Ethanol-Induced Death of Postnatal Hippocampal Neurons

Neurobiology of Disease 10, 396 – 409 (2002) doi:10.1006/nbdi.2002.0523

Ethanol-Induced Death of Postnatal Hippocampal Neurons Krista L. Moulder,* Tao Fu,* Heidi Melbostad,* Robert J. Cormier,* Keith E. Isenberg,* Charles F. Zorumski,* ,† and Steven Mennerick* ,† *Department of Psychiatry and †Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, Missouri 63110 Received August 8, 2001; revised April 23, 2002; accepted for publication May 24, 2002

Fetal alcohol exposure causes severe neuropsychiatric problems, but mechanisms of the ethanolassociated changes in central nervous system development are unclear. In vivo, ethanol’s interaction with N-methyl-D-aspartate (NMDA) and ␥-aminobutyric acid type A (GABA A) receptors may cause increased apoptosis in the immature forebrain. We examined whether ethanol affects survival of neonatal hippocampal neurons in primary cultures. A 6-day ethanol exposure killed hippocampal neurons with an LD50 of ⬃25 mM. Elevated extracellular potassium or insulin-related growth factor 1 inhibited cell loss. Although potentiation of GABA A receptors or complete block of NMDA receptors also kills hippocampal neurons, pharmacological studies suggest that ethanol’s interaction with GABA A and NMDA receptors is not sufficient to explain ethanol’s effects on neuronal survival. Ca 2ⴙ influx in response to depolarization was depressed >50% by chronic ethanol treatment. We suggest that chronic ethanol may promote neuronal loss through a mechanism affecting Ca 2ⴙ influx in addition to effects on postsynaptic GABA and glutamate receptors. © 2002 Elsevier Science (USA)

Fetal alcohol syndrome is the extreme end of a spectrum of deleterious effects of ethanol on the immature nervous system. Among the consequences of early ethanol exposure are mental retardation, hyperactivity, learning disorders, and subsequent psychiatric disorders (Famy et al., 1998; Kelly et al., 2000; Mattson & Riley, 1998). An understanding of the precise cellular changes that accompany exposure of the immature nervous system to ethanol may elucidate the neural underpinnings of these behavioral problems and offer insight into possible treatments. At present, the effects of ethanol on the immature nervous system are incompletely understood. Effects of ethanol on the cerebellum have been documented in great detail, and previous work has demonstrated changes in neurogenesis, neuronal morphology, and enhanced cell death of differentiated neurons (Ward & West, 1992). In cultured cerebellar granule neurons, ethanol either causes enhanced cell loss when administered alone or reverses the trophic action of low glutamate concentrations (Bhave & Hoff-

man, 1997; Castoldi et al., 1998; Saito et al., 1999; Wegelius & Korpi, 1995). Changes in the gross anatomy of the forebrain have also been described (Roebuck et al., 1998; Ward & West, 1992), and these changes are likely to participate in the behavioral abnormalities listed above. The details of cellular changes in the forebrain in response to ethanol are unclear, but recent evidence suggests that neonatal rats exposed to ethanol undergo enhanced apoptosis during the period of physiological neuronal death in the forebrain (Ikonomidou et al., 2000). Because the first few postnatal weeks in the rat roughly correspond to the last trimester of human embryonic development (Dobbing & Sands, 1979), the increased apoptosis is likely to be an important component of human fetal alcohol syndrome (Ikonomidou et al., 2000). Several questions remain regarding ethanol-induced neuronal death in neonatal animals. The enhanced apoptosis observed in whole animals is mimicked by N-methyl-d-aspartate (NMDA) receptor (NMDAR) antagonists and by ␥-aminobutyric acid ©

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type A (GABA A) receptor (GABAR) positive modulators, suggesting that these specific actions of ethanol may impart ethanol’s neurotoxic actions (Ikonomidou et al., 2000). However, it is difficult in these studies to exclude nutritional or metabolic consequences of ethanol exposure. Also, it is difficult to distinguish the direct effects of ethanol on susceptible neurons from ethanol’s effects on peripheral or remote regions, which in turn influence the viability of susceptible neurons. Finally, it is unclear whether longer exposure to ethanol, such as mimics the binge drinking of some heavy drinkers, might recruit additional mechanisms that contribute to cell loss. Therefore, we have begun to explore the effect of ethanol on hippocampal neurons in culture, where tighter control over several relevant experimental variables can be achieved.

METHODS Hippocampal Cultures Cultures were prepared using previously described procedures (Mennerick et al., 1995). Hippocampi were harvested from albino Sprague–Dawley rat pups at postnatal days 1–3 and dissociated by papain and mechanical dispersion. Mass cultures were prepared by plating cells at a seeding density of 1500/mm 2 onto a 35-mm culture dish containing a layer of collagen (0.5 mg/ml). For plating microcultures, cells were dispersed at 100/mm 2 onto a culture dish coated with 0.15% agarose, dried, and sprayed with collagen droplets (Mennerick et al., 1995). Cultures were used for electrophysiology 8 –17 days following plating, or as described under Results. Electrophysiology Whole-cell recordings were obtained using pipettes of 1.5- to 4-M⍀ open-tip resistance when filled with the standard pipette solution containing (in mM) KCl (130), NaCl (5), CaCl 2 (0.5), EGTA (5), and Hepes (10), pH adjusted to 7.25 with KOH. At the time of experiments, culture medium was replaced with an extracellular recording saline consisting of NaCl (138), KCl (4), CaCl 2 (2.0), MgCl 2 (1.0), glucose (10), and Hepes (10), pH 7.25 with NaOH. Drugs were delivered by local perfusion with a multibarrel pipette positioned approximately 0.5 mm from the cell of interest. For experiments examining electrophysiological responses of cells chronically treated with ethanol, the same concentration of ethanol present in the growth

