Introduction

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Excitotoxic mechanisms (i.e., excessive stimulation of glutamate receptors with a resultant disturbance of cellular [Ca²⁺]_i homeostasis) may be involved in stroke and possibly in slowly developing neurodegenerative diseases (Choi and Rothman, 1990). However, glutamate receptor stimulation is rarely a primary event in neurotoxicity. Glutamate release and glutamate-triggered excitotoxicity are rather secondary consequences of other defects or metabolic disturbances. A frequent initiating condition is energy depletion, and mitochondrial dysfunction is among the most generalized causes favoring the development of different neurodegenerative diseases (Beal, 1996). For example, Huntington's disease is modeled by exposing specific neuronal subpopulations to mitochondrial toxins (Ferrante et al., 1997), and ischemic damage can be examined in vitro after mitochondrial substrate depletion due to oxygen/glucose deprivation (Choi and Rothman, 1990).

In these models, a close relationship has been established between energy deficiency and excitotoxicity. The ATP loss resulting from decreased mitochondrial energy generation would lead to impaired function of ion pumps and partial hypopolarization of neurons, thereby releasing the voltage-dependent Mg²⁺ block of the NMDA-R-gated Ca²⁺ channel. This would make the receptor/channel hypersensitive to glutamate stimulation. NMDA-R-mediated influx of Na⁺ and Ca²⁺ then would increase energy demand and ATP depletion, enhance depolarization, trigger further [Ca²⁺]_i increase, and eventually result in further glutamate release. This putatively self-propagating process finally leading to a loss of [Ca²⁺]_i homeostasis and excitotoxicity has been termed the "energy-linked excitotoxicity hypothesis" (Henneberry et al., 1989; Zeevalk and Nicklas, 1990). Distal mechanisms (downstream of [Ca²⁺]_i increase) leading to neuronal death under such conditions may differ from those operating after simple glutamate receptor stimulation and are largely unknown.

Excitotoxicity may lead to either necrosis or to a more ordered sequence of molecular events resulting in apoptosis (Ankarcrona et al., 1995; Leist and Nicotera, 1998). Recent work in our laboratory has shown caspase activation downstream to the initial [Ca²⁺]_i increase in a model of indirect excitotoxicity, where NMDA-R activation was triggered by NO (Leist et al., 1997a, 1997d). In parallel, studies in vivo have suggested that caspases may be involved in neuronal damage after stroke (Loddick et al., 1996). In all these models, the exact primary insult is little characterized and possibly not limited to mitochondrial dysfunction; therefore, it is still unclear how a primary mitochondrial impairment can lead to excitotoxic apoptosis.

The neurotoxin MPTP induces a Parkinson's disease-like syndrome in humans and animals via its active metabolite, MPP⁺ (Tipton and Singer, 1993). Although MPP⁺ affects only certain neurons in vivo due to pharmacokinetic reason, the substance inhibits mitochondrial function in any cultured neuronal or non-neuronal cell or isolated mitochondria. A known molecular target of MPP⁺ is the mitochondrial respiratory chain complex I (NADH-ubiquinone-oxidoreductase) (Nicklas et al., 1985; Kilbourn et al., 1997), and no other main target has been characterized to date, despite extensive studies. In vivo (i.e., in the presence of glutamatergic cortical inputs), the primary impairment of mitochondrial respiration by MPP⁺ sensitizes neurons to secondary excitotoxic damage. Thus, blockade of glutamate receptors (Turski et al., 1991; Srivastava et al., 1993) or prevention of the synthesis of endogenous NO (Schulz et al., 1995; Hantraye et al., 1996), a mediator closely linked to excitotoxicity, has been found to reduce MPP⁺ toxicity in animals.

In the current study, we exposed neurons to the mitochondrial poison MPP⁺ or other respiratory chain inhibitors. CGC have been shown to be susceptible to MPP⁺ (Marini et al., 1989). Thus, we used here neuronal cultures, which mimic the very high density of excitable glutamatergic synapses present in vivo. Densely plated CGC, differentiated for 8 days in K⁺-supplemented medium, represent an ideal culture system for this purpose. We examined in this culture system (i) whether excitotoxic mechanisms contribute to MPP⁺ toxicity, (ii) whether such excitotoxicity results in apoptosis, (iii) under which conditions the execution of excitotoxic death triggered by MPP⁺ or other mitochondrial toxins involves caspases, and (iv) which apoptotic features depend on the activity of protease families activated by excitotoxic disruption of [Ca²⁺]_i homeostasis.

