Where indicated, the buffering of the cytosolic Ca2+ levels was achieved by pre-incubating HT22 cells with 20 μM BAPTA-AM for 30 min prior to the treatment with L-glutamate. For the measurement of cell viability, aliquots of the media were removed at the indicated time points and assayed for lactate dehydrogenase (LDH) release, according to the manufacturer's instructions (Roche, Lewes, UK). Each sample of LDH was compared to the total, Triton X-100 releasable LDH from the same well at the end of each experiment. Morphological evidence for chromatin condensation was obtained by the DNA-specific dye Hoechst 33258 (2 μg/ml) as previously described (Jones et al., 1998). DNA fragmentation was measured by flow cytometric analysis of propidium iodide-stained cellular DNA as previously reported (Macanas-Pirard et al., 2005). In brief, DNA fragmentation was analyzed by flow cytometric detection of hypodiploid DNA. Cells were detached by trypsinization, combined with medium containing floating cells, and centrifuged at 100 g for 5 min. The pellets were fixed in ice-cold 70% (v/v) ethanol in PBS overnight at 4 °C by gradual addition while vortex mixing. The cells were subsequently stained with 10 μg/ml propidium iodide and treated with 1 mg/ml RNase for 30 min at 37 °C before analysis using a Beckman Coulter Epics XL flow cytometer (argon laser, excitation wavelength 488 nm). A minimum of 20,000 events were acquired in list mode while gating the forward and side scatters to exclude propidium iodide-positive cell debris and analyzed in FL-3 for the appearance of the sub-G1 peak. Caspase and calpain activity assays. In situ caspase-3 activity was analyzed using a previously described probe consisting of DsRed and EYFP linked by a 19 amino acid chain containing the caspase-3 cleavage site Asp-Glu-Val-Asp (called pFRET-casp-3) (Elphick et al., 2006). In brief, cells were transfected with pFRET-casp3 by using FuGENE 5 transfection reagent (Roche, Lewes, UK) according to manufacturer's instructions. Following a 72-hour incubation, transfected cells were then exposed to L-glutamate and confocal laser scanning microscopy images were subsequently recorded (Zeiss 510 META) using a 488 nm argon laser excitation, with the emission channel split at 545 nm enabling separation of EYFP (b545 nm) and DsRed (N545 nm) as described previously (Elphick et al., 2006). The proteolytic processing of pro-caspase-3 was assayed by flow cytometric detection of HT22 cells immunostained for active caspase-3 with a phycoerythrin-conjugated anti-caspase-3 antibody (1:20), according to the manufacturer's instructions (BD Pharmingen, Oxford, UK) (Macanas-Pirard et al., 2005). For the assessment of caspase-3/-7 (DEVDase) and calpain activities, HT22 cells were seeded in 175 cm2 flasks and treated with L-glutamate as described above. At the indicated time points, cells were harvested, resuspended in caspase lysis buffer (40 mM sucrose, 50 mM NaCl, 5 mM EGTA, 2 mM MgCl2, 10 mM HEPES, pH 7.0), freeze-thawed three times and the supernatant collected (15,000 ×g, 30 min). Eighty μg of sample protein was assayed for DEVDase activity with 40 μM Ac-DEVD-AFC (λex = 355 nm; λem = 480 nm) in caspase assay buffer (100 mM HEPES, 10% sucrose, 0.1% CHAPS, 10 mM dithiothreitol, pH 7.25) (Jones et al., 1998). Additionally, the cells were resuspended in calpain assay buffer (115 mM NaCl, 1 mM KH2PO4, 5 mM KCl, 2 mM CaCl2, 1.2 mM

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Fig. 2. L-Glutamate cytotoxicity is enhanced by Z-VAD-FMK and caspase-8 and caspase-9 inhibition, but not by caspase-3/-7 inhibition. Cells were pre-incubated for 30 min with either the pan-caspase inhibitor Z-VAD-FMK (50 μM) (panel A), or inhibitors of caspase-8 (Ac-IETD-CHO, 100 μM) (panel B), caspase-9 (Ac-LEHD-CHO, 100 μM) (panel C) and caspase-3 (Ac-DEVDCHO, 50 μM) (panel D). Cells were then exposed to L-glutamate (3 mM) and LDH release monitored over 24 h. The data represent the mean (±SEM) of four independent experiments. Comparisons of means were made using a one-way ANOVA followed by Bonferroni's post hoc test (⁎ = p b 0.05 and ⁎⁎⁎ = p b 0.001 versus L-glutamate treatment; ns = not significant).