medium was also added to the recording chamber to avoid studying ethanol withdrawal effects. Current amplitudes in the absence of ethanol were then studied by washing away ethanol with local perfusion of ethanol-free saline. Except for experiments in which NMDAR-mediated excitatory postsynaptic currents (EPSCs) were specifically examined, 25 ␮M d(⫺)-2amino-5-phosphonopentanoic acid (d-APV) was included in the bath solution. For NMDAR-mediated EPSCs, bath Mg 2⫹ was omitted and 10 ␮M glycine plus 1 ␮M NBQX (6-nitro-7-sulphamoylbenzo[f]quinoxaline-2,3-dione) were added to the bath solution. For examination of responses to exogenous NMDA, the bath solution contained 0.5 mM CaCl 2, no added MgCl 2, 10 ␮M glycine, and 5 ␮M NBQX. Other alterations to these standard solutions are given in the text and figure legends. Autaptic release of neurotransmitter was stimulated in voltage clamped solitary microculture neurons with a 2.0-ms voltage pulse to 0 mV. The holding potential before and after the stimulus was ⫺70 mV. Stimulation rates for all experiments were ⬍0.05 Hz. Control conditions were interleaved with experimental conditions whenever possible to account for any time-dependent changes in transmission. EPSCs were easily distinguished from GABAergic IPSCs inhibitory postsynaptic current on the basis of PSC time course (Mennerick et al., 1995). Previous experiments have shown that all PSCs in postnatal hippocampal microcultures are either glutamatergic or GABAergic, defined by kinetics of the PSC and sensitivity to appropriate receptor antagonists (Mennerick et al., 1995). Membrane capacitance, determined for experiments measuring the chronic effect of ethanol on calcium currents (Fig. 8), was estimated from the integral of capacitive transients in response to 10-mV (20 ms) hyperpolarizing voltage pulses. For these experiments, the sample rate was 100 kHz, and the four-pole Bessel signal filter of the patch amplifier was set to a cutoff frequency of 50 kHz. The instantaneous amplitude of the capacitive transient was estimated by fitting the 10 –99% decay region of the transient with a double exponential (Mennerick et al., 1995) and extrapolating the fit to the onset of the voltage pulse. Data were collected using an Axoclamp 1-D patchclamp amplifier (Axon Instruments, Foster City, CA) interfaced to a Digidata 1200 acquisition board and a Pentium II computer. Synaptic responses were sampled at 5–10 kHz. The PClamp software suite (Axon Instruments) was used for data acquisition and analysis. Data were plotted using Sigma Plot (SPSS soft©

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398 ware). Data are presented in the text and figures as means ⫾ standard error.

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RESULTS Chronic Ethanol Exposure Kills Hippocampal Neurons in Vitro

Immunostaining Cultures were fixed for 5 min in 4% paraformaldehyde, 0.2% glutaraldehyde in phosphate-buffered saline. Cells were permeablized with 0.1% Triton X-100, blocked with 10% serum, and incubated for 3–12 h in primary antibody. Caspase 3 primary antibody (New England Biolabs) was used at a final dilution of 1:500. Signal amplification was achieved using a Vector Elite kit (Vector) with diaminobenzidine as chromogen or a tyramide signal amplification kit (NEN) used according to the manufacturer’s instructions.

Survival and Cell Counts Survival was typically assessed at 10 days in vitro, 6 days following administration of ethanol or other toxins. Protective agents were coadministered at the time of toxin treatment. Survival was assessed with counts of remaining neurons under phase-contrast optics using an inverted Nikon TE300 microscope. In several experiments (N ⫽ 2 experiments at 6 days of ethanol treatment and N ⫽ 2 at 3 days of ethanol treatment), blinding the counter to experimental conditions did not affect quantification. Cells were counted as live if somas retained a smooth, phase-bright appearance with visible processes. In some cases Hoechst 33342 or immunostains for activated caspase 3 were used to quantify dying neurons, as described under Results. Differences from untreated control cultures from the same plating were used to quantify cell loss. Ethanol was added directly to the culture medium to initiate exposure. Cultures were then transferred to a closed, humidified chamber containing 500 ml of ethanol at the same concentration as the culture medium (Isenberg et al., 1992). Control dishes were placed in an identical chamber containing a water bath. Culture chambers were gassed daily with a mixture of 5% CO 2 /95% air. In six platings, we measured ethanol (spectrophotometric assay employing alcohol dehydrogenase/NAD; Sigma, St. Louis, MO) in dishes to which 230 mg/dL (50 mM) was added. After 6 days of exposure, the measured concentrations of ethanol were 1.5 ⫾ 1.5 mg/dL (control) and 200.1 ⫾ 3.8 mg/dL ethanol. No correction was made for the small loss of ethanol over time. 2002 Elsevier Science (USA) All rights reserved.

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To determine whether forebrain neurons are sensitive to ethanol-induced cell death, we treated hippocampal neurons for 6 days beginning at 4 –5 days in vitro with various concentrations of ethanol. This is a protocol similar to that recently used by us to demonstrate the neurotoxicity of GABAR positive modulators in this same preparation (Xu et al., 2000). We found that chronic ethanol treatment induced death of hippocampal neurons in a concentration-dependent manner. Dying neurons were found in controls and more prominently in ethanol-treated cultures. The profile of dying neurons was characterized by condensed, fragmented nuclei, as is often associated with apoptotic cell deaths (Figs. 1A and 1B). Significant loss of neurons compared with control cultures was detected with as low as 10 mM ethanol (Fig. 1C; N ⫽ 4 experiments, P ⬍ 0.05). Fits to pooled data from four experiments suggested that 25 mM ethanol represented the LD50 concentration for hippocampal neurons. This places the concentration at which ethanol effectively kills hippocampal neurons in the range at which ethanol has important effects on GABARs, NMDARs, and perhaps other ion channels (Crews et al., 1996) in addition to being a clinically relevant concentration. The effect of ethanol on survival was specific to neurons. In five experiments in which cultures were treated with 50 mM ethanol, astrocyte numbers, assessed by counting Hoechst-33342-stained nuclei, were not significantly different than in control cultures (16 ⫾ 9% loss of astrocytes, P ⬎ 0.05). In contrast, at DIV 10, following 6 days of 50 mM ethanol exposure, ethanol-treated cultures exhibited higher ratios of fragmented neuronal nuclei to normal nuclei (85 ⫾ 5% of neurons possessed fragmented nuclei, N ⫽ 3 platings) compared with control cultures (51 ⫾ 2% fragmented nuclei, P ⬍ 0.05). This result also highlights the significant levels of natural cell death that occur in these cultures. We did not perform a full evaluation of the time course of ethanol-induced neuronal loss to avoid evaporation of ethanol from the culture medium during evaluation. However, in several experiments, we examined cells treated with 50 mM ethanol at days 3– 4 of treatment and found only a 35 ⫾ 5% cell loss relative to control cultures (N ⫽ 18 experiments, P ⬍ 0.01). Therefore, the cell death was slow, similar to the slow death occurring with GABAR overstimulation

