RESULTS

NMDAR overstimulation induces neuronal cell death

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Articles by Montal, M.  Previous Article  |  Next Article  Volume 16, Number 19, Issue of October 1, 1996 pp. 6125-6133 Copyright ©1996 Society for Neuroscience Mitochondrial Dysfunction Is a Primary Event in Glutamate Neurotoxicity Alejandro F. Schinder1, Eric C. Olson1, Nicholas C. Spitzer1, 2, and Mauricio Montal1 1 Department of Biology and 2 Center for Molecular Genetics, University of California at San Diego, La Jolla, California 92093-0366 ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION FOOTNOTES REFERENCES ABSTRACT Excitotoxic neuronal death, associated with neurodegenerative disorders and hypoxic insults, results from excessive exposure to excitatory neurotransmitters. Glutamate neurotoxicity is triggered primarily by massive Ca2+ influx arising from overstimulation of the NMDA subtype of glutamate receptors. The underlying mechanisms, however, remain elusive. We have tested the hypothesis that mitochondria are primary targets in excitotoxicity by confocal imaging of intracellular Ca2+ ([Ca2+]i) and mitochondrial membrane potential () on cultured rat hippocampal neurons. Sustained activation of NMDA receptors (20 min) elicits reversible elevation of [Ca2+]i. Longer activation (50 min) renders elevation of [Ca2+]i irreversible (Ca2+ overload). Susceptibility to NMDA-induced Ca2+ overload is increased when the 20 min stimuli are applied to neurons pretreated with electron transport chain inhibitors, thereby implicating mitochondria in [Ca2+]i homeostasis during excitotoxic challenges. Remarkably, exhibits prominent and persistent depolarization in response to NMDA, which closely parallels the incidence of neuronal death. Blockade of the mitochondrial permeability transition pore by cyclosporin A allows complete recovery of and prevents cell death. These results suggest that early mitochondrial damage plays a key role in induction of glutamate neurotoxicity. Key words: calcium; cyclosporin; excitotoxicity; imaging; mitochondria; NMDA receptor; neuronal death INTRODUCTION Excitotoxicity, the process by which overactivation of excitatory neurotransmitter receptors leads to neuronal cell death, must be understood in terms of the intervening cascade of events (Choi, 1987, 1988, 1995). Compelling evidence supports a key role for massive Ca2+ influx through the NMDA receptor (NMDAR) channel as a trigger of glutamate neurotoxicity: (1) selective blockade of NMDARs markedly reduces the extent of excitotoxic damage (Choi et al., 1988; Michaels and Rothman, 1990); (2) neurons survive when challenged in absence of extracellular Ca2+ or presence of intracellular Ca2+ ([Ca2+]i) chelators (Choi, 1987; Tymianski et al., 1993a,b); and (3) glutamate receptor overstimulation induces elevations of [Ca2+]i that precede cell death (de Erausquin et al., 1990; Michaels and Rothman, 1990; Randall and Thayer, 1992; Tymianski et al., 1993a,b; Witt et al., 1994). Oxidative stress is thought to be a contributing factor in glutamate neurotoxicity through generation of oxygen radicals that disrupt cell functions (Dykens et al., 1987; Dawson et al., 1991; Coyle and Puttfarcken, 1993; Lafon-Cazal et al., 1993; Lipton et al., 1993). However, the link between elevation of [Ca2+]i, free radicals, and neuronal death remains unclear (Choi, 1995). Mitochondria are unique among cell organelles in their involvement in the concerted consumption of oxygen, production of oxygen radicals, and mobilization of [Ca2+]i (Gunter and Pfeiffer, 1990; Thayer and Miller, 1990; Miller, 1991; Beal, 1992; Coyle and Puttfarcken, 1993; Rizzuto et al., 