Nitric-oxide-induced depolarization of neuronal mitochondria: implications for neuronal cell death

Molecular and Cellular Neuroscience 24 (2003) 1151–1169

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Nitric-oxide-induced depolarization of neuronal mitochondria: implications for neuronal cell death Nina J. Solenski,a,* Vannessa K. Kostecki,a Serena Dovey,b and Ammasi Periasamyc a

Department of Neurology, University of Virginia Health Sciences System, Charlottesville, VA 22908, USA b School of Medicine, University of Virginia Health Sciences System, Charlottesville, VA 22908, USA c Biological Sciences and Biomechanical Engineering, University of Virginia Health Sciences System, Charlottesville, VA 22908, USA Received 2 July 2003; revised 8 August 2003; accepted 21 August 2003

Abstract Nitric oxide (NO•) has known toxic effects on central nervous system neurons. This study characterized the effect of NO• on mitochondrial membrane changes by exploring the relationship among NO•, excitatory receptor activation, and the induction of peroxynitrite, a highly toxic NO• reactant, to neuronal injury. Cultured rat cortical neurons were exposed to the NO• generator, diethylenetriamine/ nitric oxide adduct, and were examined for signs of cell death, mitochondrial membrane potential changes (⌬␺m), and the induction of a mitochondrial permeability transition (MPT). Neurons were also examined for nitrotyrosine (NT) immunoreactivity, a marker of reactive nitrogen species (RNS) formation. Neurons exposed to NO• or to N-methyl-D-aspartate (NMDA) exhibited similar rapid depolarization of mitochondria, which was prevented by an NMDA receptor antagonist. Electrophysiological studies demonstrated NO• potentiation of NMDA-induced NMDA receptor currents. NO• and NMDA-treated neurons had evidence of mitochondrial-specific NT immunoreactivity that was prevented by a SOD/catalase mimetic (EUK-134). EUK-134 treatment reduced both NO• and NMDA-induced NT formation and neuronal cell death. EUK-134 did not prevent NO-induced ⌬␺m but partially prevented NMDA-induced ⌬␺m loss. Although NO• and NMDA both induced MPT and MPT inhibitors prevented NO-induced ⌬␺m, they did not result in significant neuroprotection, in contrast to treatment designed to decrease peroxynitrite formation. These data suggest that NO-induced NMDA receptor activation is closely linked to intramitochondrial NO-peroxynitrite/RNS formation and thereby acts as a major mediator of neuronal cell death. © 2003 Elsevier Inc. All rights reserved.

Introduction The neurotoxic effects of the diffusible gas, nitric oxide (NO•), are extensive (Dawson et al., 1993; Palluy and Riguad, 1996; Lipton et al., 1993; Lipton, 1999). As a toxin, NO• is a well-established inhibitor of the mitochondrial electron transport system (ETS) (Brorson and Zhang, 1997; Brorson et al., 1999), causing mitochondrial adenosine triphosphate (ATP) depletion and loss of mitochondrial respiration. The latter has only recently been directly linked to apoptotic cell death (Almeida et al., 2001). NO• can also form a variety of toxic reactants, including peroxynitrite

* Corresponding author. Department of Neurology, Box 800394, University of Virginia Health Sciences System, Charlottesville, VA 22908, USA. Fax: ⫹1-434-982-1726. E-mail address: [email protected] (N.J. Solenski). 1044-7431/$ – see front matter © 2003 Elsevier Inc. All rights reserved. doi:10.1016/j.mcn.2003.08.011

anion, ONOO-, a product of NO• and superoxide anion radical, O•⫺ 2 . Superoxide anion radical is a normal byproduct of the mitochondrial ETS and is normally harmlessly detoxified by manganese superoxide dismutase (MnSOD) enzyme systems (for review see MacmillanCrow and Cruthirds, 2001); however, if this system fails in the presence of NO•, O•⫺ 2 rapidly forms peroxynitrite. As a potent reactive oxidative species, peroxynitrite can modulate oxidative signaling and induce damage through the modification of nucleic acids, lipids, and proteins. Peroxynitrite formation has been implicated with neural ischemic damage (Eliasson et al., 1999) in causing evidence that it can react with the mitochondrial ETS and with constituents of the mitochondrial transition pore with the induction of a mitochondrial permeability transition (MPT) (Cassina and Radi, 1996; Radi et al., 1994; Aulak et al., 2001; Brookes et al., 2000; Vieira et al., 2001). Peroxynitrite can also nitrate

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a variety of proteins involved in cell death and survival including MnSOD, cytochrome-c (MacMillan-Crow and Thompson, 1999; Reynolds, 1999), and caspase-3. Thus, the nitrosylating effect of peroxynitrite and other reactive nitrogen species may have a profound effect on protein function, with subsequent impact on cell viability (Leist et al., 1997), and is currently an area of intense research. Intriguingly, new data indicate that neuronal mitochondria produce in situ NO• with unclear functional significance. These new data make NO• an attractive candidate for mediating in situ mitochondrial damage. Prior to this discovery it was believed that pre- and postsynaptic locations of nNOS represented main sources for neuronal NO•-related toxicity. Much less is known regarding how or whether mitochondrial in situ formation of NO• and/or peroxynitrite formation plays a direct role in mediating the observed mitochondrial membrane changes or whether it acts as an essential cell signal to mediate cell death or survival mechanisms (Radi et al., 2002). Recently it has been suggested that mitochondria contain nitric oxide synthase (mtNOS) in an isoform similar to constitutive neuronal NOS (Ghafourifar and Ritcher, 1997; Kanai et al., 2001; Elfering et al. 2002). Both enzymes are calcium-activated and are upregulated during brain injury (Lacza et al., 2001). A plausible, but unproven hypothesis is that endogenous mtNOS can be activated during conditions of oxidative stress and locally produce NO• or peroxynitrite anion in sufficient amounts to damage the ETS or to change mitochondrial permeability to trigger secondary cell death signals. It is also known that mitochondria play a critical role in determining the mode of excitotoxic cell death in neurons (Ankarcrona et al., 1995). N-methyl-D-aspartate (NMDA)mediated mitochondrial depolarization has also been extensively reported (White and Reynolds, 1996; Schinder et al., 1996) and may be a consequence of NMDA receptor activated rise in calcium (Ca2⫹) levels within mitochondria as they perform their important role of buffering Ca2⫹ (Peng and Greenamyre, 1998; Peng et al., 1998; Castilho et al., 1999). It has also long been observed that inhibiting neuronal NOS can prevent excitotoxic-induced neuronal injury (Dawson et al., 1991, 1996; Hoyt et al., 1992; Chang et al., 2000). Recent data suggest that at least in hippocampal neurons both Ca2⫹ and NO• are required for excitotoxicinduced mitochondrial depolarization (Keelan et al., 1999). These important data suggest that NO• is an important feature of the excitotoxic-mediated mitochondrial injury. Integrating these seemingly diverse observations into a unifying mechanism of neurotoxicity is challenging but now seems possible in light of new data on the biochemistry of mitochondria and the confirmation of the presence of mtNOS. NMDA receptor activation with increased intracellular Ca2⫹ level may trigger mitochondrial Ca2⫹ uptake activating mtNOS to produce NO• and/or generate peroxynitrite within mitochondria. Thus, the present emerging data suggest that NO• and peroxynitrite are potentially critical links among NMDA receptor activation, Ca2⫹ dys-

regulation, and neurotoxicity as regulated in part by the mitochondria (Almeida et al., 1999). One aim of this study is to demonstrate and characterize this critical link by identifying whether in situ peroxynitrite formation is related to either NMDA- or NO•-mediated mitochondrial dysfunction and if so to better define its role in neuronal injury. Parts of this paper have been published previously in abstract form (Solenski et al., 2002).