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We tested for other similarities with death by overinhibition (Xu et al., 2000). Chronic incubation of neurons in elevated KCl protects hippocampal neurons from death occurring from GABAR overstimulation (Xu et al., 2000) and protects against other types of apoptotic cell death (Franklin & Johnson, 1992). KCl also protects against ethanol-induced apoptosis of cerebellar granule neurons in vitro (Wegelius & Korpi, 1995). We tested the effect of 30 mM KCl, added simultaneously with ethanol to the culture medium at 4 days in vitro. As in our studies of GABA modulators, we found that KCl significantly protected against both spontaneous neuronal loss and cell death induced by 50 mM ethanol (Figs. 2A and 2B). Figure 2B shows specific KCl protection against ethanol-induced cell loss, as opposed to the “natural” cell loss. Counts from cultures treated with ethanol alone are normalized to

FIG. 1. A, B. Phase-contrast (A1, B1) and fluorescence (A2, B2) photomicrographs of control cultures (A1, A2) and ethanol-treated cultures (B1, B2) after 6 days of exposure to 100 mM ethanol. The cultures were aldehyde fixed, stained with Hoechst 33342, and visualized with epifluorescence. Note the increased presence of bright, fragmented nuclei (open arrows) in the ethanol-treated cultures at the expense of normal, intact nuclei (solid arrows). The same field of cells is represented in the phase contrast and fluorescence images for each condition. The scale bar indicates 55 ␮m. C. Concentration response relationship for cell survival following 6 days of exposure to ethanol (EtOH). The solid line represents a fit to the Hill equation with a half-maximal concentration of 25 mM. Values were obtained by counting smooth, phase-bright neuronal somas with intact neurites and normalizing counts to control cultures from the same litter and plating (N ⫽ 4 experiments).

(Xu et al., 2000) but in contrast to more rapid forms of cell death resulting from, for instance, glutamate toxicity.

FIG. 2. Depolarization and IGF-1 are partially protective against 50 mM ethanol-induced neuronal death. A. Effect of 30 mM additional KCl on neuronal survival. Survival is plotted as (experimental counts)/(control counts)-1. Thus, positive values depict increased survival relative to control, and negative values depict decreased survival. Note that KCl alone promotes survival of cultures as previously reported (Xu et al., 2000). B. The ethanol effects in A have been normalized to the respective control condition to highlight the reduction in ethanol-induced death by KCl. Ethanol-induced death was significantly reduced by KCl (P ⬍ 0.01, N ⫽ 5 experiments). C, D. Similar experiments performed with the neurotrophic factor insulin related growth factor 1 (IGF, 50 ng/ml). Again, survival of control cultures was improved (C), and ethanol-induced death was significantly reduced (D) by IGF treatment (P ⬍ 0.05, N ⫽ 5 experiments).

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400 untreated cultures, and counts from ethanol plus KCl are normalized to KCl alone. The results show that KCl significantly reduced the degree of ethanol-induced death in the cultures (P ⬍ 0.05, N ⫽ 5 experiments). Many forms of apoptosis are also sensitive to neurotrophic factor intervention. We used IGF-1 (50 ng/ mL), coapplied at the onset of ethanol exposure, to determine the sensitivity of ethanol-induced cell loss to peptide trophic support. IGF-1, like KCl, increased the number of surviving control neurons and also protected against ethanol-induced death (P ⬍ 0.05; Figs. 2C and 2D). Figure 2D shows normalized data for IGF-1 similar to those depicted in Fig. 2B. Protection by chronic depolarization and by growth factors is consistent with the idea that ethanol-induced neuronal loss is largely apoptotic, as chronic depolarization by KCl is protective against many forms of apopotic cell death (Collins & Lile, 1989; Franklin & Johnson, 1992; Gallo et al., 1987). Apoptosis is often defined biochemically by the activation of effector caspases. We immunostained hippocampal cultures for activated caspase 3, an effector caspase in many forms of neuronal apoptosis (Thornberry & Lazebnik, 1998; Wilson, 1998). Immunostaining for activated (cleaved) caspase 3 has advantages over other measures of caspase activity because it permits the identification of the cell type containing the activated caspase, a useful feature in our mixed cultures of astrocytes and neurons. We immunostained cells treated with ethanol for 3 days and found a significant increase in the number of immunopositive neurons in ethanol-treated cultures versus control cultures (Figs. 3A–3G). In these same experiments the average number of healthy unstained neurons, counted under phase-contrast optics, was 52.9 ⫾ 6.5 for control cultures and 35.7 ⫾ 5.3 for ethanol-exposed cultures (N ⫽ 18 experiments at treatment days 3– 4; P ⬍ 0.001). Thus, the percentage of cells immunoreactive for caspase 3 to healthy neurons was 18 ⫾ 3% in control cultures and 53 ⫾ 9% in ethanol-treated cultures (Fig. 3G). At later time points the ratio of caspase 3 positive neurons to unstained, healthy neurons remained higher than that in control cultures (data not shown). However, the absolute number of immunoreactive cells did not remain elevated, presumably because of complete loss of many neurons at these later time points. Most neurons exhibiting immunoreactivity for activated caspase 3 exhibited shrunken somas and few remaining neurites, similar to the profile of neurons containing fragmented, condensed nuclei when 2002 Elsevier Science (USA) All rights reserved.