1993; Jouaville et al., 1995; White and Reynolds, 1995). Given that excessive Ca2+ accumulation in mitochondria uncouples electron transfer from ATP synthesis (Beatrice et al., 1980; Gunter and Pfeiffer, 1990; Bernardi et al., 1994), and considering that impairment of energy metabolism (Wang et al., 1994) increases generation of free radicals (Halliwell and Gutteridge, 1989; Coyle and Puttfarcken, 1993), mitochondria emerge as a plausible link between elevation of [Ca2+]i and glutamate neurotoxicity. The pivotal role of mitochondria in excitotoxicity has been highlighted recently by the deterministic influence of mitochondrial function in the decision between apoptotic or necrotic cell death (Ankarcrona et al., 1995; Bonfoco et al., 1995) and by the protective action of transforming growth factors presumably conferred by enhancing mitochondrial energetics (Mattson et al., 1993; Prehn et al., 1994). Here, we focus on the following question: is the mitochondrion the sensor that converts elevation of [Ca2+]i from a physiological modulator into a trigger for cell death? We show that mitochondria are involved in Ca2+ sequestration during an excitotoxic insult (Schinder et al., 1995) and that this Ca2+ current induces sustained mitochondrial depolarization, which closely parallels the incidence of neuronal death. Our findings suggest that neuronal injury follows from excessive influx of Ca2+ via the NMDAR that, in turn, attenuates the mitochondrial membrane potential () leading to the opening of the permeability transition pore (PTP), further collapsing . Blockade of the PTP with cyclosporin A (CsA) leads to the recovery of and promotes cell survival. The results outline a novel way of thinking about excitotoxic cell death, shifting the focus of attention from a global event staged by intracellular Ca2+ toward a more compartmentalized question centered on mitochondrial homeostasis. MATERIALS AND METHODS Hippocampal cultures. Mixed hippocampal neuronal/glial cultures were established as described (Schinder and Montal, 1993). Briefly, hippocampi from E17-E19 rat embryos were incubated in basic saline solution (BSS) containing (in mM): 137 NaCl, 3.5 KCl, 0.4 KH2PO4, 0.33 Na2HPO4 · 7 H2O, 5 trimethylaminoethanesulfonic acid, pH 7.4, and 10 glucose with 0.25% trypsin for 15 min at 37°C. Tissue was washed and dissociated by trituration with a Pasteur pipette. Cells were centrifuged for 5 min at 200 × g, and the pellet was resuspended in BSS. Cells were plated (105 cells/cm2) and maintained as described. Cell death assays. In vitro excitotoxicity is commonly investigated using agonist exposures that range from 5 min to 24 hr (Choi, 1987; Weiss et al., 1994). In our conditions, mild and severe types of neuronal death were selectively achieved with exposures of 20 and 50 min, respectively. Culture medium was removed, neurons were rinsed with 10 ml of BSS + 1.1 mM CaCl2 + 10 µM glycine, and treated with 200 µM NMDA (dissolved in the same solution) for either 20 (de Erausquin et al., 1990; Michaels and Rothman, 1990; Lafon-Cazal et al., 1993; Witt et al., 1994) or 50 min (Tymianski et al., 1993a,b) at 23 ± 1°C. Treatments were terminated by addition of culture medium (Schinder and Montal, 1993) supplemented with 20 µM dizolcipine [MK-801] (Research Biochemicals, Natick, MA) to prevent activation of NMDAR attributable to residual NMDA or glutamate released from synapses (Hartley and Choi, 1989). Cultures were returned to the incubator and cell death was assessed 18-24 hr later by trypan blue staining (0.4%). When indicated, CsA (Sandoz, Basel, Switzerland) was dissolved in ethanol or dimethyl sulfoxide (DMSO; 42 mM) and added to solutions held at 