	Experimental Procedures

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Materials. Fluo 3-acetoxymethyl ester, Fura 2-acetoxymethyl ester, calcein acetoxymethyl ester, TMRE, SYTOX, ethidium homodimer-1, and H-33342 were obtained from Molecular Probes (Eugene, OR). The caspase substrate DEVD-afc, MPP⁺, DCK, DNQX, and 6,7-dichloroquinoxaline-2,3-dione were obtained from BIOMOL (Hamburg, Germany). MK801 came from RBI (Biotrend Chemikalien GmbH, Köln, Germany). succ-LLVY-amc; calpain inhibitors I, II, and III (Ac-Leu-Leu-L-norleucinal, Ac-Leu-Leu-L-methional, and calp III, z-Phe-chloromethylketone (cmk); and the caspase-inhibitors DEVD-CHO, z-VAD-fluoromethylketone (fmk), Ac-YVAD-cmk, or Ac-YVAD-2,6-dimethylbenzoyloxymethylketone (ICE II) and z-D-2,6-dichlorobenzoyloxymethylketone (cbk) (ICE III) were obtained from Bachem Biochemica (Heidelberg, Germany). DEVD-fmk was from Enzyme Systems (Dublin, CA). Fluorescein-labeled annexin V (annexin V) was from Boehringer-Mannheim (Mannheim, Germany). Clostridial toxins were generously supplied by Dr. C. Montecucco (University of Padova, Padova, Italy). Solvents and inorganic salts were from Merck (Darmstadt, Germany) or Riedel-de Haen (Seelze, Germany). All other reagents not further specified were from Sigma (Deisenhofen, Germany).

Animals. PARP^/ mice (C57Bl/6 × 129/Sv background) or corresponding wild-type animals were generously provided by Dr. Zhao-Qi Wang (IARC, Lyon, France) (Wang et al., 1995). All animals used for cell preparations were typed by Southern blotting (Wang et al., 1995) to verify the genotype. For other experiments, 8-day-old specific pathogen-free BALB/c mice were obtained from the Animal Unit of the University of Konstanz. All experiments were performed in accordance with international guidelines to minimize pain and discomfort (National Institutes of Health guidelines and European Community Council Directive 86/609/EEC).

Cell culture. Murine CGC were prepared as described previously (Schousboe et al., 1989). Neurons were plated onto 100 µg/ml (250 µg/ml for glass surfaces) poly-L-lysine (molecular mass, >300 kDa)-coated dishes at a density of ~0.25 × 10⁶ cells/cm² (800,000 cells/ml; 500 µl/well; 24-well plate) and cultured in Eagle's basal medium (GIBCO, Paisley, Scotland) supplemented with 10% heat-inactivated fetal calf serum, 20 mM KCl, 2 mM L-glutamine, 100 units/ml penicillin, and 100 µg/ml streptomycin. Forty-eight hours after plating, cytosine arabinoside (10 µM) was added to the cultures. Neurons were routinely used at 8-10 DIV unless otherwise indicated. Glial fibrillary acid protein-positive cells were <5%.

Cytotoxicity assays. Cultures were exposed to MPP⁺ or rotenone in their own medium. The culture medium was exchanged for a CSS (120 mM NaCl, 25 mM HEPES, 25 mM KCl, 1.8 mM CaCl₂, 4 mM MgCl₂) plus 15 mM glucose 4 hr after the start of the incubation, and the cells were left in this medium until toxicity parameters were read (usually 18 hr for MTT or nuclear morphology). Inhibitors were added routinely 30 min before exposure to MPP⁺.

Ca²⁺ measurements. The [Ca²⁺]_i was measured by imaging neurons loaded with fluorescent Ca²⁺ indicators. To monitor dynamic changes of Ca²⁺, CGC were loaded in their original medium with 1 µM fluo-3 acetoxymethyl ester for 10 min at 37°. The medium then was exchanged for CSS, in which CGC were incubated for 5 min to allow complete de-esterification of fluo-3. The medium was exchanged again for the original neuron-conditioned complete Eagle's basal medium supplemented with 20 mM HEPES (MPP⁺ as stimulus) or for CSS (without Mg²⁺) (50 µM glutamate or 200 µM NMDA as stimulus) or for CSS plus 2 µM MK801 (200 µM kainate as stimulus). CGC were allowed to equilibrate at room temperature for 10 min before the exposure. Images were collected using the 488-nm excitation and 520-nm emission wavelengths and a Leica DM-IRB microscope equipped with a 40× NA 1.0 lens. Data from 10-20 neurons were recorded at 3-60-sec intervals with a Leica TCS 4D confocal system. Relative mean fluorescence levels from defined areas corresponding to the position of neuronal cell bodies were recorded over the time course of the experiment, and the values were arbitrarily set to 1 at the beginning of each experiment. To determine absolute [Ca²⁺]_i, we used the low K_d indicator Fura 2-acetoxymethyl ester (2.5 µM; Molecular Probes, Eugene, OR), which reports [Ca²⁺]_i exactly in the low concentration range (500 nM). This allowed us to examine whether MK801 would maintain [Ca²⁺]_i to base-line levels in MPP⁺-treated cells. A Leica M-IRB microscope equipped with a computer-controlled filter wheel (Sutter, Novato, CA), quartz optics, and a Dage-72 (Dage-MTI; Michigan City, IN) charge-coupled device (CCD) camera [756 (H) × 581 (V) pixels] coupled to a videoscope GEN-III image intensifier was used for imaging (_ex-1 = 340 nm, _ex-2 = 380 nm, _em = 505 nm). Videomicroscopy data were analyzed using software from Imaging Research (St. Catherine's, Ontario, Canada). [Ca²⁺]_i was determined by in situ calibration using the equation [Ca²⁺]_i = K_d × (R  R_min)/(R_max  R) × S_f2/S_b2, with K_d(25°) = 264 nM. To determine R_min, cells were washed twice with calibration buffer (120 mM NaCl, 25 mM HEPES, 15 mM glucose, 25 mM KCl, 2 mM MgCl₂, 2 mM EGTA) and equilibrated for 20 min in calibration buffer supplemented with 5 µM ionomycin. Ca²⁺ (5 mM) and ionomycin (10 µM) were added to saturate Fura-2 with Ca²⁺ and calculate R_max. Autofluorescence was measured after the addition of 5 mM MnCl₂.