MgSO4, 25 mM HEPES, pH 7.4, supplemented with 60 μM Suc-LLVYAMC) in black 96-well plates and assayed for calpain activity (λex = 355 nm; λem = 460 nm) in the absence or presence of 50 μM ALLN. Cytochrome c immunocytochemistry. Following fixation in 4% w/v paraformaldehyde, cells were washed in blocking buffer (PBS supplemented with 3% bovine serum albumin and 0.05% saponin, pH 7.2) and incubated overnight (4 °C) in 1:100 monoclonal anticytochrome c antibody (Macanas-Pirard et al., 2005). Following further washes, the cells were incubated in 100 μl of 1:100 diluted FITC-conjugated anti-mouse secondary antibody (1 h, room

temperature). After final washes, cells were then mounted in VectaShield prior to confocal laser scanning microscopic analysis as described above. Western blotting. Harvested cells were resuspended in lysis buffer (0.15 M NaCl, 50 mM Tris–Cl supplemented with 0.2% (w/v) SDS, 1% (v/v) Nonidet P40, 1 mM Na3VO4, 1 μg/ml leupeptin, 1 μg/ml aprotinin, 1 μg/ml pepstatin, pH 7.5) and centrifuged (15,000 ×g, 15 min, 4°°C) and the resultant supernatants were harvested. Protein concentrations were determined using the BioRad protein assay. Western blotting was then performed as described previously (El-Hassan et al., 2003).

Fig. 1. L-Glutamate induces cell shrinkage and chromatin condensation but not either DNA fragmentation or cytochrome c release. A. L-Glutamate induces a concentration-dependent cytotoxicity. Cells were treated with 0.1–100 mM L-glutamate over 24 h and LDH release monitored. Cell death is expressed as a percentage of total cellular LDH release, with each data point representing the mean (± SEM) of four independent experiments. B. Lack of DNA fragmentation in L-glutamate-induced cell death. Representative histograms of DNA content are shown for either 1 μM staurosporine-treated (18 h, dotted line) or 10 mM L-glutamate-treated (24 h, solid line) cells. Ethanol-fixed cells were stained with 20 μg/ml propidium iodide and then analyzed by flow cytometry as described under Materials and methods for the appearance of a sub-G1 peak as indicated by the bar in the histogram. C. LGlutamate induces cell shrinkage. Cells were exposed to either vehicle control (H2O) (i), 3 mM L-glutamate (ii), 50 μM Z-VAD-FMK (iii) or 50 μM Z-VAD-FMK + 3 mM L-glutamate (iv) for 24 h. Formaldehyde-fixed cells were then viewed using DIC. D. Quantitative analysis of L-glutamate-induced cell shrinkage. Cells were exposed to either vehicle control (H2O) (left panel) or 5 mM L-glutamate (right panel) for 18 h and analyzed by flow cytometry for forward and side scatter. E. L-Glutamate induces chromatin condensation. Cells were exposed to either vehicle control (H2O) (i), 3 mM L-glutamate (ii), 50 μM Z-VAD-FMK (iii) or 50 μM Z-VAD-FMK + 3 mM L-glutamate (iv) for 18 h. Formaldehyde-fixed cells were then stained with Hoechst 33258. Cells displaying nuclear chromatin condensation are indicated by arrows. Magnification, 400×. F. Lack of mitochondrial cytochrome c translocation in L-glutamateinduced toxicity. Cells were treated with either vehicle control (H2O) (i), 3 mM L-glutamate (ii), 50 μM Z-VAD-FMK (iii), 50 μM Z-VAD-FMK + 3 mM L-glutamate (iv) for 18 h or antiCD95 monoclonal antibody (Jo2, 200 ng/ml) plus cycloheximide (1 μg/ml) for 27 h (v). Following fixation, cells were immunostained for cytochrome c and analyzed via confocal laser scanning microscopy. Scale bar = 50 (C) or 20 (F) μm.