Results NO• induces neuronal apoptotic and necrotic death in a time- and concentration-dependent manner and caspase-3 and cytochrome-c release In order to better characterize the role of NO• in neuronal death we first needed to confirm that NO•-mediated death occurred in our cell culture system. Primary cortical neurons (14 –17 days in vitro [DIV]) were exposed to diethylenetriamine/nitric oxide adduct (DETA-NO•) and reproducibly underwent both apoptotic and necrotic forms of cell death with necrosis predominating at the higher concentrations of DETA-NO•. Fig. 1 illustrates that as DETA-NO• concentration increased there was loss of diffuse cytosolic calcein-AM staining (green) and an increase in propidium iodide (PI) staining (red), indicating cell death. Residual calcein AM stain of unclear localization is present, perhaps marking calcium ion, but is not indicative of viability. Tetramethylrhodamine methyl ester (TMRM) labeling of mitochondria was also diminished, indicating mitochondrial injury and depolarization; at 1000 ␮M DETA-NO• there was evidence of severe membrane lysis consistent with necrosis and near absence of TMRM staining. By 24 h, an average of 71–90% of neurons demonstrated these findings when exposed to 3 h of 1000 ␮M DETA-NO•, compared to control nontreated neurons. When a total initial cell count was performed prior to NO• treatment and compared to total posttreatment cell counts, there was an additional estimated 15–20% total cells “unaccounted for,” likely due to severe membrane lysis and cellular degradation. In order to equate DETA-NO• concentration to NO• concentrations, we measured the amount of NO• produced at increasing doses of DETA-NO•; as seen in Fig. 2A, 1000 ␮M DETA-NO• generates approximately 7 ␮M NO• by 4 h. This concentration is similar to reported concentrations of NO• measured with brain microdialysis during brain ischemia/reperfusion (1–10 ␮M) (Malinski et al., 1993). In cultures 14 –17 DIV, total cell death was NO• concentration-dependent with an EC50 of 62 ␮M concentration (Fig. 2B). Under certain conditions NO• can induce apoptosis in cortical neurons (Palluy and Rigaud, 1996); therefore, NO•-treated neurons were examined for established signs of late-stage apoptosis including microscopic evidence of DNA fragmentation/condensation. Fig. 3A illustrates typical neurons stained with a DNA dye following exposure to increasing concentra-

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Fig. 1. Montage of LSCM fluorescent images of neurons (17 DIV) of either A, a, control neurons, B,b, neurons treated with either 30 ␮M DETA-NO•, or C,c, or with 1000 ␮M DETA-NO• for 3 h and then assayed 24 h later for signs of cell death. In A, B, and C, neurons were stained with both calcein-AM (green) and propidium iodide (PI, red) to determine viability. In a, b, and c neurons were stained with a triple label of calcein AM, PI, and the mitochondrial membrane potential dye, TMRM (red). The far right pictures labeled Mito are an enlargement of the area in the white box demonstrating TMRM-labeled mitochondria (arrows). As seen in A, a, control healthy neurons show typical intact cytoplasmic green staining and robust TMRM staining, respectively. In contrast, neurons exposed to 1000 ␮M DETA-NO• show depleted cytoplasmic-specific staining with severely disrupted plasma membranes (C), increased red nuclear stain, and severely reduced or absent TMRM fluorescence compared to control (c), all signs of severely injured or dead neurons. Original magnification 60⫻ oil immersion.

tions of NO• compared to untreated neurons. An average of 74% (SEM ⫹/⫺ 8%) of neurons by 24 h had evidence of apoptotic cell death when exposed to 1000 ␮M DETA-NO• compared to 20% of the control nontreated neurons; this percentage was similar to that at 6 h, suggesting that rapid apoptotic changes were occurring. NO• was associated with increasing neuronal DNA condensation and/or fragmentation at all concentrations tested (10, 30, 100, 300, and 1000 ␮M of DETA-NO•) at both the 6- and the 24-h time points (3 h exposure) compared to controls (Fig. 3B). There was no statically significant difference in percentage cell death between the 6- and 24-h time points, indicating that at each concentration maximum apoptotic changes occurred by at least 3 h of NO• exposure time; a concentration response curve was plotted for both the 6- and the 24-h time points (Fig. 3C). Short NO• exposure times of 5 or 20 min did not result in a statistically significant difference in cell death compared to time-matched controls (Fig. 3D). The nontreated controls had a similar rate of cell death throughout the 24 h of the experiment (range 29–38 ⫹/⫺ 4%). The activation of caspase-3 and the release of the proapoptotic factor cytochrome-c (cyto-c) can be associated with ap-

optotic-like cell death. Additionally, cyto-c translocation from the mitochondria to the cytosol prior to MPT may be the result of NO•-peroxynitrite-mediated nitration (Hortelano et al., 1999; Ghafourifar et al., 1999; Alonso et al., 2002). Neurons maintained in glial-enriched medium and exposed to NO• had a linear increase in fluorescent intensity consistent with increasing caspase-3 activation over time, which was blocked in the presence of the caspase-3 inhibitor, DEVD-fmk (Fig. 4A). Neurons exposed to DETA-NO• (10, 100, or 1000 ␮M; 37°C) for 3 h in glial-enriched medium and assayed at 6 and 24 h resulted in more diffuse cytoplasmic staining, particularly in the more injured neurons, while the nontreated cells demonstrated more punctate mitochondrial located cyto-c immunoreactivity. Fig. 4B demonstrates typical neurons exposed to 1000 ␮M NO• for 3 h versus nontreated controls. NO• causes rapid concentration-dependent mitochondrial depolarization which is prevented by MTP inhibitors NO•-induced mitochondria membrane dysfunction in cortical neurons has been only recently reported in a similar cell system (Almeida et al., 2002). Our goal was to further

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Fig. 2. A. Typical concentration–response curve indicating the concentration of NO• released by 0, 10, 30, 100, 300, 500, and 1000 ␮M DETA-NO• after approximately 4 h using a porphyrinic NO• microsensor. NO• measurements were determined under the same experimental conditions used in this study using an NO• calibration curve prepared from known concentrations of the NO• generator, SNAP (S-nitroso-N-acetyl-D,L-penicillamine), according to the manufacturer’s protocol. At the highest concentration tested, 1000 ␮M, DETA-NO• released approximately 7 ␮M NO• (dotted lines). B. Concentration– response curve for DETA-NO• induced neuronal cell death as determined using the live/dead assay (see Experimental Methods). Cells were exposed to 0, 10, 30, 100, 500, and 1000 ␮M DETA-NO• for 3 h and the percentage cell death was determined at 24 h. EC50 was calculated to be 62 ␮M. Percentage cell death was calculated as the ratio of the number of cells with positive PI staining to the total neurons studied. An average of 70 neurons were examined per group.