Electrophoretic assays. Fodrin proteolysis was analyzed by immunoblot. CGC cultures were lysed in RIPA buffer (150 mM NaCl, 50 mM Tris, 1% Nonidet P-40, 0.25% sodium deoxycholate, 1 mM EGTA) supplemented with protease inhibitors (1 mM phenylmethylsulfonyl fluoride, 1 mM benzamidine, 1 mM iodoacetate, 1 mM iodoacetamide, 40 µM leupeptin, 10 µg/ml antipain, 5 µg/ml pepstatin). Before lysis, cultures were stained with 0.5 µM SYTOX to control the percentage of cells with intact membranes, which was >95% for all samples analyzed. Protein was determined using the bicinchoninic acid method (BioRad, München, Germany), and 5 µg of protein/lane was loaded onto 8% polyacrylamide gels. Proteins were separated under reducing conditions at 60 mA and then blotted onot a nitrocellulose membrane (Amersham-Buchler, Braunschweig, Germany) in a BioRad semidry blotter at 2.6 mA/cm² for 50 min using a Towbin buffer system. Blots were blocked for 1 hr and then incubated with anti-fodrin monoclonal antibody (clone 1622, 1:1000) from Chemicon (Temecula, CA) diluted in TNT (150 mM NaCl, 50 mM Tris, pH 8.0, 0.05% Tween 20) for 1 hr at room temperature. Specifically stained bands were detected by enhanced chemiluminescence (Amersham) using a peroxidase-coupled secondary antibody. For Western blot analysis of procaspase-3 (14% gel, 60 µg of neuronal protein/lane), we used a primary rabbit anti-human procaspase-3 polyclonal antibody (1:1000, no. 06753; Upstate Biotechnology, Lake Placid, NY) recognizing the 32-kDa murine procaspase but not the cleavage products.

Mitochondrial function and integrity. ATP was measured luminometrically after lysis of cells in ATP-releasing agent (Sigma) with a commercial kit (Boehringer-Mannheim) as described previously (Leist et al., 1997c). The was monitored by loading cells with the fluorescent indicator TMRE (5 nM: _ex, 568 nm; _em, 590 nm). Under these conditions, fluorescence completely disappeared on loss of (50 µM glutamate or 50 µM CCCP) in neurons that retained plasma membrane integrity. For the experiments, at least three culture dishes were imaged in at least six cell preparations for each data point presented. For semiquantitative analysis of , we added red fluorescent beads of 2-µm diameter and calibrated fluorescence intensity (Molecular Probes, Eugene, OR) to the cultures. The standardized fluorescence of the beads was used as an internal standard to normalize the fluorescence intensities of digitized images of TMRE-loaded neurons. The fluorescence of neurons treated with 20 µM CCCP was used as reference for depolarized mitochondria.

Enzymatic assays. Caspase-3-like activity (measured by DEVD-afc cleavage) was assayed as described previously (Leist et al., 1997b) with the following modifications: CGC were pelleted in PBS supplemented with 5 mM EDTA, 1 µg/ml leupeptin, 1 µg/ml pepstatin, 1 µg/ml aprotinin, and 1 mM PEFA-block. Lysis was performed in 25 mM HEPES, 5 mM MgCl₂, 1 mM EGTA, 0.5% Triton X-100, 1 µg/ml leupeptin, 1 µg/ml pepstatin, 1 µg/ml aprotinin, and 1 mM PEFA-block, pH 7.5. The fluorimetric assay was carried out in microtiter plates with a substrate concentration of 40 µM and a total protein amount of 5 µg. Cleavage of DEVD-afc was followed in reaction buffer (50 mM HEPES, 10 mM dithiothreitol, 1% sucrose, 0.1% 3-[(3-cholamidopropyl)dimethylammonio]propanesulfonate) over a period of 30 min at 37° with _ex = 390 nm and _em = 505 nm, and the activity was calibrated with afc-standard solutions. Calpain activity was determined by a kinetic fluorimetric assay as described previously (Leist et al., 1997d). Glucose concentrations in the medium were determined according to the hexokinase/glucose dehydrogenase method using a commercial kit (Sigma).

Visualization of PS translocation. Cells stained with different fluorescent probes were imaged on a Leica DM-IRBE microscope equipped with a computer-controlled z-stage and connected to a TCS-4D UV/VIS confocal scanning system (Leica AG, Benzheim, and Leica Lasertechnik, Heidelberg, Germany). The staining protocol for fluorescein-conjugated annexin V (detection of PS on the outer leaflet of the plasma membrane) was adapted for neuronal cultures as follows: CGC were grown on glass-bottomed culture dishes and incubated with MPP⁺ or rotenone with or without inhibitors. At the end of the incubation periods, 0.5 µg/ml H-33342 was added to the cultures to later visualize chromatin structure. After a 10-min incubation at 37°, CGC were washed for 10 sec with binding buffer (10 mM HEPES, 140 mM NaCl, 2.5 mM CaCl₂, 10 mM MgCl₂) and subsequently incubated for 2 min in the dark with annexin V diluted 1:100 in binding buffer. After a new wash with binding buffer, stained cultures were immersed in binding buffer supplemented with 0.25 µM ethidium homodimer-1 and visualized by three-channel confocal microscopy (blue, chromatin structure, green, annexin binding, red, membrane integrity) using a 63×/NA 1.32 UV-corrected lens.