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ROS measurements. HT22 cells were seeded in 25 cm2 flasks and treated with L-glutamate in the presence of the caspase inhibitor ZVAD-FMK or a corresponding volume of DMSO as described above. At the indicated time points, cells were harvested, resuspended in 1 ml Dulbecco's PBS supplemented with 10 μM CM-H2DCFDA and incubated for 30 min at 37°°C. After washing, cells were analyzed using a Beckman Coulter Epics XL flow cytometer (Beckman Coulter, Fullerton, CA) (argon laser, excitation wavelength 488 nm). A minimum of 10,000 events were acquired in list mode while gating the forward and side scatters to exclude cell debris and analyzed in FL-1. Statistical analysis. All data are expressed as means (±SEM) of at least three independent experiments. Comparisons of treatments against controls were made using one-way ANOVA followed by Bonferroni's post hoc test with GraphPad Prism (v. 4) software. Significance difference was set at p b 0.05.

Results L-glutamate induces chromatin condensation and cell death without cytochrome c release

Initial experiments confirmed a previous report that a 24-hour exposure to 3–10 mM L-glutamate induced marked HT22 cytotoxicity (Davis and Maher, 1994) (Fig. 1A). Morphological investigation of the cells exposed to L-glutamate revealed cell shrinkage (Fig. 1C). This was confirmed by flow cytometric analysis as evidenced by a decrease in Forward Scatter (Fig. 1D). L-glutamate also induced marked chromatin condensation (Fig. 1E). Both cell shrinkage and chromatin condensation are signs of apoptosis; however, in contrast to classic apoptosis, nuclear and DNA fragmentation were both absent (Figs. 1B and E). Moreover, no mitochondrial cytochrome c release occurred during L-glutamate-induced cytotoxicity, evidenced by retained punctate

Fig. 3. L-Glutamate-induced ROS production is enhanced by Z-VAD-FMK. (A) Cells were exposed to L-glutamate (3 mM) and loaded with CM-H2DCFDA at the indicated time points followed by DCF fluorescence measurement as described under Materials and methods. (B) Cells were pre-incubated for 30 min with either the pan-caspase inhibitor Z-VAD-FMK (50 μM), NAC (1 mM) (alone or in combination) or a corresponding volume of carrier solvent (DMSO) before exposure to L-glutamate (3 mM or 5 mM) for 6 h and measurement of DCF formation by flow cytometry. (C) The data represent the mean (±SEM) of four independent experiments. (D and E) NAC prevents cell shrinkage. Cells were exposed to either vehicle control (H2O) (left panel) or 3 mM L-glutamate (right panel) for 16 h in the absence or presence of Z-VAD-FMK (50 μM) or NAC (1 mM) (alone or in combination) or a corresponding volume of carrier solvent (DMSO) and analyzed by flow cytometry for forward and side scatter. In E, the histograms were gated for high forward scatter (healthy cells) and low forward scatter (shrunken cells). Each bar represents the mean (±SEM) of three independent experiments. The cells exposed to Z-VAD-FMK or NAC alone were not significantly different from untreated controls. (F) NAC prevents L-glutamate-induced cell death. Cell death is expressed as a percentage of total cellular LDH release, with each bar representing the mean (± SEM) of four independent experiments. Comparisons of means were made using a one-way ANOVA followed by Bonferroni's post hoc test (⁎ = p b 0.05).