characterize the mechanism by which NO• mediates these critical changes. The effect of NO• on mitochondrial membrane potential (⌬ ␺m) was examined by labeling neurons with the mitochondrion-selective membrane potentialdriven dye, TMRM. At low concentrations and following exposure to DETA-NO•, TMRM fluorescence does not undergo dequenching or increase of signal; Fig. 5A demonstrates typical cortical neurons with TMRM-labeled mitochondria. In all experiments the ETS uncoupler, carbonyl cyanide m-chlorophenylhydraxone (CCCP), is added at the end of the experiment as a positive control, resulting in a rapid collapse of ␺m and loss of TMRM signal (Fig. 5B). At all concentrations of NO• tested (10, 100, 1000 ␮M) a progressive ⌬ ␺m (depolarization) was seen compared to nontreated cells (Fig. 5C); the ⌬ ␺m from baseline was 14, 37, and 62%, respectively, after 10 min of treatment. As demonstrated in Fig. 5B NO•-induced depolarization was rapid on onset and once initiated continued for the full 20-min incubation time. Infrequently, at the 1000 ␮M concentration an immediate moderate to large rapid depolarization followed by a slower rate of depolarization occurred. Although the majority of neuronal mitochondria underwent some form of depolarization (83 ⫹/⫺ 10%) approximately 17–25% of cells did not, indicating some heterogeneity in the response. The magnitude of loss of TMRM signal following DETA-NO• treatment ranged from 20 to 70% from the baseline signal in individual neurons (when assayed at the 15-min posttreatment time point). The removal of the DETA-NO• (1000 ␮M) after 1, 2, or 5 min of exposure time did not significantly alter the rate of depolarization compared to controls, suggesting an immediate and rapid effect

on membrane depolarization once initiated (data not shown). It has recently been demonstrated that NO• can induce MPT in a concentration-dependent manner in isolated mitochondria (Brookes et al., 2000). We therefore wished to test whether NO• concentration-dependent ⌬ ␺m preceded induction of MPT in our cortical neurons. As seen in Fig. 6A, the addition of 50 ␮M of bongkrekic acid (BA), an inhibitor of mitochondrial adenine nucleotide translocator, prevented the typical NO•-induced depolarization compared to those treated with NO• alone. Experiments using 1 ␮M cyclosporin (CsA), a less specific inhibitor of the mitochondrial transition pore, also resulted in a statistically significant prevention of the NO•-induced mitochondrial depolarization compared to the non-CsA NO•-treated group at all time points tested (Fig. 6B). MTP inhibitors do not prevent NO•-induced cell death MPT is involved in both necrotic and apoptotic cell death (reviewed in Crompton, 1999), and since we had established that MTP inhibitors prevent NO• concentration-dependent ⌬␺m, we next tested whether this inhibition resulted in neuroprotection. The addition of BA (50 ␮M) was not neuroprotective, and, in fact, potentiated NO•-induced cell death when assayed at the 6- and 24-h time points. Despite a short incubation time at which the BA control alone was not neurotoxic, there was no statistically significant change in NO•-induced death in the BA-treated vs untreated groups compared to controls (Fig. 6C). We repeated the experiments identically with low-concentration CsA(1 ␮M) and as seen in Fig. 6C, CsA did not inhibit NO•-induced cell death but instead potentiated cell

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Fig. 3. A. Montage of neurons stained with Hoechst DNA dye. Neurons were treated with medium. ⫹/⫺ 1000 ␮M DETA-NO• for 3 h and were assayed 24 h later (37°C). Control neurons demonstrate normal ovoid-shape nuclei (a, b; solid arrows) in contrast to those exposed to DETA-NO•, which demonstrate condensed/fragmented DNA consistent with apoptotic cell death (c,d;open arrows). Original magnification 60⫻. B, C. Concentration–response curve and bar graph of DETA-NO• - induced apoptotic neuronal cell death as determined by percentage DNA fragmentation/condensation. Cells were exposed to 0, 30, 100, 300, or 1000 ␮M DETA-NO• for 3 h and percentage cell death was determined at 6 and 24 h. Percentage cell death was calculated as the ratio of the number of cells with DNA fragmentation/condensation as determined by Hoechst dye staining to the total neurons studied. An average of 100 neurons were examined per group; experiment was repeated in triplicate; data are means ⫹/⫺ SEM. D. Bar graph demonstrating the percentage of DNA fragmentation/ condensation measured following increasing exposure time to 1000 ␮M DETA-NO• (gray bars) compared to time-matched controls (black bars). Data are means ⫹/⫺ SEM from triplicate experiments with approximately 200 cells per group. (*P ⬍ 0.05; ANOVA).

death similar to that observed with BA treatment; treatment with NO• in the presence of BA killed 81 ⫹/⫺ 7% of neurons while NO• in the presence of CsA, killed 99 ⫹/⫺ 3% by 24 h. Neither the CsA nor the BA control group was associated with an increase in cell death compared to the untreated group. Reduction of ⌬␺m by NO• is prevented by NMDA receptor inhibition Recent data suggest that glutamate receptor-mediated NOS activation could trigger MPT pore opening (Almeida et al., 2001) and that NO•-dependent damage to

neuronal mitochondria may involve the NMDA receptor (Stewart et al., 2002). Therefore, we next examined the role of NO• and the NMDA receptor in our system. In cortical neurons treated with DETA-NO• (1000 ␮M), the addition of the NMDA receptor antagonist, APV 2-amino-5-(10 ␮M) statistically and significantly attenuated DETA-NO•-induced ⌬ ␺m compared to controls alone; APV treatment alone did not alter ⌬ ␺m (Fig. 7A). To directly test the effect of NO• at the NMDA receptor, whole cell currents were measured using patch-clamp techniques in cortical cells at 7, 14, and 21 DIV. To perform these studies neurons were exposed to an extra-

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Fig. 4. A. Bar graph illustrating the effect of DETA-NO• on caspase-3 enzyme activity based on measurement of intensity of fluorescence of a caspase-3 cleaved substrate compared to nontreated cells. Neurons (14 DIV) were exposed to either 0 or 1000 ␮M DETA-NO• for 0, 2, 6, or 24 h, and relative fluorescent intensity was determined. Data are means ⫹/⫺ SEM from duplicate experiments with approximately 20 –25 cells per group. (*P ⬍ 0.05; ANOVA with pairwise multiple comparisons using the Tukey test). B. Enlarged epifluorescent images of cyto-c immunoreactivity within neurons under either control conditions (top, CON) or following treatment with 1000 ␮M DETA-NO• for 3 h (lower panel, ⫹ NO•). Control neurons demonstrate typical punctate areas of staining (open arrows) versus less intense punctate staining and more diffuse cytosolic staining indicating cyto-c release in the DETA-NO• neurons (closed arrows). Original magnification 60⫻.

cellular solution containing 10 ␮M NMDA ⫹/⫺ 1000 ␮M DETA-NO• or artificial cerebrospinal fluid (control). Neurons (14 DIV) treated with DETA-NO• demonstrated sustained potentiation of the NMDA-induced current by 68 ⫹/⫺ 10% compared to controls (Fig. 7B) (Solenski et al., 2002). Upon addition of DETA-NO there was no increase in the background-recorded current, suggesting a direct effect on neuronal plasma membrane potential.