Statistics. Toxicity experiments were run in triplicate and repeated in three to eight cell preparations. Statistical significance was calculated on the original data sets using the Student's t test. When variances within the compared groups were not homogeneous, the Welch test was applied. Western blots and measurements of [Ca²⁺]_i were repeated in at least three independent cell preparations.

	Results

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A role for the NMDA-R in MPP⁺-induced CGC apoptosis. MPP⁺ caused apoptosis of differentiated (8 DIV) CGC in a concentration range of 25-100 µM (Fig. 1). Toxicity of MPP⁺ was evident in cells exposed to MPP⁺ for 2-4 hr, unlike the slow toxicity observed in an earlier report (Marini et al., 1989). Susceptibility of CGC to such rapidly developing MPP⁺-induced apoptosis was strongly dependent on the cell differentiation state. CGC were hardly sensitive toward MPP⁺ during the first 2 DIV, whereas they became progressively more susceptible to induction of apoptosis elicited by MPP⁺ with increasing time in culture, as described elsewhere for other excitotoxic stimuli (i.e., NMDA, NO, or ONOO) (Leist et al., 1997a) (Fig. 1).

View larger version (18K): [in this window] [in a new window]   Fig. 1.   MPP+-induced apoptosis in CGC cultures from wild-type and PARP/ animals. Cultures from wild-type CGC of different culture age (wt) or PARP-deficient mice (PARP/) were challenged with MPP+ concentrations as indicated. The percentage of neurons with apoptotic nuclei was determined after 7 hr by staining with fluorescent chromatin dyes and counting of the cells. Data are mean ± standard deviation of triplicate determinations.

View larger version (34K): [in this window] [in a new window]   Fig. 2.   Prevention of MPP+-induced CGC apoptosis by blockade of Ca2+ channels. Top, CGC were preincubated for 30 min with solvent (0.4% DMSO; con) or 2 µM MK801, 500 µM AP5, 5 mM Mg2+, 1 µM verapamil (Ver), 1 µM nifedipine (Nif), 500 nM tetrodotoxin (TTX), 400 µM N-methyl-L-arginine (NMA), or 1 mM nitroarginine (NA) before they were challenged with 50 µM MPP+. Medium was exchanged for CSS after 4 hr, and cell death parameters were determined after 18 hr. Apoptosis was quantified by scoring condensed neuronal nuclei (note inverted scale); mitochondrial function was determined by measuring MTT-reduction capacity. Bottom, CGC were incubated as described above with different concentrations of inhibitors of the NMDA-R glycine site (DNQX, 6,7-dichloroquinoxaline-2,3-dione, or DCK) plus 50 µM MPP+. Apoptosis was quantified after 18 hr. Data are mean ± standard deviation of triplicate determinations.

Rapid ATP depletion as a possible trigger of MPP⁺ excitotoxicity. Because energy depletion is known to trigger excitotoxic processes in different experimental systems, we examined intracellular ATP levels after treatment of CGC with MPP⁺. ATP declined slowly during the first 30 min of treatment with MPP⁺, and it was dissipated 90% after 3-4 hr. Treatment with MK801 delayed the loss of ATP by 30-60 min; however, at 3-4 hr after exposure to MPP⁺, ATP also was depleted in these cells (Fig. 3A). These findings suggest that MPP⁺ caused a primary ATP depletion that was largely independent of the secondary excitotoxicity. This also implied that ATP depletion alone was not immediately lethal to neurons but rather sensitized them to glutamate and facilitated glutamate release. Under our culture conditions (residual glucose concentrations of ~1 mM in the medium), glycolytic ATP production alone was not sufficient to maintain intracellular ATP concentrations at control levels after exposure to MPP⁺. Accordingly, the addition of 10 mM glucose delayed MPP⁺ toxicity, which suggests that glycolysis remained functional. Also, when we measured the glucose consumption of neurons, the basal rate of 20 nmol/10⁶ cells/hr increased to ~400 nmol/10⁶ cells/4 hr in the presence of MPP⁺, regardless of whether MK801 was included in the culture dish.