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(mitochondrial) and absence of nuclear cytochrome c staining throughout the cytotoxic insult (Fig. 1F). This is clearly different from the distinct relocalization of cytochrome c with loss of punctate distribution and marked nuclear staining observed following Fas (CD95) activation in HT22 cells (Fig. 1F). Caspase inhibition enhances L-glutamate-induced cytotoxicity through increased ROS production As expected, induction of HT22 cell apoptosis by a known apoptotic trigger (1 μM staurosporine) was significantly reduced by the pan-caspase inhibitor Z-VAD-FMK (50 μM) (data not shown). Surprisingly, Z-VAD-FMK induced a marked enhancement of 3 mM L-glutamate-induced cytotoxicity (Fig. 2A). Alone, Z-VAD-FMK failed to induce significant chromatin condensation or cytochrome c release. However, the combination of Z-VAD-FMK plus L-glutamate resulted in an increase in the number of shrunken cells (Fig. 1C) and cells possessing chromatin condensation (Fig. 1D) that paralleled the increase in cytotoxicity (Fig. 2A). As with L-glutamate alone, the combination of L-glutamate plus Z-VAD-FMK also failed to cause cytochrome c release (Fig. 1E). In order to identify the caspases responsible for this enhancement of L-glutamate-induced cell death, HT22 cells were pretreated with a range of caspase inhibitors. Inhibition of the execution caspases-3 and -7 by pretreating the cells with Ac-DEVD-CHO (50 μM) for 30 min failed to significant influence 3 mM L-glutamate-induced cytotoxicity. In contrast, both the caspase-9 inhibitor Ac-LEHD-CHO (100 μM) and the caspase-8 inhibitor Ac-IETD-CHO (100 μM) significantly enhanced L-glutamate-induced cell death (Fig. 2). Given the critical role of ROS in initiating the cascade of events leading to the death of HT22 cells exposed to glutamate (Tan et al., 1998a), we tested the hypothesis that caspase inhibition caused an exacerbation of ROS production. In agreement with earlier studies (Tan et al., 1998a; Noh et al., 2006; Ha and Park, 2006), 3 mM L-glutamate induced a time-dependent increase in ROS production as evidenced by the increased number of cells which had converted CM-H2DCFDA to its oxidized fluorescent DCF product (Fig. 3A). Incubating HT22 with the pan-caspase inhibitor Z-VAD-FMK for over 6 h did not lead to any detectable change in CM-H2DCFDA oxidation, showing that caspase inhibition by itself did not cause any

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ROS formation (Figs. 3B and C). In clear contrast, the combination of 3 mM L-glutamate with the caspase inhibitor resulted in an increase in ROS formation that was significantly higher than that reached with 3 mM L-glutamate alone and reached levels similar to those obtained with 5 mM L-glutamate (Fig. 3C). The increase in ROS formation following caspase inhibition manifested itself not only by an increase in the population of the cells showing enhanced levels of DCF formation (Fig. 3C) but also in the intracellular levels of DCF in that population (Fig. 3B). Even though the lack of linearity of the response of H2DCF based probes to ROS precludes quantification of intracellular ROS levels, our findings nevertheless strongly suggest that the pan-caspase inhibitor Z-VAD-FMK increased the rate of glutamateinduced ROS formation. A 30-min pretreatment of HT22 cells with the antioxidant NAC (1 mM) completely prevented the increase in ROS formation caused by L-glutamate or the combination of L-glutamate and Z-VAD-FMK (Fig. 3B). To investigate the relationship between caspase inhibition, ROS formation and cell death, the effect of NAC on cell morphology and viability was also examined. Figs. 3D and E show that L-glutamate-induced cell shrinkage was completely blocked by NAC. Likewise, the enhanced cell shrinkage caused by L-glutamate under conditions of caspase inhibition was equally well prevented by NAC. The flow cytometric data were confirmed by microscopic examination of the cells and for each treatment, NAC prevented the morphological changes induced by L-glutamate and the combination of L-glutamate and Z-VAD-FMK (data not shown). Further investigation of the protective effects of NAC revealed that the antioxidant fully protected the cells from the loss of cell viability following exposure to L-glutamate or the combination of L-glutamate and Z-VAD-FMK (Fig. 3F). Lack of execution caspase activation in oxidative L-glutamate toxicity In agreement with the lack of cytoprotection by caspase inhibition, no significant increased caspase-3 activity was detected during L-glutamate insults. This was evidenced by no significant increase in DEVDase activity following a 24 h treatment period with 3 mM L-glutamate (Fig. 4A) and the absence of proteolytic processing of procaspase-3, assayed by the appearance of immunoreactive caspase-3 p20/p17 subunit using flow cytometry (data not shown). In addition,