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stained with Hoechst 33342 (Figs. 1A, 1B, and 3A–3E). In some experiments, we double labeled cultures with activated caspase 3 antibody and with Hoechst 33342 to stain nuclei. There was excellent overlap of caspase3-labeled cytoplasm and condensed, fragmented neuronal nuclei (data not shown). On occasion, we observed immunoreactive neurons with largely intact neurites, presumably in the early stages of apoptosis (Fig. 3E). At any given time point, these immunoreactive neurons of relatively normal morphology constituted a small fraction of all immunoreactive neurons, suggesting that the time that neurons remain morphologically normal following caspase 3 activation is probably brief. As an additional control, we examined cleaved caspase 3 immunoreactivity in cultures treated with NMDA to promote acute excitotoxicity. Cultures of the same age as those used for ethanol-associated activated caspase 3 staining were treated with 100 ␮M NMDA for 30 – 60 min. Cells were fixed following this insult, stained for activated caspase 3, and counted. The NMDA treatment resulted in a similar amount of cell loss as 3– 4 days of ethanol treatment (35 ⫾ 16% death, N ⫽ 3 experiments). However, while ethanol treatment resulted in more than a doubling of caspase immunoreactive neurons relative to controls (Fig. 3F), we found no increase in the number of caspase 3 immunoreactive neurons in the NMDA-treated cultures. In fact, we found a trend toward fewer activated caspase 3 immunoreactive neurons (⫺32 ⫾ 14% change relative to sibling control dishes, N ⫽ 3, P ⫽ 0.07, Fig. 3H). This experiment demonstrates that caspase cleavage is not simply a nonspecific result of cell death. Taken together, the results in Fig. 3 suggest an important component of caspase 3 associated apoptotic cell loss, although they do not exclude the participation of other mechanisms. GABA Receptors and Ethanol Effects The time course of cell loss with ethanol exposure in vitro is similar to the death time course we previously observed in these postnatal cultures with GABAR overstimulation. Also, ethanol has been shown to enhance GABAR function acutely. However, ethanol’s GABA A potentiating properties have been debated and are not apparent under all conditions (Criswell et al., 1993; Soldo et al., 1994; Weiner et al., 1997). We therefore sought to test the hypothesis that GABAR overactivity is responsible for ethanol’s effects on neuronal survival. To determine whether an acute GABA potentiating action of ethanol may participate in the

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FIG. 3. Caspase 3 activation is associated with 50 mM ethanol-induced neuronal loss. A–D. Phase-contrast (A, C) and brightfield (B, D) photomicrographs depicting control (A, B) and ethanol-treated (C, D) cultures fixed and immunostained for activated caspase 3 following 3 days of exposure to 50 mM ethanol. The open arrows in B and D mark examples of four neurons immunoreactive for caspase 3. The solid arrows denote four examples of healthy, unstained neurons, which can be more clearly visualized in the phase-contrast panel (A). Note that most immunopositive cells have shrunken cell bodies and no remaining neurites. The scale bar in C represents 50 ␮m and applies to A–D. E. Higher power brightfield photomicrograph of another field of ethanol-treated cells. One positive cell, presumably in the early stages of apoptosis, still has largely intact, immunoreactive processes (vertical open arrow). Scale bar represents 35 ␮m. F. Summary of the absolute number of cells per 20⫻ microscope field exhibiting immunoreactivity for activated caspase 3 after 3– 4 days of incubation in 50 mM ethanol (open bar) or in sibling control cultures (solid bar) (N ⫽ 18 experiments, P ⬍ 0.01). G. Summary of the ratio of caspase positive cells (counted with brightfield optics) to healthy cells (counted with phase-contrast optics) after 3 days of ethanol treatment (N ⫽ 18, P ⬍ 0.01). H. Summary of the absolute number of activated cells exhibiting caspase 3 immunoreactivity following exposure to 100 ␮M NMDA for 30 – 60 min (N ⫽ 3 experiments).

chronic effect of ethanol on neuronal survival, we examined the effect of ethanol on autaptic IPSCs of solitary hippocampal neurons. Examining evoked autaptic IPSCs allows a straightforward assessment of whether ethanol acutely affects presynaptic and/or postsynaptic signaling. Surprisingly, we could not detect an effect of 100 mM ethanol on GABAR-mediated postsynaptic currents (Fig. 4A). In contrast, other postsynaptic modulators, such as pentobarbital (Fig. 4B) and neurosteroids (not shown), robustly potentiated IPSCs. It is possible that ethanol does not affect GABARs in solitary GABA neurons. Alternatively, ethanol may not effectively potentiate the actions of

brief, high concentrations of GABA present at synapses. To test these possibilities, we examined the effect of acute and chronic ethanol exposure on responses to low concentrations of exogenously applied GABA in the same mass cultures in which chronic ethanol toxicity studies were performed. We explored whether chronic incubation of hippocampal cells in 50 mM ethanol for 4 –5 days (before maximum death was achieved) altered the current density of GABAR-mediated currents or induced sensitivity of GABARs to ethanol. In 21 control (untreated) neurons and 21 ethanol-treated cells (4 –5 days) from three independent ©