View larger version (98K): [in this window] [in a new window]   Fig. 3.   NMDA-R and prevention of mitochondrial damage caused by MPP+. A, CGC were preincubated for 30 min with solvent (0.4% dimethylsulfoxide) or 2 µM MK801 and then challenged with 50 µM MPP+. At the times indicated, the medium was removed, cells were lysed, and ATP was determined luminometrically. Intracellular ATP content is expressed as percentage of that measured in untreated cells (control) at t = 0 (2.8 nmol of ATP/mg of protein). Viability and were determined in parallel cultures from the same preparation. TMRE fluorescence did not significantly drop in CGC treated with MK801 + MPP+, whereas it was fully lost after 60-90 min in cells treated with MPP+ only. Viability of MPP+ + MK801-treated cells was maintained for 16 hr, whereas nuclear condensation in MPP+-treated neurons started after 45 min. Data represent mean ± standard deviation of triplicate determinations. B, CGC incubated in control medium, with 50 µM MPP+, or with MPP+ plus 2 µM MK801 were stained after 4 hr with H-33342 (chromatin), calcein (cell membrane integrity), and TMRE (mitochondrial ). Samples were imaged by confocal microscopy using fluorescence and polarization contrast (PC). Mitochondria from the axodendritic network (TMRE neurites) and those within the cell body (TMRE somata) were imaged at different focal planes. MPP+-treated neurons had condensed nuclei, whereas the plasma membrane (calcein staining) was still intact. TMRE staining in both neurites and somata disappeared. Only astrocytes (fifth row, middle: one astrocyte is shown with the cell body in the lower right corner and processes extending to the left) retained (TMRE staining). MK801 prevented both nuclear condensation and loss of mitochondrial function. All experiments were repeated in at least six cell preparations with similar results. Scale, width of each micrograph corresponds to 108 µm in CGC cultures. C, CGC were preincubated with or without 2 µM MK801 and challenged with 50 µM MPP+. Cytochrome c release into the cytosol was determined by Western blot. Cells treated with 0.5% Triton X-100 (Triton) were analyzed to indicate the maximal cytochrome c release. D, CGC were incubated with 25 nM rotenone or with rotenone plus 2 µM MK801. They were stained after 4 hr with H-33342 (chromatin). In rotenone-treated cells, all TMRE staining (mitochondrial ) was lost, and nuclei appeared condensed. Astrocyte nuclei and their mitochondria were unaffected (astrocyte in lower right corner with processes extending up and to the left). MK801 prevented nuclear condensation and loss of TMRE.

Prevention of mitochondrial damage by NMDA-R block. In this system, MPP⁺ may damage mitochondria by two ways: (i) by direct radical-induced damage and (ii) by triggering excitotoxicity, which results in Ca²⁺ overload and subsequent mitochondrial failure. We distinguished between these two mechanisms by comparing MPP⁺-triggered mitochondrial effects in the presence and absence of NMDA-R blockers. First, changes in were examined by staining CGC with the mitochondrial potential-sensitive dye TMRE (Fig. 3B). CGC pretreated with MK801 (Fig. 3B) or AP5 (not shown) and then exposed to MPP⁺ showed no significant loss of . In neurons exposed to MPP⁺ alone, ATP depletion was paralleled by a complete loss of , in analogy with a model of direct excitotoxicity (Ankarcrona et al., 1995). Thus, loss of seemed to be due to indirect excitotoxicity and preceded nuclear condensation and loss of membrane integrity. MPP⁺-induced loss of TMRE fluorescence was selective for neurons because astrocytes within the same culture dish lost neither TMRE fluorescence nor viability. To confirm this finding, also was monitored with the indicators JC-1 (Ankarcrona et al., 1995) and Mito-Tracker Red (CMX-rosamine). All dyes yielded similar results, but TMRE was most convenient for routine measurements because it was less photolabile than JC-1 and more suitable for monitoring long time periods (24 hr) than CMX-Ros, which reacts with cellular thiols.

NMDA-R block inhibits loss of mitochondrial membrane potential and toxicity triggered by classic mitochondrial inhibitors. In view of these findings, we considered that NMDA-R-dependent toxicity might be triggered by other compounds affecting mitochondria. To evaluate this possibility, we exposed CGC to low concentrations of mitochondrial poisons such as oligomycin, CCCP, or rotenone. The latter blocks the respiratory chain at a site similar to that affected by MPP⁺ (Kilbourn et al., 1997). Toxicity of oligomycin, CCCP, or rotenone was significantly (50% in all cases) reduced/delayed by MK801 (not shown). Pretreatment with MK801 completely prevented the toxicity of low concentrations of rotenone (25-50 nM) and maintained in short term incubation (Fig. 3D).

The role of NO generation and exocytosis in MPP⁺ toxicity. Endogenous NO production contributes to MPTP-induced pathology in vivo (Hantraye et al., 1996). Thus, we tested whether endogenously formed NO was involved in our system. Inhibitors of the neuronal NO synthase did not reduce MPP⁺-induced apoptosis, nor did they modify other cytotoxicity parameters (Fig. 2A). One mechanism potentially involved in neuronal excitotoxic death is the activation of the enzyme PARP downstream to NO-mediated DNA damage (Zhang et al., 1994). The mechanism of MPP⁺-induced excitotoxicity in CGC was, however, unrelated to PARP activation because neurons prepared from PARP^/ mice were equally sensitive as those from wild-type mice (Fig. 1). Together, these findings suggested that MPP⁺ has the potential to activate excitotoxic mechanisms independent of NO generation.