Fig. 4. L-Glutamate cytotoxicity does not involve caspase-3 activation. (A) A representative experiment showing caspase-3 activity measured in lysates from cells treated for 18 h either with vehicle control (♦), 3 mM L-glutamate (▾) or 1 μM staurosporine (□) by monitoring the release of AFC from DEVD-AFC. (B) Cells, transfected with pFRET-casp-3, were exposed to either vehicle control (H2O) (i), 10 mM L-glutamate (ii), vehicle control (0.1% DMSO) (iii) or 1 μM staurosporine (iv) for 18 h and examined by confocal laser scanning microscopy. Caspase-3-mediated cleavage of the reporter construct is evidenced by Fluorescence Resonance Energy Transfer (FRET) loss (EYFP-DsRed), with a resultant loss of DsRed fluorescence (red) and concomitant increase in EYFP fluorescence (green). Scale bar = 20 μm.

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the in situ cleavage of caspase-3 substrates was investigated using pFRET-casp3. Control HT22 cells expressing this FRET construct showed enhanced red fluorescence as a result of energy transfer from EYFP to DsRed due to the caspase-3 containing linker region between the two fluorescent proteins being intact (Fig. 4B). The positive inducer of apoptosis and caspase-3 activator staurosporine caused a reduction of FRET from donor EYFP to acceptor DsRed as a result of cleavage of the linker region. This was manifested by an increase in EYFP emission and a corresponding decrease in DsRed fluorescence. In clear contrast, 10 mM L-glutamate-treated HT22 cells showed no loss of FRET demonstrating a lack of in situ caspase-3 activation induced by L-glutamate. The involvement of other proteases in oxidative L-glutamate toxicity The above results have clearly shown that execution caspases are neither activated nor mediate the cytotoxicity of L-glutamate. Instead, the initiation caspases-8 and -9 appear to play a cytoprotective role against L-glutamate cytotoxicity. To investigate whether other proteases that are unrelated to caspases are involved in the mechanism of cell death, we investigated the possible role of cathepsins and calpains in L-glutamate oxidative toxicity. The cathepsin B inhibitor CA-074-Me afforded weak inhibition versus 3 mM L-glutamate-induced injury and was without effect in protecting against 10 mM L-glutamate cytotoxicity (Fig. 5A). In contrast, the calpain inhibitor ALLN (10 μM)

provided near-complete cytoprotection (Fig. 5B). Since calpains require a rise in intracellular Ca2+ levels for their activation, the role of Ca2+ in L-glutamate cytotoxicity was investigated. As shown in Fig. 5C, the buffering of intracellular Ca2+ with BAPTA by pre-loading the cells with 20 μM BAPTA-AM for 30 min significantly suppressed L-glutamate-induced cell death. BAPTA itself induced some cytotoxicity; however, this was significantly less than the toxicity caused by L-glutamate. Importantly, L-glutamate did not further enhance the toxicity of BAPTA. A sustained increase in cytosolic Ca2+ concentration that leads to cytotoxicity often necessitates influx of Ca2+ across the plasma membrane to replenish depleted endoplasmic reticulum Ca2+ stores. To test the requirement for extracellular Ca2+ in the cytotoxicity of L-glutamate, we buffered extracellular Ca2+ by adding 1.9 mM EGTA to the medium. Fig. 5D shows that in the absence of extracellular Ca2+, L-glutamate-induced HT22 cell death was significantly suppressed. The distinct pattern of fragmentation of the cytoskeletal protein αfodrin (αII-spectrin) is an established marker to discriminate between caspase-3- and calpain-mediated in situ proteolysis and, therefore, can be used as an assay for calpain activity in intact cells (Dutta et al., 2002). Western blot analysis revealed that in L-glutamate-treated cells, fodrin was cleaved to yield a 145 kDa fragment that represents calpain-mediated cleavage of the protein (Fig. 6A). In agreement with other findings, the caspase-3 signature cleavage product that corresponds to a 120 kDa fragment was absent, unlike the positive apoptotic control cells exposed to 1 μM staurosporine. Finally, the