Both NMDA and DETA-NO• produce mitochondrialspecific nitrotyrosine (NT) staining In our experimental paradigm it remained unclear how NO• so rapidly depolarized mitochondria in a NMDAreceptor-dependent manner. Since NMDA receptor activation is associated with increased NT immunostaining in association with acute hypoxic neuronal death (Ochiai-

Fig. 5. A. LSCM image of TMRM stained neurons in a single microscopic field of view (left). A typical sampled perinuclear region of interest is shown with an individual neuron (white box) shown at higher magnification (right) to better demonstrate TMRM labeling of individual mitochondria (arrows). B. Monochrome image of TMRM-labeled neurons at baseline, 10 min, and after the addition of CCCP in untreated CON (top) versus DETA-NO•-treated cells (1000 ␮M; bottom). C. Representative graph of the concentration effect of DETA-NO• on relative ⌬␺m measured by TMRM fluorescence versus time. A statistically significant change in mitochondrial membrane depolarization was demonstrated starting at the 5-min posttreatment time point compared to nontreated control cells at all concentrations tested (*P ⬍ 0.05). Measurements are normalized to baseline (pretreatment) readings and are based on analysis of approximately 15–20 neurons per group for this experiment.

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Fig. 6. A, B. Representative graphs of the effect of the MTP inhibitors, bongkrekic acid (BA) (A), and cyclosporin A (CsA) (B) on DETA-NO• induced ⌬ ␺m compared to nontreated controls. Continuous exposure to 1000 ␮M DETA-NO• alone resulted in a typical depolarization while coincubation with BA resulted in a statically significant prevention of the depolarizing effect (*P ⫽ 0.002). The addition of BA alone did not differ from control. Similarly, the addition of CsA statistically prevented the DETA-NO•-induced depolarization (**P ⫽ 0.001). Data are based on an average of 150 neurons per group; the experiment was repeated in triplicate. C. Bar graphs illustrating the percentage cell death measured by the calcein-AM/PI live/dead assay in the presence of DETA-NO• ⫹/⫺ BA (50 ␮M) or ⫹/⫺CsA (1 ␮M) compared to nontreated cells. Neurons were exposed to 3 h of 1000 ␮M DETA-NO• and assayed 24 h later. Data are means ⫹/⫺ SEM from duplicate experiments with approximately 20 –25 cells per group. (*P ⬍ 0.05 compared to the respective control; ANOVA with pairwise multiple comparisons using the Tukey test). Each field represents 42 s time.

Kanai et al., 1999), we next tested whether NO• and/or NMDA treatment could produce peroxynitrite as a mediator of mitochondrial membrane dysfunction. We found that when cortical neurons were exposed to either DETA-NO• or to NMDA ⫹ glycine this resulted in significant immunoreactivity for NT residues (Fig. 8A). Following treatment with NO• both a punctate and a diffuse staining pattern occurred along the cell body and in the dentritic processes compared to nontreated cells (Fig. 8B). The positive controls (peroxynitrite and an alternative NO• generator [SIN-1]) demonstrated similar cell body staining, indicating adequate antibody specificity. The negative control (degraded peroxynitrite) demonstrated no significant staining. We observed that the punctate staining pattern was similar to the pattern seen in TMRM-labeled mitochondria; therefore, we

tested whether these stained regions were associated with mitochondria. Using colocalization immunocytochemical experiments with an antibody against an established mitochondrial matrix protein (heat shock protein 75 or grp75), we determined that NT staining was specifically within mitochondria (Fig. 9). The differences in the microscopic imaging technique and postfixation techniques (colocalization experiments used membrane permeablizing techniques) account for the visual differences in NT staining pattern in Figs. 8A (laser confocal microscopy), 8B, and 9 (epifluorescent microscopy). Morphological and quantitative analysis was confined to the digital images obtained from the epifluorescent microscope using the MetaMorph software thresholding and colocalization applications.

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Fig. 7. A. Typical graph demonstrating the effect of the NMDA receptor antagonist (APV) on DETA-NO• induced ⌬␺m. The coincubation of APV (5 ␮M) with DETA-NO• (1000 ␮M) resulted in a statically significant prevention of the depolarizing effect of DETA-NO• alone (*P ⬍ 0.05). A mean of 150 neurons was assayed per group and the experiment was repeated in triplicate. B. Typical whole cell current recordings after applying NMDA (100 ␮M, bar) in cortical neurons at 7, 14, and 21 DIV ⫹/⫺ DETA-NO• (1000 ␮M). At 7 DIV, neither NMDA nor NMDA ⫹ DETA-NO• elicited a measurable current (left v. right, respectively, n ⫽ 7). At 14 and 21 DIV, NMDA currents were potentiated following DETA-NO• treatment by 68 ⫹/⫺ 10 and 73.7 ⫾ 4.1%, respectively. NMDA was applied via a modified U-tube device. Data are presented as means ⫾ SEM.

Reducing reactive nitrogen species formation reduces NT immunoreactivity and neuronal death in a NMDAreceptor-dependent manner We reasoned that if peroxynitrite was mediating NO• or NMDA-induced neuronal cell death or mitochondrial membrane changes, then preventing its formation may attenuate these effects. We chose to use a compound with a mechanism of action similar to MnSOD, an important mitochondrial enzyme which dismutases superoxide radical species, the critical substrate for peroxynitrite formation (see review Macmillan-Crow and Cruthirds, 2001). Furthermore MnSOD prevents neural apoptosis by suppressing peroxynitrite production and mitochondrial dysfunction (Keller et al., 1998). We treated neurons with a cell-permeable SOD/ catalase mimetic, “EUK-134” (manganese 3-methoxyN,N⬘-bis (salicylidene)ethylenediamine chloride; 5 ␮M) in the presence and absence of DETA-NO• (1000 ␮M) or NMDA (100 ␮M) for 3 h and reanalyzed for NT staining (Fig. 10A). We chose the compound EUK-134 since it is a manganese salen (Mn–Salen) complex with a mechanism of action similar to MnSOD and has been shown to significantly decrease nitration of enzymatic proteins in neurons exposed to a variety of oxidative toxins (Pong et al., 2000).

EUK-134 ameliorates cellular damage caused by both oxidative and nitrosative stresses (Sharpe et al., 2002). Treatment with EUK-134 significantly reduced NO•- and NMDA-induced average NT clusters by 42% in the NO•treated cells and by 58% in NMDA-treated cells when compared to untreated controls; not all neurons were protected, indicating some degree of heterogeneity (Fig. 10B). The NO•- or NMDA-untreated controls and the EUK-134 alone group had only nonspecific background staining. It was difficult to determine the effect of EUK-134 on cell viability in these ethanol–acetic acid fixed neurons; therefore, we performed separate but identical viability studies using the calcein-AM and PI live/dead assay as previously described. NO•-induced death was reduced by EUK-134 by 60% and NMDA-induced neuronal death by 59% compared to those not treated with EUK-134 (Fig. 10C). We next tested whether treatment with EUK-134 during DETA-NO• exposure prevented NO•-induced or NMDAinduced ⌬ ␺m; treatment with EUK-134 did not significantly prevent NO•-induced ⌬␺m but there was a small but statistically significant prevention of NMDA-induced ⌬ ␺m (Fig. 11A and B); the absolute difference over time was 15–22% between the EUK-134 ⫹ NMDA-treated group

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Fig. 8. A. Representative epifluorescent images of immunoreactivity of nitrotyrosine (NT), in untreated (A,a) or either NMDA ⫹ glycine treated (100 ␮M, 15 ␮M, respectively) (B,b) or DETA-NO• (1000 ␮M) (Cc)-treated neurons for 3 h. A similar intense diffuse and punctate staining pattern was observed in both the DETA-NO• -and the NMDA-treated neuons when compared to control conditions (white arrows). Original magnification 100⫻. B. Representative LSCM images of immunoreactivity for nitrotyrosine, a marker for peroxynitrite formation, in untreated (a) or in DETA-NO• (1000 ␮M ⫻ 3 h) in cortical neurons (14 DIV) (b), compared to positive (c, e) and negative (d) controls. DETA-NO•-treated cells had significant staining throughout the perikaryon and in the dendritic processi similar to the positive time-matched controls using peroxynitrite (3 mM) and SIN-1 (100 ␮M). Original magnification 100⫻.