View larger version (37K): [in this window] [in a new window]   Fig. 4.   MPP+-triggered exocytosis and autocrine excitotoxicity in CGC. A, CGC were preincubated for 10 hr with 20 ng/ml BoNT/C (130 pM) or 500 ng/ml tetanus toxin (3 nM) before they were challenged with MPP+. The medium was exchanged 4 hr after challenge for CSS, and survival was determined after 18 hr by MTT assay. Data are mean ± standard deviation from triplicate determinations. *, p  0.05. B, CGC were challenged with 50 µM MPP+ ± MK801. After 2 hr, CGC were loaded with Fura-2, and [Ca2+]i was determined by ratiometric video imaging and in situ calibration. Data are mean ± standard deviation from 16-20 neurons. Similar results were obtained in three different cell preparations. The [Ca2+]i measured in MPP+-treated CGC is close to the saturation range of the indicator and therefore may be 1 µM. *, p  0.05. C, CGC were pretreated with DMSO alone (solvent) or 2 µM MK801 (MK801) and loaded with fluo-3 acetoxymethyl ester before they were challenged with 120 µM MPP+. The increase of [Ca2+]i in time (relative fluo-3 fluorescence) was measured by confocal microscopy. Data are mean ± standard error from 10-15 neurons. D, CGC were challenged in CSS containing 1.8 mM Ca2+ (+ Ca2+) or no nominal Ca2+ ( Ca2+) with 100 µM MPP+ or 100 nM rotenone. After 3 hr, apoptosis was determined by staining of cells with H-33342 and counting of the percentage of condensed nuclei. In three independent experiments, the average apoptosis was 84 ± 7% (MPP+ + Ca2+), 88 ± 8% (rotenone + Ca2+), and 10% under all other conditions.

View larger version (39K): [in this window] [in a new window]   Fig. 5.   MK801 prevents rotenone-induced Ca2+ increase and PS translocation. A, CGC were incubated with (open symbols) or without (filled symbols) 1 µM MK801. Rotenone (1 µM) was added as indicated (arrow), and the change of [Ca2+]i was followed by confocal imaging of fluo-3 fluorescence. Data represent the mean ± standard deviation from 15-25 cells in three independent experiments. B, CGC were pretreated with 1 µM MK801 or not treated. At 2 hr after the addition of rotenone (25 nM), cells were stained with fluorescent annexin V (PS translocation) and H-33342 (chromatin structure). Fluorescence images were collected by confocal microscopy. Note that cells with only marginally condensed nuclei are already annexin V positive. Phosphatidylserine translocation (annexin V stainability) was completely prevented by MK801. Data are representative for four experiments in three independent cell preparations.

The role of proteases in MPP⁺- or rotenone-induced autocrine excitotoxicity. To further characterize the steps between NMDA-R-mediated [Ca²⁺]_i increase and cell death, we examined the role of cellular thiol proteases. Inhibition of either caspases or calpains was sufficient to block apoptosis elicited by MPP⁺ concentrations up to 50 µM (Fig. 6, A and B). The same set of inhibitors also protected CGC from the apoptosis induced by 50 nM rotenone (80% cells remained viable in the presence of each of five different protease inhibitors) (Fig. 6E). Because all available inhibitors lack an absolute specificity for a single type of proteases (Villa et al., 1997), we used three or five structurally different agents for each class of proteases with similar results. Inactive control peptides (100 µM) with similar end groups as the inhibitors were not effective. We also established sensitive in vitro assays with purified calpain or recombinant caspase-3 to test cross-reactivity of the inhibitors between the two classes of thiol proteases: for instance, z-D-cbk was found to be a highly potent inhibitor of caspase-3 (IC₅₀ < 200 nM) without any inhibitory effect on calpains (IC₅₀ > 200 µM). Calp II showed exactly opposite characteristics. Thus, both caspases and calpains seemed to be required to mediate apoptosis of CGC challenged with 50 µM MPP⁺. Cells protected by protease inhibitors remained viable for 24 hr, when they received new medium or CSS. At very high intensities of insult (e.g., 100 µM MPP⁺; >4-hr toxin exposure), the protective effect of protease inhibitors was bypassed.

View larger version (41K): [in this window] [in a new window]   Fig. 6.   Caspase or calpain inhibitors prevent MPP+ triggered intracellular proteolysis and apoptosis. CGC were incubated with (A) various caspase inhibitors or (B) calpain inhibitors at the concentrations indicated (logarithmic scale). At 30 min later, they were challenged with 50 µM MPP+. The percentage of apoptotic neurons was determined by counting the number of cell bodies. Data are mean ± standard deviation of triplicate determinations. C, Inhibition of fodrin proteolysis was studied by Western blotting of extracts from CGC pretreated with seven different concentrations of the caspase inhibitor z-D-cbk (top) or the calpain inhibitor calp III (bottom) and challenged with 50 µM MPP+. As positive control for fodrin proteolysis, we used the proapoptotic toxin colchicine (1 µM). Colchicine treatment (12 hr) produced proteolytic fragments of 150 and 120 kDa (bottom, first lane), whereas MPP+ (4-hr treatment) mainly produced the 150-kDa fragment. D, CGC were pretreated with various caspase or calpain inhibitors (100 µM) or MK801 (2 µM), or the inactive caspase inhibitor analogue z-F-cmk, before they were challenged with MPP+ (50 µM) for 4 hr. Fodrin proteolysis was analyzed by Western blot. E, CGC were pretreated with z-D-cbk (100 µM) or Ac-YVAD-cmk (YVAD; 100 µM) before they were exposed to 50 µM kainate, 25 µM glutamate, or 50 nM rotenone for 90 min. Then, medium was exchanged for CSS plus glucose. Apoptosis was quantified by staining neurons with H-33342 and determining the percentage of condensed nuclei. Data are mean values from triplicate determinations. *, p < 0.05; **, p < 0.01 solvent versus inhibitor.