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FIG. 4. Ethanol’s effect does not involve GABAR activity. A. Superimposed autaptic IPSCs from a solitary GABAergic neuron obtained in the absence and presence of 100 mM ethanol. No effect of ethanol on the IPSC was detected. For this and subsequent examples of PSCs, presynaptic stimulus transients have been blanked for clarity. B. As a positive control, the IPSC from another neuron was challenged with pentobarbital (Ptb; 50 ␮M, thick trace), which dramatically prolonged the IPSC decay. C, D. Responses to exogenous GABA (2 ␮M) were also unaffected by 100 mM ethanol in either control cultures (C) or cultures chronically treated for 5 days with 50 mM ethanol (D). E. Summary of the chronic effects of ethanol exposure on GABA responsiveness from experiments like those depicted in C and D. Steady-state current amplitude of responses to 2 ␮M GABA were normalized to cell capacitance (current density) to account for any changes in cell size induced by ethanol (N ⫽ 21 neurons from three independent platings for each condition). F. Summary of the acute effects of ethanol (100 mM) on GABA current amplitude for either control cultures (solid bar) or cultures treated chronically with 50 mM ethanol for 5 days. Ethanol was coapplied with GABA as shown in C and D. Note that the acute EtOH effect on currents was less than 5% in both untreated and EtOH-treated cells.

experiments, the current density in response to a subsaturating concentration of GABA (2 ␮M) was similar in ethanol-treated and control neurons (Figs. 4C– 4E). In a subset of these neurons, we also examined the 2002 Elsevier Science (USA) All rights reserved.

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acute ethanol sensitivity of GABA currents and again found no effect of ethanol on GABA currents elicited from chronically treated cultures (Figs. 4C, 4D, and 4F). It is possible that factors, such as neurosteroids or phosphorylation factors (Criswell et al., 1999; Harris et al., 1995), which may be important for ethanol-induced modulation of GABARs, are present during chronic experiments but are not present in the acute setting of electrophysiological experiments. In the final, and perhaps most definitive, experiment to test a role of ethanol’s GABAR potentiating activity in the neurotoxic effects of ethanol, we examined the effect of GABAR blockade on ethanol-induced toxicity. Consistent with our previous results, we found that 50 ␮M pentobarbital caused widespread neurotoxicity in hippocampal cultures (Xu et al., 2000). This toxicity was significantly reduced by coincubation with 50 ␮M bicuculline. In three experiments, pentobarbital caused a 67 ⫾ 13% reduction in viable cells over 6 days of treatment. With 50 ␮M bicuculline, death was only 36 ⫾ 8% of cultures treated with bicuculline alone (Fig. 5B). Additionally, we have previously shown that bicuculline prevents the death of cells exposed to GABA-potentiating neurosteroids (Xu et al., 2000). In contrast, bicuculline did not appreciably affect the toxicity induced by chronic exposure of cells to 50 mM ethanol (Fig. 5A). NMDA Receptor Hypofunction and Ethanol Effects Taken together, the results of Figs. 4 and 5 strongly suggest that GABAmimetic actions of ethanol do not participate substantially in the neurotoxic effects of ethanol in hippocampal cultures. We therefore turned our attention to the NMDAR antagonist effect of ethanol, another reported target relevant to ethanol’s proapoptotic effect in vivo. NMDAR antagonists are also neurotoxic to immature neurons in vivo (Ikonomidou et al., 1999). We found, consistent with previous reports (Lovinger et al., 1989; Peoples et al., 1997; Popp et al., 1999), that ethanol partially blocked NMDAR-mediated EPSCs (Fig. 6A). Supporting the suggestion that NMDAR antagonists are directly neurotoxic, we found that chronic incubation of neurons with 50 ␮M d-APV caused loss of many neurons (59 ⫾ 16% compared with untreated, N ⫽ 7 experiments). The death induced by d-APV was nearly completely prevented by coincubation with 30 mM KCl (Fig. 6B). These observations suggest that blockade of NMDARs is also toxic to hippocampal neurons and that NMDAR

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to explain the effect of ethanol on neuronal survival, then submaximum concentrations of selective NMDAR antagonists should produce cell loss similar to that observed with 50 –100 mM ethanol. We titrated the concentrations of a competitive (d-APV) and a noncompetitive (7-chlorokynurenic acid) NMDAR an-

FIG. 5. The GABAR antagonist bicuculline does not affect ethanolinduced survival of hippocampal neurons. A. Effect of 50 ␮M bicuculline coapplied with ethanol on day in vitro 4. The effect of bicuculline was normalized to sibling cultures treated with bicuculline alone as in Figs. 2B and 2D. There was no significant effect of bicuculline on ethanol-induced death (N ⫽ 4 experiments). B. Similar experiments performed with pentobarbital (50 ␮M) as toxin revealed a significant protective effect of bicuculline (N ⫽ 3 experiments, P ⬍ 0.05).

block could be relevant to ethanol’s effect on hippocampal neuron survival. To test the acute actions of ethanol on NMDARs directly, we examined isolated autaptic NMDAR-mediated EPSCs and found that ethanol blocked EPSCs by 37 ⫾ 2% at 50 mM (N ⫽ 7) and by 43 ⫾ 5% at 100 mM (N ⫽ 11), consistent with previous reports suggesting that effects on NMDA receptors are nearly maximum at 50 mM ethanol. Ethanol also slightly decreased the 10 –90% decay time of NMDAR EPSCs (23 ⫾ 5% decrease at 100 mM). In contrast, pharmacologically isolated AMPA receptor (AMPAR) mediated EPSCs were only slightly affected by 100 mM ethanol (6 ⫾ 2% depression). These data suggest that presynaptic targets cannot explain the effect of ethanol on NMDAR EPSCs and that AMPARs are not an important target of acutely applied ethanol. These data are consistent with previous results suggesting that maximum block of NMDARs is not 100%, even at high concentrations of ethanol (Lovinger et al., 1989; Peoples et al., 1997; Popp et al., 1999). Therefore, if the acute effect of ethanol on NMDARs is sufficient