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Fig. 9. Epifluorescent images demonstrating colocalization of NT staining, a marker for peroxynitrite formation, with a mitochondrial matrix protein, HSP-75. Nontreated neurons stained for the mitochondrial marker HSP-75 (red) and demonstrated minimal NT immunoreactivity (green), (CON, top); note that the CON has nonspecific NT staining of very low intensity (average color ⫽ 55 [range 1–255, 8-bit image]). In contrast, neurons treated with 1000 ␮M DETA-NO• for 3 h (⫹NO, bottom) demonstrated typical cytosolic and punctate staining with the latter colocalizing with HSP-75 staining, resulting in yellow staining (white arrows). (original magnification 60⫻).

and the NMDA-treated only group, which became statistically significant after 15 min following NMDA treatment. Discussion This study characterizes the effects and mechanisms of NO•-mediated neurotoxicity in primary cortical neurons under aerobic conditions. Based on our experimental results we conclude that NO• in micromolar concentration is a potent neurotoxin associated with mitochondrial dysfunction involving both a NMDA receptor-dependent and -independent mechanism. Specifically NO• causes a time- and concentration-dependent rapid mitochondrial depolarization closely linked to NMDA receptor potentiation. NO• and NMDA cause neuronal injury at least in part by intramitochondrial peroxynitrite formation, and NMDA-induced mitochondrial depolarization is in part dependent on peroxynitrite formation. Although NO• induces MPT which has been associated with cell death, NO•-induced MPT induction was not a primary cause for neuronal cell death in our cell system; alternatively measures to reduce peroxynitrite formation were associated with neuroprotection.

This study established that NO• under aerobic conditions results in significant neuronal cell death in a time- and concentration-dependent manner. In cortical neurons exposed to NO• both apoptosis and necrosis occurred simultaneously with the rapidity and dominant type of cell death depending on the exposure time and concentration of NO•. Since the distinction between “apoptotic” and “necrotic” forms of neuronal death is inexact (see review Roy and Sapolsky, 1999; Niquet et al., 2003) we were less interested in defining the exact type and more interested in determining whether both forms actually existed as a consequence of NO• exposure. The majority of the experiments in this study consisted of neurons exposed to high concentrations of NO• which promoted necrotic over the apoptotic type of cell death. We report rapid NO•-induced mitochondrial depolarization. This has been reported in neuronal-like and hippocampal cells mainly under conditions of oxygen– glucose deprivation, with, in some studies, speculation that MPT is critically involved (Kayahara et al., 1998; Brorson et al., 1999; Bal-Price and Brown, 2000; Kindler et al., 2003). Our data extend the findings of a related study of NO•-induced

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Fig. 10. A. Representative phase and epifluorescent microscopy images of neurons stained for NT following treatment ⫹/⫺ DETA-NO• (1000 ␮M) or ⫹/⫺ NMDA (100 ␮M) for 3 h and in the presence and absence of a SOD/catalase mimetic EUK-134 (5 ␮M) compared to nontreated controls. Far left panels demonstrate typical background NT staining (CON), which increases significantly with either DETA-NO• or NMDA treatment (center). NT staining is attenuated by coincubating with EUK-134 (far right). Neurons pretreated with EUK-134 in the NMDA group also revealed better overall cellular morphology compared to the DETA-NO• group (arrow). B. Bar graph indicating the effect of EUK-134 (5 ␮M) on NT immunoreactivity ⫹/⫺ NMDA (100 ␮M) or DETA-NO• (1000 ␮M). Neurons were exposed to DETA-NO• or to NMDA ⫹/⫺ EUK-134 for 3 h, fixed, and immunostained for NT formation. Using morphometric analysis the average number of NT cluster was measured. Data are based on 50 neurons per group; *P ⬍ 0.05. C. Bar graph indicating the effect of EUK-134 on DETA-NO-or NMDA-induced cortical neuron death using the live/dead calcein-AM/PI fluorescent assay (see Experimental Methods). Neurons were treated with EUK-134 (5 ␮M) ⫹/⫺ either NMDA (100 ␮M) or DETA-NO (1000 ␮M) for 3 h and assayed immediately. Data are based on 150 –250 neurons per group (single experiment, 50 images, 3–5 neurons/image).

mitochondrial depolarization in pure cortical cultured neurons under oxygen– glucose deprivation conditions (Almeida et al., 2002). Almeida et al. report NO•-mediated ⌬␺m as the result of ATP depletion and inhibition of mitochondrial respiration. In our study we report that a similar depolarizing effect can occur under aerobic conditions not only following exposure to NO• but also to NMDA. In addition, we report that NO•-mediated ⌬␺m can be completely prevented by antagonizing the NMDA receptor and that NO• exposure results in potentiation of NMDA-induced NMDA receptors, suggesting a critical relationship between these two neurotoxins. We cannot eliminate the possibility that laser-induced phototoxicity with NO-related free radical formation is not occurring. The relationship among NO•, Ca2⫹, free radicals, and excitotoxic-induced mitochondrial dysfunction is complex in neurons with the exact mechanism unknown (Hoyt et al.,

1992; Keelan et al., 1999; Stout et al., 1998; Scanlon and Reynolds, 1998; Ward et al., 2000). Our report corroborates other studies suggesting that NO• exhibits its effects through an NMDA-receptor mechanism (Hoyt et al., 1992; Almeida et al., 1999; Stewart et al., 2002) with the most accepted pathway being NMDA receptor Ca2⫹-mediated activation of cytosolic nNOS in susceptible neurons. In this scenario cytosolic NO• directly enters the mitochondria and imparts direct or indirect NO•-related damage. Our study focused on an alternative pathway involving the possibility of NMDAinduced activation of Ca2⫹-dependent mtNOS, resulting in similar NO•-related mitochondrial damage including the formation of peroxynitrite. This is based on compelling data that NMDA increases mitochondrial Ca2⫹ uptake (Peng and Greenamyre, 1998) and that glutamate-mediated neuronal cell death requires mitochondrial Ca2⫹ uptake (Stout et al., 1998). Our electrophysiological studies indicate that NO• at

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Fig. 11. Graphs illustrating the effect of the SOD/catalase mimetic, EUK-134, on NMDA or NO•-induced mitochondrial depolarization. Neurons were coincubated with EUK-134 (5 ␮M) ⫹ DETA-NO• (1000 ␮M) and TMRM fluorescence was measured over a 25-min period. A. The addition of EUK-134 (5 ␮M) did not statistically prevent the typical DETA-NO•-induced ⌬␺m compared to the control group. B. In contrast treatment with EUK-134 significantly but only partially prevented the NMDA-induced mitochondrial depolarization (5 ␮M). Data are means ⫹/⫺ SEM from duplicate experiments with approximately 20 –25 cells per group (*P ⬍ 0.05; ANOVA with pairwise multiple comparisons using the Tukey test).