Protease inhibitors act downstream to the Ca²⁺ influx via the NMDA-R. Two sets of experiments were performed to test whether protease inhibitors indeed acted downstream of NMDA-R-mediated [Ca²⁺]_i increase. First, we measured NMDA-R-mediated [Ca²⁺]_i increases in CGC in the presence of protease inhibitors. None of the peptide inhibitors had a significant inhibitory effect (Fig. 7). Similar experiments were performed using as agonists glutamate, kainate, or high [K⁺]. None of the protease inhibitors blocked [Ca²⁺]_i increase due to these stimuli (not shown).

View larger version (20K): [in this window] [in a new window]   Fig. 7.   Protease inhibitors do not inhibit NMDA-R-mediated neuronal Ca2+ increase. CGC were pretreated with various protease inhibitors (100 µM) or MK801 and loaded with fluo-3. They were challenged with 200 µM NMDA as indicated, and the fluorescence increase (F/F0) was recorded by confocal microscopy. Data were obtained from 10-15 neurons, and experiments were repeated in three different CGC preparations.

The role of proteases and the NMDA-R in MPP⁺-induced chromatin breakdown. Caspase-3 activation has recently been shown to be a direct upstream mechanism leading to oligonucleosomal DNA-fragmentation. In CGC, oligonucleosomal DNA fragmentation was not significantly activated within the first 4 hr after challenge, when most other processes relevant to cell death had already occurred. In glutamate-challenged CGC oligonucleosomal DNA fragmentation is a late event and may not be evident at all (Ankarcrona et al., 1995). In analogy with the glutamate model, we observed here high molecular weight DNA fragmentation into 600-, 300-, and 50-kbp fragments in CGC treated with MPP⁺. This characteristic feature of apoptotic chromatin degradation was prevented by MK801 or by inhibitors of either calpains or caspases (Fig. 8A). This suggests the involvement of a proteolytic step, downstream to the initial [Ca²⁺]_i increase, which results in chromatin breakdown. This proteolytic step does not seem to be linked to caspase-3 activation. Consistent with the lack of oligonucleosomal DNA fragmentation and with the proteolysis pattern of fodrin [caspase-3 forms a 120-kDa fragment, whereas caspase-4, caspase-2, or calpain forms mainly a 150-kDa fragment (Nath et al., 1996)], we did not detect caspase-3-like (DEVD-afc cleavage) activity in CGC challenged with MPP⁺. Notably, DEVD-afc cleavage activity instead was easily detected in the same cell preparation challenged with colchicine or low [K⁺] (Leist et al., 1997d). Further evidence for the absence of caspase-3 activation in the MPP⁺ model is the lack of processing of procaspase-3 to the active caspase. Procaspase-3 was not cleaved even at a time point where fodrin was already 90% cleaved and nuclei were condensed (Fig. 8B).

View larger version (46K): [in this window] [in a new window]   Fig. 8.   MPP+-triggers NMDA-R- and caspase-dependent apoptotic chromatin degradation but not procaspase-3 proteolysis. A, CGC were preincubated for 30 min with 100 µM Ac-YVAD-cmk, z-D-cbk, calp III, or 2 µM MK801 or solvent control only. Then, they were challenged with 50 µM MPP+ or they were left unchallenged (control CGC). After 4 hr, incubations were stopped and DNA fragmentation was analyzed by field-inversion gel electrophoresis. Less than 5% of CGC had lost membrane permeability at that time point. B, CGC were challenged with MPP+ in the presence and absence of MK801. Cells were harvested after 3 hr, when fodrin was already nearly maximally cleaved (see Fig. 6) and >80% of apoptotic nuclei were scored. Alternatively, apoptosis was induced by switching cells to low K+ medium for 24 hr (low K+; ~75% apoptotic neurons). Control cells were incubated for 24 hr in high K+ (<10% apoptotic neurons). A decrease of procaspase-3 (casp3) due to activation of the protease is seen in the low K+ model but not in the MPP+ model. Activated murine caspase-3 was not detected by the antibody used. Therefore, the blot membrane was redeveloped after stripping with an anti-actin antibody to confirm equal protein loading.

Prevention of MPP⁺-induced PS translocation by NMDA-R blockade and caspase inhibition. Translocation of PS from the inner to the outer surface of the plasma membrane is an early and specific event in apoptosis (Leist et al., 1997b, 1997d), which seems to be dependent on caspase activation, at least in tumor cells. In CGC, MPP⁺ caused PS translocation, as detected by annexin V staining, within 1-2 hr. MPP⁺-treated neurons stained intensively when the first signs of chromatin condensation were visible and well before the nucleus became fully pyknotic. CGC remained stainable for several hours until membrane permeability was lost and membranes were degraded. Virtually all neurons with condensed chromatin had lost plasma membrane lipid asymmetry (90% annexin V-positive neurons at MPP⁺ concentrations of 50 µM in four independent experiments). Often, large spherical membrane blebs were formed from the axons, which also stained positively with annexin V and had an intact membrane. Notably, annexin V binding to neuronal bodies was completely prevented by MK801 (Fig. 9) and was triggered by direct exposure to glutamate (not shown), which suggests that PS translocation is linked to NMDA-R-mediated Ca²⁺ influx and not to direct MPP⁺ actions. In addition, exposure to rotenone induced PS translocation, which was inhibited by MK801 (Fig. 5B). Annexin V labeling also was inhibited by the caspase inhibitor z-D-cbk, in agreement with the view that caspases also are pivotal for this feature of neuronal apoptosis (Fig. 9).