FIG. 6. NMDAR block is insufficient to account for ethanol-induced death. A. The effect of 100 mM ethanol on an isolated NMDAR-mediated EPSC. Note that the block of EPSCs is incomplete. B. Effect of a 6-day incubation of cultures in 50 ␮M d-APV on survival of neurons. The effect of 30 mM KCl, normalized to survival in KCl alone in sibling cultures, is also shown. d-APV-induced cell loss was significantly attenuated by KCl (P ⬍ 0.05, N ⫽ 6). C. d-APV (1 ␮M) mimicked the partial inhibition of NMDARs. D. However, neither this concentration of d-APV nor a concentration of the noncompetitive antagonist 7-chlorokynurenate (7-CK) chosen to partially block NMDA receptor-mediated EPSCs mimicked the full effect of 50 mM ethanol exposure on neuronal survival (N ⫽ 5 d-APV experiments and 4 7-CK experiments). E. Effect of chronic ethanol (50 mM) exposure on NMDA current density. A protocol similar to that used for measuring GABA current density was used (Fig. 4E). Ethanol-treated cells (EtOH) had been exposed to 50 mM ethanol for 4 –5 days. NMDA currents were elicited at a membrane potential of ⫺70mV by 10 ␮M NMDA in a bath solution containing 0.2 mM Ca 2⫹, 10 ␮M glycine, and no added Mg 2⫹. These conditions were chosen to limit desensitization of NMDA-gated responses. There was no significant difference in the NMDA current density in control and ethanol-treated cells (P ⫽ 0.73, N ⫽ 21 control and 19 ethanol-treated cells from three independent platings).

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tagonist to yield ⬃40% block of NMDARs. We found that in 20 ␮M glycine, 50 ␮M 7-chlorokynurenic acid produced 44 ⫾ 2% inhibition of NMDAR EPSCs (N ⫽ 4). Likewise, 1 ␮M d-APV produced 37 ⫾ 2% inhibition of NMDAR EPSCs (N ⫽ 4; Fig. 6C). We did not attempt to use the more commonly employed noncompetitive antagonist MK-801 because the use-dependent, slowly reversible nature of MK-801 block is clearly different from the fast onset and offset block observed with ethanol. We evaluated the effect of these submaximum concentrations of d-APV and 7-chlorokynurenate on neuronal survival. Despite the similarity in effects at NMDARs, the selective NMDAR blockers consistently caused less cell death than 50 mM ethanol. For instance, 50 ␮M 7-chlorokynurenate caused only 21 ⫾ 17% cell death over 6 days (N ⫽ 4 experiments; Fig. 6D), while 1 ␮M d-APV caused only 11 ⫾ 19% cell death (N ⫽ 5 experiments). Figure 6E shows that chronic ethanol exposure also did not cause a downregulation of NMDA current. Taken together, these results suggest that NMDAR hypofunction cannot account for all of ethanol’s toxic effect but could work in concert with other factors to induce apoptosis. Voltage-Gated Calcium Influx and Ethanol Effects In many models of activity-dependent neuronal survival, intracellular calcium levels appear to be a key determinant of survival versus apoptosis. Depression of voltage-gated Ca 2⫹ channels promotes apoptosis in other cell types (Galli et al., 1995; Koike et al., 1989). Also, moderate calcium influx through voltage-gated calcium channels is important in the neuroprotective actions of KCl (Collins & Lile, 1989; Franklin & Johnson, 1992; Galli et al., 1995). These observations are accounted for by the calcium setpoint hypothesis (Franklin & Johnson, 1992), which suggests that excessively low or high intracellular calcium levels promote apoptosis and moderate intracellular calcium elevations promote cell survival. Also, existing literature suggests that ethanol may interact directly with calcium influx in several cell types (Catlin et al., 1999). We therefore designed experiments to test directly whether depression of calcium influx causes apoptosis of hippocampal neurons and whether ethanol may directly affect calcium influx in these cells. Because survival of other cell types is especially dependent upon L-type Ca 2⫹ currents (Collins & Lile, 1989; Koike et al., 1989), we examined the effect of the L-type Ca 2⫹ channel blocker nifedipine. We found that depressing calcium influx directly with 15 ␮M nifed2002 Elsevier Science (USA) All rights reserved.

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FIG. 7. Effect of block of Ca 2⫹ influx and general activity on survival of hippocampal neurons. A. Effect of nifedipine (15 ␮M) and nifedipine plus 30 mM KCl administered over 6 days. The effect of nifedipine plus KCl is compared to sibling cultures treated with KCl alone. Note the ineffectiveness of KCl-induced depolarization in overcoming the nifedipine-induced death. B. Similar to experiments shown in Fig. 3F, the number of neurons immunoreactive for activated caspase 3 after 3– 4 days of treatment with nifedipine was increased (P ⬍ 0.05, N ⫽ 9 experiments). C. The effect on survival of inhibition of spiking activity with tetrodotoxin (100 nM) is shown. Note that TTX-induced death was significantly attenuated by KCl (P ⬍ 0.05, N ⫽ 3 experiments).

ipine over 6 days in vitro caused the loss of 77 ⫾ 5% of neurons relative to control (untreated) cultures from the same plating (N ⫽ 5 experiments; Fig. 7A). In addition, nifedipine counteracted the survival-promoting effect of KCl on hippocampal cultures (Fig. 7A). Nifedipine-treated cultures exhibited a similar profile of increased activated caspase 3 immunoreactive neurons as ethanol-treated cultures (Fig. 7B). These results show that direct depression of calcium influx causes neuronal loss, and the results are consis-