high concentrations consistently potentiates NMDA-activated NMDA receptor whole-cell currents in primary cultured cortical neurons (⬎7 DIV). This finding was somewhat unexpected. The role of NO• in NMDA receptor modulation is complex (see review by Aizenman et al. 1998). While NO• down-regulation of this receptor through a redox modulatory site is well described, NO• potentiating effects are less well defined (see the extensive reviews by Lipton et al., 1993, 1996, 1998a, 1998b; Choi and Lipton, 2000). In support of our data, there is a single related study using HEK-293 cells transfected with NMDA receptors; in this study using patch-clamp techniques, the authors report NO•-potentiated glutamate-mediated response of the recombinant NMDA receptors (Gbadegesin et al., 1999). There are some important differences between this study and ours. This study used nonneuronal cells and physiologically relevant concentrations of a different NO• generator with a substantially shorter half-life (DEA-NO• t1/2 is 2 min while DETA-NO• t1/2 is 20 h). The authors conclude that the observed NO•-induced NMDA receptor potentiation may represent a normal physiological response of NO• at the NMDA receptor. We report robust and reproducible potentiation at the same high concentrations of NO• that provoke ⌬␺m and kill neurons in our culture system. How could NO• potentiate the NMDA receptor? One explanation is that NO• is causing mitochondrial respiratory failure with ATP depletion, associated with glutamate release and subsequent NMDA receptor activation as suggested by other investigators in different experimental paradigms (Bal-Price and Brown, 2001; McNaught and Brown, 1998). Alternatively, NO• could be acting directly at the presynaptic site since studies of hippocampal synaptic vesicles demonstrate NO• triggering of neurotransmitter exocytosis and in cerebellar granule cells reveal that blocking exocytosis prevents NO•-mediated apoptosis (Sporns and Jenkinson, 1997; Leist et al., 1997). Lastly, NO• can be activating endogenous cGMP with subsequent synaptic potentiation (Bon and Garthwaite, 2001) or could be stimulat-

ing the release of glycine, a cofactor for NMDA activation (Akira et al., 1994). We are currently testing these potential explanations with ongoing electrophysiological experiments in our laboratory. We report new indirect evidence of neuronal intramitochondrial peroxynitrite/RNS formation as a consequence of treating with either NMDA or NO•. Reducing the peroxynitrite formation significantly attenuates neuronal cell injury. Reducing peroxynitrite formation, however, revealed no change in NO•induced ⌬␺m and only a partial prevention of NMDA-induced ⌬␺m. These data suggest that in neurons high-dose exogenously applied NO• may be exhibiting its neurotoxic effect more directly on neuronal mitochondria and more independently of peroxynitrite formation. Much data exist supporting that peroxynitrite and MnSOD play a central role in apoptosis and anti-apoptotic mechanisms (respectively) in neural cells (Estevez et al., 1995; Keller et al., 1998), but there far fewer reports suggesting that MnSOD protects against necrosis, particularly in neurons (Gow et al., 2000). Our data suggest that even under conditions promoting necrosis (over apoptosis) reducing peroxynitrite formation can be neuroprotective. Our data also corroborate some reports that MnSOD protects nNOS neurons from glutamate and NMDA toxicity (Gonzalez-Zulueta et al., 1998, Li et al., 1998). Since NT staining could be associated with other reactive nitrogen species, another toxic nitrogen species could be mediating cell death; due to the complexity of SOD and NO interactions in living cells we cannot unequivocally conclude that PN formation alone is responsible for our findings. Notably we chose a concentration of EUK-134 known to be neuroprotective under preincubation conditions in other cell culture systems (Pong et al., 2002) but it is possible that other concentrations of this compound or alternatively lower concentrations of DETA-NO• would have demonstrated different results. In our pilot studies using 10-fold higher and lower concentrations of EUK-134, we did not see any significant change in cell viability (data not shown). Since high concentrations of NO• promote necrosis this may explain why partial

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rather than complete neuroprotection was demonstrated with EUK-134 treatment. Alternatively, multiple mechanisms of cell death including those unrelated to NO• or peroxynitrite formation may be occurring. The finding that DETA-NO• demonstrated positive NT staining in a concentration-dependent manner was somewhat unexpected in light of a recent contradictory report (Vidwans et al., 1999), but may be supportive of NMDAreceptor mediated peroxynitrite formation. The difference could be due experimental conditions including the use of pure versus mixed cortical cultures and/or the type of NT assay. Our study used high-resolution confocal imaging at the level of the mitochondria of fixed and membrane permeabilized neurons rather than assaying for nitrosylated proteins in the culture medium. The important question is how does DETA-NO• treatment which primarily generates NO• (rather than peroxynitrite) result in our observed intramitochondrial NT staining? One explanation is that since DETA-NO• potentiates NMDA receptor activation (as shown in our electrophysiological studies), peroxynitrite formation could be the result of NMDA receptor Ca2⫹-mediated activation of mtNOS. Our data showing that EUK-134 treatment results in reduced NT immunoreactivity support this mechanism. An alternate explanation is that DETA-NO• releases NO• into the cytosol which then may be directly perturbing the ETS, causing increased superoxide free radical formation followed by peroxynitrite formation. Either of these explanations could result in the observed mitochondrial in situ NT staining pattern but we would predict more prominent cytosolic staining in the latter scenario. Experiments designed to prevent mitochondrial Ca2⫹ uptake (in progress) or the use of specific inhibitors of mtNOS when available could help differentiate between these two possibilities. A final important finding is that although MTP inhibitors prevented NO•-induced mitochondrial depolarization they did not result in neuroprotection by 6 or 24 h of observation. These data suggest that although NO• can depolarize mitochondria at least in part by facilitating MTP as shown by others (Brookes et al., 2000; Kindler et al., 2003), preventing NO•-mediated MPT is not a prominent mechanism of NO•-induced cell death in our neuronal culture system. These data are intriguing since BA and CsA have been shown to prevent NMDA-induced apoptosis in a variety of cell types including neurons (Schinder et al., 1996; Budd et al., 2000). This may be related to the use of high DETA-NO• dose promoting necrotic cell death. In a single contrasting study using younger-aged cortical neurons BA did not prevent NO•-induced mitochondrial depolarization; culture conditions including age, the presence of glia, or the simultaneous use of oligomycin ⫹ BA may be responsible; the same report, however, corroborates our findings of a lack of BA to attenuate NO•-mediated cell death (Almeida et al., 2001). There are important caveats with this study including defining the role of glia in modulating our results; for example, our electrophysiological recordings were per-

formed in the physical absence of astroglia cells which play an important role in neurotransmitter regulation; additional evidence is necessary to determine whether this NO•-induced NMDA evoked receptor potentiation also occurs in vivo. It is also well-recognized that the biochemistry of NO• and of NO• generators is complex, particularly in the neuron with multiple locations for its endogenous formation, routes of action, and NO•-related reactants being formed, depending on environmental conditions or the redox milieu. In our experimental paradigm we deliberately chose a high concentration of DETA-NO• with a concentration of NO• equivalent to that measured during ischemic brain injury although the true endogenous [NO•] released in vivo is unknown. At these concentrations secondary NO•-related effects such as activation of cGMP could be occurring. Although NT staining is consistent with peroxynitrite staining and is an accepted marker of such, recent evidence indicates that a variety of other reactive nitrogen species (RNS) maybe important in the formation of NT. Notably, however, peroxynitrite is the most prominent RNS in vivo. Corroborating evidence of NO• or NMDA-mediated nitrosylated mitochondrial membrane or matrix proteins would be useful. In summary we conclude that that NO•-mediated mitochondrial dysfunction and neurotoxicity involve both NMDA receptor and peroxynitrite-dependent and -independent mechanisms. We report new data on NO•-mediated potentiatation of NMDA-induced receptor activation and evidence of in situ mitochondrial formation of reactive nitrogen species as a consequence of either NO• or NMDA exposure. NMDA receptor activation and mitochondrial NO• formation are intricately related to mitochondrial function and ultimately to cell viability. Better understanding of how mitochondria provide the critical link between NO•– NMDA receptor activation and neuronal cell death has significant implications for excitotoxic-mediated human disease states including ischemic stroke. In particular, more research is needed to elucidate the specific consequences of in situ mitochondrial NO•-related perturbation on membrane protein signaling and other functions on the activation or alteration of less-defined enzymes systems such as mtNOS or the effect on mitochondrial Ca2⫹ buffering or ionic homeostasis in general.