Fig. 5. L-Glutamate-induced cytotoxicity is inhibited by ALLN and extracellular Ca2+ chelation, but not via cathepsin B inhibition. Cells were exposed to either L-glutamate for 24 h in the absence or presence of 10 μM CA-074, a cathepsin B inhibitor (A), 10 μM ALLN, a calpain inhibitor (B), cytosolic Ca2+ buffering with BAPTA (C) or extracellular Ca2+ chelation with 1.9 mM EGTA (D) for 24 h. Data represents the mean (±SEM) of four independent experiments. Comparisons of means were made using a one-way ANOVA followed by Bonferroni's post hoc test (⁎ = p b 0.05; ⁎⁎ = p b 0.01; ⁎⁎⁎ = p b 0.001 versus L-glutamate treatment; ns = not significant).

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Fig. 6. L-Glutamate induces calpain activation. (A) Cells were treated with 10 mM L-glutamate (24 h), 1 μM staurosporine (STS, 18 h) or their corresponding vehicles (H2O, lane 3; 0.1% DMSO, lane 2). Total cell lysate protein was separated by 6% SDS-PAGE prior to α-fodrin cleavage product immunoblotting. Arrows indicate naïve α-fodrin substrate and corresponding 145 kDa- and 120 kDa-cleavage fragments. Densitometric analysis of the 145 kDa band revealed a greater than 5.4 fold increase over vehicle-treated controls. (B) Cells were treated with either H2O vehicle (▴) or 3 mM L-glutamate ( ) for 6 h and then incubated with the calpain substrate Suc-LLVY-AMC. A representative experiment depicting changes in fluorescence (liberated AMC) over a 2-hour period is shown.

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presence of active calpain in cell lysates following L-glutamate exposure was confirmed by an increase in the rate of cleavage of the calpain substrate Suc-LLVY-AMC (Fig. 6B). Discussion Exposure of HT22 cells to high levels of L-glutamate results in oxidative cell death via blockade/reversal of the xc− transporter and subsequent intracellular glutathione depletion (Tan et al., 1998b; Satoh et al., 2000; Dargusch and Schubert, 2002; Rossler et al., 2004). This study has used this model of oxidative cell death to study the role of proteases in oxidative neuronal cell death. HT22 cells exposed to a cytotoxic concentration of L-glutamate were characterized by distinct chromatin condensation, but without DNA fragmentation or apoptotic body formation. Therefore, the nuclear morphological biochemical features induced by L-glutamate in HT22 cells only incompletely reflected the classic apoptotic morphological features of chromatin condensation, chromatin margination to the nuclear envelope and nuclear fragmentation to form apoptotic bodies. A similar form of chromatin condensation has recently been observed in cerebellar granule neurons exposed to L-glutamate (Bezvenyuk et al., 2003). The most likely reason for the incomplete apoptosis is the lack of execution caspase activation. The late apoptotic stages of the nuclear changes require caspase-6-mediated lamin A cleavage (Ruchaud et al., 2002) as well as the presence of ATP (Kass et al., 1996). Indeed, when execution caspases are not activated (El-Hassan et al., 2003; Bezvenyuk et al., 2003), are inhibited (Johnson et al., 2000) or are not expressed (Zheng et al., 1998), chromatin condensation occurs in cells undergoing