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tent with the idea that depression of calcium influx may be important in the death produced by overinhibition and chronic ethanol treatment of hippocampal neurons. We also tested a global blocker of cellular activity, tetrodotoxin (TTX), for its effect on neuronal survival. Like d-APV, GABA potentiators, and nifedipine, TTX also caused slow loss of hippocampal neurons (Fig. 7C). Like d-APV and GABA potentiators, but unlike nifedipine, TTX caused cell loss that was reduced by coincubation in 30 mM KCl (Fig. 7C). Ethanol interacts directly with voltage-dependent Ca 2⫹ currents, either positively or negatively depending on the ethanol exposure time, cell type, and possibly other variables (Catlin et al., 1999). We therefore tested whether an interaction with voltage-gated Ca 2⫹ channels could participate in the cell death that could not be fully explained by GABAR overactivation and NMDAR hypofunction. In other cell types, L-type calcium channel function is depressed by ethanol, while chronic incubation upregulates calcium currents (Catlin et al., 1999). Neither effect has been previously reported in hippocampal neurons. We examined effects of ethanol on high voltage-gated Ca 2⫹ currents in hippocampal neurons using whole-cell voltage clamp recordings. To enhance current size, we used 4 mM extracellular Ba 2⫹ (other divalents omitted) as the charge carrier. We found that 50 mM ethanol, applied acutely, had no effect on voltage-gated Ba 2⫹ currents in hippocampal neurons. However, chronic incubation for 5 days in 50 mM ethanol depressed Ba 2⫹ current density by about half in hippocampal neurons (Figs. 8A and 8B). It should be noted that while the average size of neurons in ethanol-treated cultures was often smaller than in control cultures (presumably due to ongoing ethanol-induced death), some selection bias was exercised in these experiments to record from cells of approximately the same size under the various treatment conditions. As a result of this attempt to record from similarly sized neurons, we saw no difference among experimental groups in the membrane capacitance estimated from the integral of capacitive current transients in response to hyperpolarizing voltage pulses (Fig. 8C). Thus, the difference in calcium current density was present even in cells that were chronically treated with ethanol but which were still apparently normal morphologically. To test whether the effect on calcium current density is related to NMDAR hypofunction induced by 50 mM ethanol, we examined the effect of 50 ␮M 7-chlorokynurenate treatment for 5 days on whole-cell Ba 2⫹ currents. We found no significant difference in the

FIG. 8. Depression of calcium currents by chronic ethanol exposure may underlie apoptosis induced by ethanol. A. Ca 2⫹ currents elicited by voltage steps from ⫺70 to 0 mV in a control neuron and a neuron from a culture treated for 5 days with 50 mM ethanol. Ethanol was also present in the recording solution from the cell chronically treated with ethanol. The capacitance of the control cell was 20.9 pF and that of the ethanol-treated cell was 18.4 pF. B. Summary of the current density from control cultures, ethanol (EtOH; 50 mM) -treated cultures, and cultures treated with 50 ␮M 7-chlorokynurenate (7-CK). Numbers below labels indicate the number of cells tested under each condition (from three to five independent platings). Difference from control was statistically significant for ethanol (P ⬍ 0.05) but not for 7-chlorokynurentate. C. In the same cells, there was no difference in measured membrane capacitance.

density of voltage-gated Ca 2⫹ current in cultures treated with low concentrations of the selective NMDAR blocker (Fig. 8B). Therefore, we conclude that the chronic ethanol-induced depression of voltage-gated Ca 2⫹ currents is unrelated to the effect of ethanol on NMDARs but likely contributes significantly to the death of hippocampal neurons. Because L-type channels have been implicated in the survival of other neurons (Koike et al., 1989), we next sought to determine whether L-type Ca 2⫹ chan©

2002 Elsevier Science (USA) All rights reserved.

406 nels might be preferentially depressed by ethanol treatment. In control and ethanol-treated cultures, we examined the acute nifedipine sensitivity of calcium current evoked by a depolarizing pulse to 0 m V. We detected no difference in the percentage of high-voltage activated Ba 2⫹ current blocked by 5 ␮M nifedipine (27 ⫾ 8% depression for ethanol-treated and 26 ⫾ 9% depression for control cultures; N ⫽ 22 and 23 cells, respectively, from three independent platings). This degree of nifedipine sensitivity is consistent with other estimates of the contribution of L-type calcium current to total high-voltage activated current in these cells (Nakashima et al., 1998). In the same cells in which nifedipine sensitivity was examined, ethanol induced an overall 37 ⫾ 8% decrease in calcium current density (P ⬍ 0.05). Because nifedipine sensitivity remained unchanged, these results suggest that ethanol depresses L-type calcium current as well as other subtypes of high-voltage activated calcium currents. Therefore, although nifedipine can mimic the effect of ethanol on survival, depression of L-type current by ethanol may not completely account for ethanol’s effects on survival. It is likely that depression of at least one other calcium channel subtype and depression of NMDA-receptor-mediated neurotransmission also participate in ethanol’s overall effects on survival of hippocampal neurons.

DISCUSSION Consistent with the idea that ethanol is directly toxic to forebrain neurons, we find that ethanol exposure is toxic to hippocampal neurons in vitro, with cell loss occurring at concentrations as low as 10 mM (⬃40 mg %). Although we have recently shown that GABAmimetics are also toxic to hippocampal neurons over a similar time course, the ethanol neurotoxicity is not explained by the GABAmimetic activity of ethanol. Similarly, NMDA antagonist activity is not sufficient to explain ethanol’s effect on neuronal survival in culture. We find that ethanol administered chronically, but not acutely, depresses voltage-activated calcium currents. Because direct blockers of calcium currents are also toxic to hippocampal neurons, we propose that in addition to effects on postsynaptic receptors, ethanol directly depresses calcium signaling in hippocampal neurons, leading to cell death, apparently through an apoptotic mechanism. Although we have not exhaustively tested all criteria for the mechanism of ethanol-induced cell death, four lines of evidence suggest that a major component 2002 Elsevier Science (USA) All rights reserved.