Experimental methods Preparation/maintenance of primary cortical neuron cultures Primary neuronal cultures were prepared using an adaptation of the “Banker” technique in which neurons were grown on coated glass coverslips and inverted over a separate nutritive glial support (Banker et al., 1998). Astroglial cells were first prepared from 1- to 2-day-old postnatal rat

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pups; brains were removed, and the debrided cerebral hemispheres were minced and treated with 1% DNAse with trypsin and the supernatant was filtered. The supernatant was centrifuged and resuspended in glial medium and the cells were counted and plated; cells reached confluency in 10 –14 days and were fed every 4 days. Once glial cells were confluent, in a separate protocol, a timed pregnant E19 Sprague–Dawley rat was euthanized and the fetuses and their brains were quickly removed; the cortex was debrided, dissected, trypsinized, washed, and dissociated. Cells were plated onto specially cleaned, poly-L-lysine-coated coverslips; after neurons attached they were inverted over previously prepared 2-week-old glial cultures using Teflon support rings. Neuronal maintenance medium consisted of MEM with Earle’s salts, pyruvic acid, glucose, ovalbumin, and N2 supplement. N2 supplement contains insulin, progesterone, putrescine, selenium dioxide, transferrin, and MEM with Earle’s salts. Glial medium contained MEM with Earle’s salts, glucose, pyruvic acid, and horse serum. Neurons were fed every 10 days. By day 14 the neurons formed synapses as determined by colocalization of the NR1 (NMDA receptor subunit) with the synaptic vesicle protein, synaptophysin (Solenski et al., 2002), and by electrophysiological studies demonstrating both GABA and NMDA receptor activation. Neurons stained positively for MAP with minimal contaminating glial elements as determined by GFAP staining. For consistency, all experiments were performed 14 –17 DIV. All procedures involving the use of animals were approved by the University of Virginia Animal Care and Use Committee and strictly followed the NIH guidelines for the use of laboratory animals. Measurement of NO• generated by DETA-NO• The amount of NO• released by DETA-NO• (SigmaAldrich Co., St. Louis, MO) was measured using a standard NO• electrode (Iso-NO, World Precision Instruments, Stevenage, UK); the electrode was calibrated with known concentrations of SNAP (S-nitroso-N-acetylpenicillamine) under reducing conditions according to the manufacturer’s instructions; NO• released by 10, 30, 100, 300, 500, and 1000 ␮M DETA-NO• was measured at room temperature (RT) in medium (pH 7.4) mimicking experimental conditions, including that the DETA-NO• solution was allowed to sit for a minimum of 1 h prior to NO• measurement; in all experiments DETA-NO• was allowed to sit for 1 h prior to its use. DETA-NO•, a nitric oxide adduct, was chosen as the NO• generator due to its long half-life of approximately 20 h, particularly under tissue culture conditions (Fitzhugh and Keefer, 2000). We first determined the concentration of NO• generated by DETA-NO• as used in our cell system; fresh solutions of 10, 30, 100, 300, 500, and 1000 ␮M of DETA-NO• were prepared under conditions identical to those used during the experiments. The concentration of NO• was determined at 1– 6 h after preparation using a standard NO• porphyrinic microsensor (WPI). Previous re-

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ports suggest that DETA-NO•-released NO• concentration plateaus within minutes (Griffiths and Garthwaite, 2001). EC50 was calculated from the concentration–response curve with the curve fit being determined by the software program Prism 3.0 (Graph Pad Software, Inc., San Diego, CA). In order to maximize NO• effects, we elected to use 1000 ␮M of DETA-NO• for the majority of experiments unless otherwise indicated. Measurement of cell viability— calcein-AM/propidium iodide and DNA staining Neuronal cultures were preloaded with calcein-AM (10 ␮M; Molecular Probes, Eugene, OR), and with PI (3 ␮g of 1 mg/ml stock, Molecular Probes) followed by either no or varying concentrations of DETA-NO• (25°C). Positive green (calcein-AM) and red (PI) labeling indicating live versus dead cells, respectively, was determined using laser confocal scanning microscopy (LCSM) techniques with a three channel Nikon PCM2000 confocal microscopy system coupled to a Nikon TE-200 epifluorescence microscope using PC-based C-Imaging software for digital image acquisition (Compix Imaging SimplePCI; C-Imaging Systems, Cranberry Township, PA). High-resolution digital images were further processed using Adobe Photoshop graphic software (Adobe, San Jose, CA). The absorption maximum for PI is 535 nm and the fluorescence emission maximum (EMmax) is 617 nm (488 line of an argon-ion laser); EX/ EMmax for calcein-AM is 494 nm and 517 nm (543 line of the Green HeNe laser.). Three sets of cells were labeled for each experimental variable. Total cell number was determined prior to experimentation in 10 separate (adjacent) fields of view using an inverted Zeiss phase microscope (40⫻ objective). The percentage of labeled to total cell number was determined to estimate cell detachment (evidence of late necrosis). Percentage cell survival was equal to live cells divided by the sum of dead and alive cells. To examine DNA structure, cells were stained with Hoechst 33342 for 10 min, washed, and examined under the microscope at UV excitation/emission 350/461 wavelength (nm). DNA morphology was examined for “fragmentation” defined as a cluster of punctate staining DNA or for “condensation” defined as stained DNA that was greater than or equal to 50% of the median control DNA circumference (age-matched). The effect of exposure and concentration of NO• on cell death was determined according to a standard paradigm; in some experiments neurons were treated for 3 h with NO• and then assayed for cell death at both 6 and 24 h following the NO• treatment. In a second set of experiments, neurons were continuously exposed to NO• for 1, 3, 6, and 24 h and immediately assayed for signs of cell death. Unless otherwise indicated each experiment was repeated in triplicate. This paradigm yielded approximately 50 –200 cells per group depending on the experiment.