37°C under vigorous stirring to a final concentration of 1.6 µM. To mimic those conditions used in imaging experiments, CsA was added 40 min before the NMDA challenge and removed after 2 hr. A minimum of 500 cells were counted blind on each culture dish. For each experiment, two identical culture dishes were treated for control and 20 and 50 min exposure to NMDA, or, in the CsA experiments, control and 20 min NMDA in absence and presence of CsA. Calcium imaging. Neurons were Fluo-3-loaded for 75 min at a bath concentration of 5.5 µM Fluo-3 AM (Molecular Probes, Eugene, OR) dissolved in DMSO (Kao et al., 1989). Imaging was performed with an argon laser confocal system (Bio-Rad MRC600) equipped with 488 nm excitation/515 nm emission filters on a Zeiss microscope (Thornwood, NY) with a 20× water immersion objective (numerical aperture 0.4). Neurons were constantly superfused at a rate of 3 ml/min with BSS + 1.1 mM CaCl2 + 2 mM MgCl2 + 10 µM glycine (control solution). NMDA was applied in Mg2+-free solution. Images were sampled and collected at 8-15 sec intervals for 2 hr (up to 4 hr for 50 min NMDA experiments) and recorded on videotape. Slower sampling rates (1 image/2 min) and lower laser intensity (1/6 of standard power) yielded identical results, suggesting that phototoxic effects are minimal. Signals obtained during Ca2+ overload were not attributable to saturation of dye, as fluorescence increased further after addition of 5 µM ionomycin (Calbiochem, La Jolla, CA) in the presence of 20 mM CaCl2, pH 8.0. Digital images were acquired either on-line or off-line with a Macintosh computer equipped with a frame grabber (Data Translation) using Image 1.47 software (W. Rasband, National Institutes of Health). A total of 20-25 neurons was studied in each experiment; somatic regions of ~170 pixels were chosen for quantitation of fluorescence intensity. Regions without neurons or processes were used for background subtraction. For calibration experiments (Kao et al., 1989; Vergara and DiFranco, 1992), [Ca2+]i was calculated as [Ca2+] = Kd(F  Fmin)/(Fmax  F), where Kd was assumed to be 400 nM, Fmax was measured at the end of the experiment by treating neurons with 5 µM ionomycin, Fmin was derived from Fmax, and background fluorescence was obtained after addition of 0.02% saponin. Imaging of . Neurons were incubated for 30 min in the presence of 0.1 µM tetramethylrhodamine ethylester (TMRE; Molecular Probes). TMRE was added to all solutions used during these experiments, which typically lasted 2 hr. When indicated, 1.6 µM CsA was present in all solutions throughout the experiment, with a preincubation time of 40 min before the NMDA challenge. Three to seven neurons were studied in each experiment. Imaging (Farkas et al., 1989) was as described above, except that samples were collected at 1 image/90 sec and a 514/550 nm filter and a 40× water immersion objective (numerical aperture 0.9) were used. The confocal pinhole was closed to 1/5 to 1/3, and signals were amplified 1.5× with a digital zoom. Fluorescence intensity was quantified in a perinuclear ring (~1000 pixels) in which mitochondrial clusters are located (see Fig. 4B). Baseline fluorescence (F0) was measured as average normalized fluorescence emitted during the initial 5 min of the experiment. Control experiments showed that selective depolarization of the plasma membrane (35 mM KCl, 0 Ca2+) evoked minor changes in mitochondrial fluorescence. Results indicate that decrements of mitochondrial fluorescence primarily reflect mitochondrial depolarization and not reequilibration of dye in response to NMDA-induced plasma membrane depolarization. This assertion was confirmed by the