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apoptosis but in the absence of nuclear fragmentation and apoptotic body formation. A potential candidate that could mediate chromatin condensation in the absence of caspases is apoptosis-inducing factor (AIF) (Susin et al., 1999). However, its ability to translocate to the nucleus and cause chromatin condensation is restricted by its mitochondrial localization. The data presented here show that cytochrome c was not released from mitochondria during L-glutamate-induced cell death. Therefore, AIF is unlikely to play a role in oxidative L-glutamate-induced nuclear changes because of the absence of increased outer mitochondrial membrane permeability which is regarded as a prerequisite for the release of AIF from mitochondria (Arnoult et al., 2002). In contrast, another study (Andreau et al., 2004) has reported preapoptotic chromatin condensation, induced by staurosporine in HeLa cells that preceded both cytochrome c and AIF release and caspase activation. Whether this preapoptotic, caspase- and AIF-independent chromatin condensation is identical to the L-glutamate-induced chromatin condensation observed here remains to be determined, and is a focus of our ongoing investigations. An earlier study (Stanciu and DeFranco, 2002) had reported an inhibition of oxidative HT22 cell death by the pan-caspase inhibitor ZVAD-FMK. Likewise, the caspase inhibitor Ac-YVAD-CMK was found to prevent L-glutamate-induced HT22 cell death in several studies (Tan et al., 1998b; Dargusch and Schubert, 2002). However, we and others (van Leyen et al., 2005; Zhang and Bhavnani, 2006) were unable to demonstrate any protection under similar experimental conditions. A likely reason for this apparent discrepancy is that instead of assaying cell viability using 3-(4,5-dimethyldiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) reduction, we quantified cell death directly by assaying lactate dehydrogenase release from compromised cells. Previous studies comparing the two methods in neuronal cells revealed that MTT reduction provides an inaccurate measurement of cell death (Lobner, 2000; Zhao et al., 2002). Indeed, MTT reduction is dependent on cellular NAD(P)H content and mitochondrial and nonmitochondrial dehydrogenase activities and therefore changes in MTT reduction may not reflect cell death (Dhanjal and Fry, 1997). In contrast, LDH release provides robust evidence for plasma membrane damage in the absence of interference by the cell's metabolic status. We found that Z-VAD-FMK significantly enhanced L-glutamateinduced cell death, an effect that was, at least partly, reproduced by inhibiting the initiation caspases-8 and -9. The ability of Z-VAD-FMK to enhance cell death has previously been observed in tumor necrosis factor- and ROS-mediated cell death models (Vercammen et al., 1998; Kim and Han, 2001; Cauwels et al., 2003; Liu et al., 2003; Prabhakaran et al., 2004; Martinet et al., 2006; Uchiyama et al., 2007; May and Madge, 2007). It is noteworthy that in these studies, caspase inhibition led to a switch from apoptosis to necrosis. Cells like L929 mouse fibrosarcoma cells (Yu et al., 2006) and murine embryonic fibroblasts (May and Madge, 2007) respond to Z-VAD-FMK with enhanced ROS formation. In L929 cells this was attributed to catalase degradation with downstream activation of autophagic cell death (Yu et al., 2006). The consequences of caspase inhibition are very much different from what is reported here. Z-VAD-FMK did not cause an increase in ROS formation in the absence of L-glutamate and did not induce a switch in the type of cell death. Instead, the enhanced execution of cell death was along the same pathway triggered by L-glutamate with no apparent morphological or biochemical difference. Recently, it has been reported that the inhibition of caspase-9 in lymphoblastic cells, whether by LEHD-FMK or by RNA interference-mediated caspase-9 downregulation, lead to an increase in apoptosis (rather than a switch to necrosis) in response to trophic signal removal and staurosporine (Shah et al., 2004). Likewise, caspase-8 activation is now recognized to play an important role in B-cell survival (Dohrman et al., 2005; Lemmers et al., 2007). Therefore, an additional role for caspases (and in particular initiation caspases) appears to exist which counteracts HT22 cell death. The molecular mechanism underlying this caspase-