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Measurement of relative ⌬ ␺m using laser scanning confocal microscopy (LSCM) Cells were loaded in triplicates with the dye, tetramethylrhodamine methyl ester dye in Hank’s buffered saline solution with Hepes (TMRM;100 nM; Molecular Probes). Depending on the experiment, cells were mounted on a chamber and fluorescent images of the cells were obtained using LSCM. Baseline images were obtained by rapid scanning at 1-min intervals for the first 4 min; at 5 min DETANO• was added to the cell chamber and rapid scanning continued for a total of approximately 14 min or fields. At the conclusion of each experiment (usually at 15 min) the protonophore (uncoupler) CCCP (5 ␮M; Sigma Chem. Co.) was added as a positive control to determine maximal membrane potential collapse. For TMRM the excitation/emission wavelength is 525/575 nm; a HeNe Green laser (543nm) was used with a 505-nm dichroic mirror and a 515-nm long-pass filter on the same LSCM system described above. The exact parameters needed to ensure fast scanning time with maximal resolution while ensuring minimal fluorescent quenching, photobleaching, and bleedthrough artifacts (when dual probes were used) were previously determined and standardized for all experiments. All data were normalized to baseline ⌬ ␺m and expressed as the percentage of relative fluorescence units. Three sets of cells were labeled and analyzed per group and each experiment was repeated in triplicate. The selected region of interest for analysis for this study was the perinuclear region of the neuron; approximately 90 –100 neurons were studied per group within each experiment. Measurement of caspase-3 activation Cells were incubated in the presence of either 500 ␮M DETA-NO• or 500 ␮M DETA-NO•⫹ 100 ␮M DEVD-fmk (CalBiochem) for 0, 2, 6, 12, and 24 h at 37°C. At the indicated assay time, cells were gently washed three times with phenol-free buffer and were incubated with PhiPhiLux caspase substrate reagent (OncoImmunin, Inc., Gaithersburg, MD, final concentration 2 ␮M) for 30 min at 37°C. Cells were then gently washed three times and immediately imaged. Cells were scored “positive” only if they demonstrated a fluorescence above an average background fluorescence; relative intensity was measured on raw images using Adobe PhotoShop software 6.0.

Cytochrome-c immunocytochemical staining Cortical neurons were fixed with 4% paraformaldehyde for 30 min at 25°C and washed with buffer. Cultures were blocked for 30 min at in goat serum and Triton X-100 in buffer and then incubated with primary antibody 1:1000 mouse anti-cyto c (Pharmingen) for 16 h at 4°C. Cultures were then washed again and incubated with FITC-conjugated secondary antibody in PBS (goat anti-mouse IgG1 Alexa 488, PharMingen) for 1 h at RT, washed, and mounted. Slides were viewed through the 40⫻ objective magnification lens of an Eclipse TE200 light microscope with a 100-W illumination pillar and system condenser and a four-filter linear slider. One multiband filter was used to view colocalized antibodies and two of three single-band filters, ultraviolet, blue, and green, were used to view the 350-, 488-, and 594-nm fluorescent markers, respectively. Antibodies were viewed with the appropriate filter cube and images were taken of individual neurons using a Photometric CoolSNAP CCD digital camera system with image capturing using MetaMorph version 6.01 software application (Limerick, Ireland) and further processed in Adobe Photoshop 6.0 (San Jose, CA). Images were collected for each primary antibody and of the colocalization of the two antibodies. Ten to 20 fields (80 –100 neurons) were imaged for each experimental group. 3-Nitrotryrosine immunocytochemical staining The method used was adapted from Viera et al. (1999) and as listed in the Upstate USA, Inc., manufacturer’s instructions. Neurons were fixed in ice-cold EtOH–acetic acid mixture for 1 min, washed with buffer, and blocked with 1% BSA–PBS for 30 min at RT and washed again. Cultures were incubated overnight at 4°C in primary antibody solution 1:50 rabbit anti-nitrotyrosine (Upstate) in 1% BSA– PBS. Cultures were washed and incubated with secondary antibody 1:50 FITC anti-rabbit IgG (Vector Labs) in 1% BSA–PBS for 1 h at RT and mounted. Controls were similarly prepared and processed; peroxynitrite (3–30 mM, Upstate) and 3-morpholinosydnonimine (SIN-1, 100 ␮M, Sigma) served as positive controls and PBS as the negative control; they were similarly prepared and processed. Fluorescent images were obtained using the LSCM system; 20 –30 neurons were imaged and analyzed using MetaMorph (Dowington, PA) Vers. 6.01 morphometric software for each experimental group.

Mitochondrial transition pore inhibition 3-NT and anti-grp 75 coimmunocytochemical staining Neurons were incubated with and without either 50 ␮M BA or 1 ␮M CsA [separate experiments] 20 min prior to the addition of 1000 ␮M DETA-NO• in the presence of glialenriched medium only. Each group of cells was loaded with calcein-AM and with PI, and positive green and red labeling indicating live versus dead cells, respectively, was determined using LSCM techniques as described above.

Neurons were fixed in ice-cold EtOH–acetic acid mixture, washed with buffer, followed by blocking with 1% BSA–PBS and Triton X-100 at 25°C, and washed again. Cultures were incubated at 4°C in primary antibody solution rabbit anti-nitrotyrosine IgG (Upstate, NY) and mouse antigrp75 IgG (Stressgene Biotech., BC, Canada) in 1% BSA–

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PBS. Cultures were washed and incubated with secondary antibody 1:50 FITC anti-rabbit (Vector Labs) and goat-antimouse IgG Alexa Fluor 594 in 1% BSA–PBS for 1 h at RT, washed again, and mounted. Fluorescent images were obtained using a Nikon Eclipse TE200 epifluorescent microscope as described above. MnSOD-EUK-134 studies For EUK-134 studies neurons were either pretreated for 1 h or coincubated with 5 ␮M EUK-134 followed by the addition of either DETA-NO• (1000 ␮M) or NMDA (100 ␮M) for 3 h. All groups consisted of two to three coverslips of neurons and analysis was performed using the microscope method described in the cyto-c analysis section (above) except analysis of each group was performed on 50 separate digital images (approximately 100 cells total). EUK-134 was a kind gift from Eukarion, Inc. (Bedford, MA). Electrophysiological studies Patch electrodes were pulled on a horizontal FlamingBrown microelectrode puller; resistances were 4 – 8 M⍀. Electrodes were filled with a sterile-filtered internal recording solution of (in mM) potassium gluconate, (145), MgCl2 (2.0), N-[2-hydroxyethyl]piperazine-N⬘-[2-ethansulfonic acid acid] (Hepes) (10.0), ethylene glycol-bis(␤-aminoethyl ether) N,N,N⬘,N⬘-tetraacetic acid (EGTA) (5.0), and dipotassium ATP (3.0), pH 7.3, osmolarity 275–280 mOsm. Prior to recording, coverslips containing cortical neurons were placed in a polystyrene culture dish with sterile filtered external recording solution containing NaCl (145), CaCl2 (1.0), KCl (5.0), glucose (10.0), Hepes (10.0), and glycine (0.01), pH 7.4. The osmolarity was adjusted to 320 –325 mOsm with sucrose. Whole cell recordings were made at RT with an Axopatch 1-D patch clamp amplifier (Axon Instruments, Union City, CA) and low-pass filtered at 2 kHz and recorded at 10 –20 kHz. Data were recorded to a personal computer with Axoscope 7.0 data acquisition software using a Digidata 1200 interface (Axon Instruments). NMethyl-D-aspartate (10 ␮M) was applied using a modified T-tube drug application device (Greenfield and Macdonald, 1996). Statistics Statistical analysis of TMRM fluorescence changes compared to control and/or of survival data was performed by repeated measures ANOVA, ␹2 analysis of contingency tables, or a Student t test if individual data time points were being compared to control; multiple linear regression analysis was also performed (SigmaStat Ver. 2.0, SPSS).

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Acknowledgments This study has been supported in part by NIH-NINDS Grant NS01857 and a grant from the American Federation of Aging Research (AFAR) to NJS. The authors are grateful to Cassie Gregory and Ashley Renick for the preparation and maintenance of cortical neuronal cultures and to Drs. Jaideep Kapur and Patrick Mangan for their electrophysiological expertise.

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