finding that 100 µM carbonyl cyanide m-chlorophenylhydrazone, uncoupler of oxidative phosphorylation, decreases TMRE fluorescence (Farkas et al., 1989) to an extent comparable to that produced by the NMDA treatments. Fig. 4. NMDAR overstimulation collapses . A, Digital images of neurons exposed to TMRE before removal of NMDA (top frame indicated by 0:00:00 time), during 20 min stimulation with 200 µM NMDA, and after removal of NMDA (wash). Collapse of is shown as a decrease in fluorescence intensity during exposure to NMDA. Partial repolarization occurs after the stimulus is removed. Pseudocolor scale represents arbitrary fluorescence intensity values ranging from 0 to 255. Scale bar, 20 µm. B, Time course of measured in arbitrary fluorescence units for neurons stimulated with NMDA for 0 (control), 0.5, 20, and 50 min and recorded during 2 hr. Each trace represents mitochondrial fluorescence from an individual neuron normalized to its initial baseline value (F0), and each set of recordings belongs to a single representative experiment. Inset, Perinuclear ring of mitochondrial fluorescence used for quantitation. C, Mitochondrial fluorescence signals, as shown in B, are quantified as Peak Depolarization = Fmin/F0, an indicator of transient mitochondrial depolarization, and Recovery Ratio = Fend/F0, an indicator of mitochondrial repolarization after cessation of the NMDA stimulus. Bars plotted for 0, 0.5, 20, and 50 min NMDA represent mean ± SEM for n = 28, 28, 74, and 84, respectively. For both bar charts, ANOVA analysis revealed that populations are different with p < 0.00001. **, Statistically significant differences (p < 0.0001) compared with all other treatments using the post hoc test. D, Effect of [Ca2+]ext on depolarization of elicited by 20 min NMDA. Bar chart for Peak Depolarization and Recovery Ratio was calculated and plotted as in C. *, Statistically significant difference with p < 0.002. Inset, Comparison of the time courses of in the presence of [Ca2+]ext = 1.1 or 5.5 mM. Traces are averages from all recordings (5.5, n = 33; 1.1, n = 74). [View Larger Version of this Image (50K GIF file)] All experiments reported were performed at 23 ± 1°C on neurons cultured for 14-17 d. Statistics. Pooled data are presented as mean ± SEM. For comparisons between two groups, a two-tailed Student's t test was used. When three or more groups were compared (Figs. 1 [Ca2+]i (4) followed by a slow decrease to a lower level lasting several minutes (5), which gradually increases to achieve a higher plateau [Fplateau, (6)]. [Ca2+]i returns to baseline levels shortly after the NMDA stimulus is removed (7). Record obtained from the neuron marked with an arrowhead. Scale bar, 40 µm. B, Same as A, but neurons were stimulated for 50 min with NMDA. A similar time course was recorded, but heightened [Ca2+]i (6) failed to return to baseline levels [Ca2+ overload, (7)]. Calibration of [Ca2+]i yielded the following values: (1) [Ca2+]baseline = 29 ± 2 nM; (2) [Ca2+]K+ = 0.91 ± 0.10 µM; (4) [Ca2+]peak = 1.5 ± 0.2 µM; (5) [Ca2+]valley = 0.29 ± 0.03 µM; (6) [Ca2+]plateau = 2.1 ± 0.2 µM; n (number of cells) > 47. C, Extent of Ca2+ overload versus duration of the NMDA challenge. Ca2+ overload was calculated as the ratio of the fluorescence value at the end of the experiment [Fend, (7)] to the highest value achieved during the challenge [Fplateau, (6)]. Data are mean ± SEM. ANOVA analysis revealed that populations are different with p < 0.00001. *, Statistically significant difference (p < 0.005) obtained using the post hoc test compared with all other treatments; **p < 0.0001. Numbers in parentheses denote population sizes. {/ANNT;64064n;;66880n;72864n}