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mediated effect appeared to be linked to the level of ROS production. As reported here, even though Z-VAD-FMK by itself had no effect on basal ROS production, when combined with L-glutamate, the levels of ROS produced were approximately doubled. Therefore, L-glutamateinduced toxicity involves increased cytotoxic ROS production that is accelerated by caspase inhibition. We propose that the initiation caspases limit ROS production by L-glutamate but the target protein involved remains to be identified. It has been suggested that caspasemediated cleavage of ROS-generating proteins such as receptor interacting protein (RIP) and phospholipase A2 may constitute a potential mechanism (Cauwels et al., 2003; Chipuk and Green, 2005). The cathepsin B inhibitor CA-074-Me provided only a very small inhibition from 3 mM L-glutamate-induced injury and was without effect in protecting against the cytotoxicity of 10 mM L-glutamate. This suggests that cathepsins are not the major executioner protease in this model of oxidative toxicity. Instead, a critical role of calpains in the mechanism of L-glutamate-induced oxidative injury has been evidenced by the inhibition of cell death by the calpain inhibitor ALLN. Calpains were activated 6 h following L-glutamate exposure, and this required an increase in cytosolic Ca2+ concentration which was dependent on extracellular Ca2+ influx. An increase in Ca2+ levels in HT22 cells exposed to L-glutamate had been previously documented (Tan et al., 1998a; Herrera et al., 2007). Previous studies have shown that calpains are involved in a complex cross-talk with caspases. Calpains are known to cleave Bax and to generate a fragment with enhanced pro-apoptotic activity that ultimately causes the release of cytochrome c from mitochondria and induction of apoptosis (Wood et al., 1998; Toyota et al., 2003). Yet, calpains also suppress caspase-dependent apoptosis by proteolytically degrading caspase-9 and caspase-3 (Lankiewicz et al., 2000; Bizat et al., 2003). In contrast, in our L-glutamate-induced oxidative toxicity model the absence of cytochrome c release supports a model where calpain independently induces cell death in the absence of caspase involvement. In summary, this report provides evidence that calpains mediate L-glutamate-induced oxidative toxicity in the murine HT22 neuronal cell line to induce a caspase-independent form of apoptosis. Furthermore, a novel cell protection role by caspases that operates by suppressing ROS formation was identified. These findings may have significant implications when considering treatment avenues for oxidative neurodegenerative diseases. Acknowledgments We thank Dr Pamela Maher (Scripps Research Institute, California) for donating the HT22 cell line and to Dr Alessandro Natoni (MRC Toxicology Unit, Leicester, UK) and Dr Afshin Samali (National University of Ireland, Galway, Ireland) for their help and advice. This work was supported by TEKES (Finland). References Andreau, K., Castedo, M., Perfettini, J.L., Roumier, T., Pichart, E., Souquere, S., Vivet, S., Larochette, N., Kroemer, G., 2004. Preapoptotic chromatin condensation upstream of the mitochondrial checkpoint. J. Biol. Chem. 279, 55937–55945. Arataki, S., Tomizawa, K., Moriwaki, A., Nishida, K., Matsushita, M., Ozaki, T., Kunisada, T., Yoshida, A., Inoue, H., Matsui, H., 2005. Calpain inhibitors prevent neuronal cell death and ameliorate motor disturbances after compression-induced spinal cord injury in rats. J. Neurotrauma 22, 398–406. Arnoult, D., Parone, P., Martinou, J.C., Antonsson, B., Estaquier, J., Ameisen, J.C., 2002. Mitochondrial release of apoptosis-inducing factor occurs downstream of cytochrome c release in response to several proapoptotic stimuli. J. Cell Biol.. 159, 923–929. Bannai, S., 1986. Exchange of cystine and glutamate across plasma membrane of human fibroblasts. J. Biol. Chem. 261, 2256–2263. Bezvenyuk, Z., Miettinen, R., Solovyan, V., 2003. Chromatin condensation during glutamate-induced excitotoxicity of celebellar granule neurons precedes disintegration of nuclear DNA into high molecular weight DNA fragments. Mol. Brain Res. 110, 140–146. Bizat, N., Hermel, J.M., Humbert, S., Jacquard, C., Creminon, C., Escartin, C., Saudou, F.,

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