View larger version (82K): [in this window] [in a new window]   Fig. 9.   MPP+ triggers NMDA-R- and caspase-dependent apoptotic PS translocation. CGC were incubated with MPP+ for 2 hr before they were stained with FITC-annexin V and H-33342 and imaged by confocal microscopy. Top, an xy section through the cell bodies (4 µm from coverslip). Apoptotic nuclei seem to be condensed and hyperfluorescent and are surrounded by annexin V-stained membranes. A transversal section (xz plane) demonstrates that the entire surface of the apoptotic neurons plus some non-chromatin-containing blebs formed from the axonal projections are surface stained with annexin V. Neurons that had been preincubated with z-D-cbk or MK801 did not condense and did not stain with annexin V.

	Discussion

Top Summary Introduction Procedures Results Discussion References

MPP⁺ elicits autocrine excitotoxicity in CGC. Our results show that MPP⁺ induces apoptosis of cultured CGC by eliciting autocrine excitotoxicity. MPP⁺ can induce either apoptosis or necrosis in vivo (Tatton and Kish, 1997), and previous studies have shown that either form of cell death can be induced in vitro, by mechanisms unrelated to excitotoxicity (Marini et al., 1989; Du et al., 1997b). Low MPP⁺ concentrations predominantly elicit apoptosis, whereas high concentrations trigger necrosis (Hartley et al., 1994; Du et al., 1997b). In animals, removal of glutamatergic inputs (decortication), blockers of glutamate release, or NMDA-R antagonists reduce MPTP- and MPP⁺-induced striatal damage and dopamine depletion (Srivastava et al., 1993) or loss of dopaminergic neurons in the substantia nigra (Turski et al., 1991).

Mechanisms of indirect excitotoxicity. Several mechanisms have been implicated in direct excitotoxic neuronal death, including excessive NO production and subsequent activation of PARP (Zhang et al., 1994), mitochondrial alterations (Ankarcrona et al., 1995), and protease activation (Siman and Noszek, 1988; Du et al., 1997a). Although brain NO synthase has an aggravating role in some models of excitotoxicity, NO plays a minor role as a direct mediator of toxicity in CGC (reviewed in Leist et al., 1997a). CGC strongly express nitric oxide synthase and therefore may have developed mechanisms to prevent direct NO toxicity. Consistent with this assumption, MPP⁺ toxicity was not altered by nitric oxide synthase inhibitors. PARP activation is triggered by DNA damage, which may be caused by NO (Zhang et al., 1994). We found no alteration in the cell death rate or mode of cell death in cells prepared from PARP^/ mice, which further supports the lack of involvement of NO in MPP⁺ toxicity. In contrast, in CGC stimulated with MPP⁺, mitochondrial dysfunction and protease activation seem to be key events, which mediate excitotoxic cell death.

Proteases in excitotoxic death. Apoptosis is associated with the activation of a proteolytic cascade, probably involving different sets of proteases, which operate virtually at all stages of the cell death program (i.e., signaling, control, and execution) (Villa et al., 1997). Here, we found that both caspases and calpains were activated downstream to the NMDA-R-mediated [Ca²⁺]_i increase and upstream of PS translocation, nuclear condensation, and DNA fragmentation. Under appropriate conditions, cells pretreated with protease inhibitors survived MPP⁺ challenge for several days. Obviously, different sets of proteases can interact to cause neuronal death; examples include caspases and calpains (Nath et al., 1996; Jordán et al., 1997; current study), different caspases and serine proteases (Stefanis et al., 1997), and caspases plus the proteasome.

Excitotoxic apoptosis. Excitotoxic cell death may occur by either apoptosis or necrosis. Seemingly divergent observations are reconciled by the finding that the intensity of insult may determine the mode of cell death (e.g., Ankarcrona et al., 1995; Du et al., 1997a; Leist and Nicotera, 1998) and that excessive Ca²⁺ entry may convert the mode of cell death in some cases from apoptosis to necrosis (reviewed in Leist et al., 1997d). Intracellular ATP levels seem to be critical in determining the shape of cell death (Ankarcrona et al., 1995; Leist et al., 1997c), and ATP may be required at multiple sites. For instance, ATP/dATP is required for the activation of caspase-3, which is a central protease in many apoptosis models. Here, we show that apoptosis proceeds in the absence of procaspase-3 processing. This also may explain the lack of oligonucleosomal DNA fragmentation, which is a direct consequence of caspase-3 activation. In MPP⁺-treated neurons, other cell death proteases may be activated by mechanisms not directly requiring ATP.