NMDA produces transient or sustained increases in [Ca²⁺]_i depending on pulse duration

The susceptibility to NMDA-induced Ca²⁺ overload is augmented by mitochondrial function inhibitors

Effects of mitochondrial function inhibitors on 

CsA

NMDA collapses 

Blockade of the mitochondrial PTP with cyclosporin A promotes recovery of the NMDA-induced collapse of and prevents neuronal death

DISCUSSION

Time-lapse video imaging of shows that depolarization is a consequence of NMDAR overstimulation (Fig. 4); whereas mitochondrial depolarization evoked by 20 min NMDA does not revert to baseline levels (Fig. 4B), elevation of [Ca²⁺]_i does (Fig. 1, Table 1). These results have three major implications: (1) sustained mitochondrial depolarization is not attributable to tonically elevated [Ca²⁺]_i; (2) mitochondrial homeostasis is not linked to [Ca²⁺]_i after removal of the excitotoxic stimulus; and (3) persistent mitochondrial depolarization is an expression of mitochondrial dysfunction.

Findings on cell death, [Ca²⁺]_i homeostasis, and are summarized in Table 1. Results from 20 min NMDA treatments reveal an apparent mismatch between the extent of Ca²⁺ overload and the percentage of dead neurons. This suggests that although existing ATP pools may be sufficient to drive Ca²⁺ extrusion systems and recover [Ca²⁺]_i homeostasis (Fig. 1A,C), early neuronal damage occurs as revealed by the collapse of (Fig. 4) and the induction of cell death. In contrast, the extent of Ca²⁺ overload observed for 50 min treatments (Fig. 1B) parallels the incidence of neuronal death, suggesting exhausted ATP pools. Ca²⁺ overload therefore may be secondary to mitochondrial dysfunction, and not a causal event for cell death (Fig. 6). This conjecture is supported by the results presented in Figure 2, in which inhibition of mitochondrial function leads to Ca²⁺ overload; for instance, [Ca²⁺]_i responses to 50 min NMDA are mimicked by 20 min NMDA in neurons pretreated with oligomycin. Moreover, in agreement with previous findings (Tymianski et al., 1993a), Ca²⁺ overload occurs much earlier than loss of plasma membrane integrity, as indicated by the ability of neurons to retain the Ca²⁺ indicator Fluo-3 in the cytoplasm for more than 4 hr. Thus, Ca²⁺ overload may be an indicator of cell death, but recovery is not an indicator of cell survival. This notion resolves discrepancies in correlations of [Ca²⁺]_i postinsult and neuronal death (de Erausquin et al., 1990; Michaels and Rothman, 1990; Randall and Thayer, 1992; Witt et al., 1994).

Mitochondria are major buffering compartments for elevations of [Ca²⁺]_i in response to brief exposures to glutamate in cortical neurons (White and Reynolds, 1995). Inhibition of mitochondrial function by picomolar concentrations of antimycin A or rotenone abolishes Ca²⁺ sequestration in response to prolonged NMDA treatments (Fig. 2), indicating that mitochondria act as a primary buffer of [Ca²⁺]_i also under excessive insults. A unique function of mitochondria (as a main source of ATP) is highlighted by the finding that oligomycin or higher concentrations of antimycin A or rotenone eliminate the recovery phase, presumably by compromising ATP-dependent Ca²⁺ extrusion systems (Fig. 6). Thus, mitochondria, as intracellular Ca²⁺ buffers, set a point of vulnerability whereby if the boundary between mitochondrial function and dysfunction is crossed, cell death ensues. As discussed below, this crossroad is probably determined by the PTP (Fig. 5).

Our population study suggests a correlation between mitochondrial dysfunction and cell death, but does not establish a causal link (Table 1). If mitochondrial impairment were a primary event in the induction of excitotoxic cell death, then neuronal death might be prevented by treatments that protect mitochondrial homeostasis. As shown in Figure 5, the NMDA-induced collapse of recovers completely in the presence of CsA. Accordingly, neuronal death is reduced by 65% under these conditions compared with neurons challenged in the absence of CsA (Fig. 5C). NMDARs are not a target of CsA, because voltage-clamp recordings show that currents elicited by NMDA remain unchanged in the presence of CsA (data not shown). The fact that complete recovery of is accompanied by a substantial, yet partial, reduction of cell death might indicate that pathways that are parallel and/or downstream to mitochondria are also activated by NMDAR overstimulation (Fig. 6). Immunosuppressant drugs, such as CsA, display numerous intracellular binding sites (Kunz and Hall, 1993) and have been shown to reduce excitotoxic neuronal death in cortical neurons, presumably by inhibition of a pathway that activates nitric oxide synthase (NOS) (Dawson et al., 1993). However, because CsA is a potent blocker of the PTP (Petronilli et al., 1993; Bernardi et al., 1994; Nicolli et al., 1996) and because our results show a direct effect of CsA on mitochondrial (Fig. 5A,B), it appears unlikely that preservation of mitochondrial function and promotion of cell survival are attributable to inhibition of NOS, yet this cannot be ruled out. Our findings suggest that persistent mitochondrial depolarization results from opening of the PTP [presumably as a consequence of combined mitochondrial depolarization and elevation of intramitochondrial Ca²⁺ (Bernardi et al., 1994)] and that maintenance of mitochondrial homeostasis can protect hippocampal neurons from excitotoxic cell death (Fig. 6).

In view of these findings, glutamate excitotoxicity can be visualized as a cascade of events that starts with overstimulation of NMDARs, which elicit massive Ca²⁺ influx and abnormally high [Ca²⁺]_i (Fig. 6). Ca²⁺ is sequestered into the mitochondrial matrix, driven by the proton electrochemical gradient generated by the electron transport chain, depolarizing (Akerman, 1978; Gunter and Pfeiffer, 1990; Loew et al., 1994). This reduction in the electrochemical gradient decreases ATP synthesis at a time of great demand by the plasma membrane Ca²⁺ pump and, indirectly, by the Na⁺/Ca²⁺ exchanger (White and Reynolds, 1995). Concurrent buildup of intramitochondrial [Ca²⁺] and attenuation of induce opening of the mitochondrial inner membrane PTP, thereby allowing ionic diffusion and collapsing (Beatrice et al., 1980; Gunter and Pfeiffer, 1990; Bernardi et al., 1994; Hoek et al., 1995). The drop of cellular ATP levels, in concert with the generation of free O radicals consequent to high ATP consumption (Halliwell and Gutteridge, 1989; Coyle and Puttfarcken, 1993; Reynolds and Hastings, 1995), is a primary cause of cell death (Siesjö, 1992). The boundary between cell survival and cell death relies on a subtle balance of mitochondrial function/dysfunction, centered on . This model predicts that neuronal lifespan may be prolonged by specific inhibition of mitochondrial Ca²⁺ uptake and that susceptibility to excitotoxic death may be influenced by the energetic state of neurons before and during the insult.

Our findings establish a direct coupling between excitotoxicity and shifts of redox potentials and membrane potentials of mitochondria inside living neurons. This evidence strengthens the involvement of mitochondria in the neurotoxicity associated with mitochondrial complex I inhibitor 1-methyl-4-phenylpyridinium (Saporito et al., 1992), amyotrophic lateral sclerosis (Brown, 1995; Wong et al., 1995), apoptosis (Newmeyer et al., 1994; Vayssière et al., 1994; Zamzami et al., 1995), and age-related disorders (Luft, 1994; Brouillet et al., 1995). These findings may provide useful guidelines for the development of the next generation of drugs targeted against neurodegeneration and stroke.

FOOTNOTES

Received May 15, 1996; revised July 17, 1996; accepted July 19, 1996.

This work was supported by grants from National Institutes of Health (GM-49711 to M.M., NS-15918 to N.C.S.) and the U.S. Army Medical Research and Material Command (DAMD 17-93-C-3100 to M.M.). We are indebted to P. Bernardi, F. H. Gage, Y. Mika, C. D. Patten, J. L. Vergara, G. Zlokarnik, and the members of the Montal lab for valuable discussions and comments. We thank Drs. D. Römer, E. Rissi, G. Engel, and H. Widmer from Sandoz (Basel, Switzerland) for kindly providing CsA.

Correspondence should be addressed to Mauricio Montal, Department of Biology, University of California at San Diego, 9500 Gilman Drive, La Jolla, CA 92093-0366.

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