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Moulder et al.

of ethanol-induced death in vitro is apoptotic. Ethanolinduced cell loss is sensitive to depolarization, characteristic of other forms of apoptosis (Koike et al., 1989). The ethanol-induced cell loss is also sensitive to neurotrophic peptides (IGF-1), also characteristic of various forms of apoptotic cell death (Galli et al., 1995). Ethanol-induced death is associated with nuclear fragmentation, which has been used as a hallmark of apoptosis (although the specificity of this marker has been questioned: Gwag et al., 1997; Ishimaru et al., 1999). Finally, caspase 3, which is a downstream effector in many forms of neuronal apoptosis (Wilson, 1998), is clearly activated in ethanol-treated neurons. Other criteria have also previously been used to assess apoptotic versus nonapoptotic cell death. For instance, sensitivity of death to caspase inhibitors has been used to help define apoptotic cell death. While apoptosis of sympathetic neurons deprived of trophic support is protected by caspase inhibition (Deshmukh et al., 1996), activity-dependent apoptosis of cerebellar granule neurons is associated with caspase 3 activation but is relatively insensitive to caspase blockade (Miller et al., 1997a). In preliminary experiments, we found no effect of the pan caspase inhibitor zVADfmk when coapplied with ethanol for 3 days (N ⫽ 9, data not shown). This may suggest a mechanism similar to activity-dependent apoptosis of cerebellar granule neurons and that parallel caspase-dependent and caspase-independent pathways contribute to hippocampal neuronal loss with ethanol. Ultrastructural criteria have also been used to define apoptosis (Ishimaru et al., 1999). Forebrain neurons in situ exhibit apoptotic ultrastructural profiles upon ethanol exposure (Ikonomidou et al., 2000). Future work will determine whether hippocampal neurons exposed to ethanol in vitro exhibit a similar progression of apoptotic ultrastructural changes. Our results demonstrate that cultured postnatal hippocampal neurons are susceptible to cell death in response to prolonged ethanol exposure. In particular, our results suggest that recently observed susceptibility of widespread regions of the forebrain to ethanol exposure in vivo is likely due at least partly to direct effects of ethanol on susceptible neurons rather than to (1) indirect polysynaptic effects of ethanol or (2) metabolic by-products of ethanol. Our results do show some differences from in vivo results, however, which are likely due to differences in the in vivo and in vitro environments. The time course of cell death is notable. In vitro, the time course of cell death is similar among ethanol and selective NMDA and GABA receptor modulators,

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with death occurring over several days. In vivo results suggest that exposure to 0.2 g/dL ethanol can result in a dramatic increase in the number of apoptotic neurons evaluated 24 h following exposure (Ikonomidou et al., 2000). The time course of apoptosis seen after administration of ethanol to the whole animal is also similar to that observed with administration of selective NMDAR antagonists or GABAR potentiators. These similarities and differences between the models may suggest either differences in the inherent susceptibility of neurons or differences in the presence of apoptotic modulating factors in the two environments. The longer exposure time to chronic ethanol in vitro may invoke additional mechanisms (i.e., Ca 2⫹ current down-regulation) that are not involved in the in vivo model because of the short exposure times involved. Our results suggest that GABAR potentiation is insignificant in vitro and that incomplete NMDAR blockade, which mimics ethanol’s maximum effects at NMDARs, is at best mildly toxic. In vivo, a combination of GABA potentiation and NMDAR block is sufficient to recapitulate ethanol-induced apoptosis. It seems likely that factors are present in vivo, notably neurosteroids (Criswell et al., 1999), that enhance ethanol’s effects at GABARs and perhaps enhance ethanol’s pro-apoptotic ability. It is also possible that differences in the phosphorylation status of GABARs in vitro vs in vivo may affect susceptibility to ethanol modulation (Wafford & Whiting, 1992). Other in vitro models of CNS neurons have shown that ethanol can either be directly apoptotic or enhance death in response to withdrawal of KCl support (Bhave & Hoffman, 1997; Castoldi et al., 1998; Saito et al., 1999; Wegelius & Korpi, 1995). As with forebrain neurons, this death has been proposed to result primarily from the action of ethanol at NMDARs. Our results show that low concentrations of specific inhibitors of NMDARs do not cause the same degree of neuronal loss as ethanol causes. At least one other effect in addition to NMDAR and GABAR actions appears necessary to explain the full toxicity promoted by ethanol in vitro. Hippocampal neurons are thus one of many cell types dependent upon basal activity levels and calcium influx for survival (Xu et al., 2000; Mennerick & Zorumski, 2000). In many cell types, KCl protection from apoptotic cell death is via promotion of Ca 2⫹ influx through L-type Ca 2⫹ channels (Collins & Lile, 1989; Franklin & Johnson, 1992; Gallo et al., 1987; Koike et al., 1989). This appears to be true in our system as well, as nifedipine blocks the protection afforded by KCl. There is debate about the down-

stream mechanisms of moderately elevated Ca 2⫹. In some systems, there is evidence for Ca 2⫹-dependent upregulation of peptide neurotrophic factor release or sensitivity (Ghosh et al., 1994; Meyer-Franke et al., 1998). In other systems, it has been proposed that Ca 2⫹ may interact intracellularly with biochemical pathways accessed by conventional peptide trophic factors (Miller et al., 1997b; Yano et al., 1998). We also observe protection from ethanol-induced death by IGF-1, consistent with an apoptotic mechanism of ethanol-induced death. As with KCl, the mechanisms underlying IGF-1 protection are unclear. It is possible that, similar to KCl, IGF-1 directly upregulates Ca 2⫹ infux (Blair et al., 1999). In hippocampal neurons, depression of Ca 2⫹ influx through voltage-gated Ca 2⫹ channels is an additional participant in the total ethanol-induced loss observed. This ethanol-induced depression of Ca 2⫹ influx through voltage-gated Ca 2⫹ channels may be one reason that depolarization by KCl is only partially effective in protecting against ethanol-induced neuronal loss. It seems likely that down-regulation of Ca 2⫹ influx through NMDARs and voltage-gated Ca 2⫹ channels acts in concert to render hippocampal neurons apoptotic, but we cannot exclude a contribution of other mechanisms, perhaps unrelated to cellular electrical activity and Ca 2⫹ influx, to ethanol-induced cell death. Because of the membrane permeability of ethanol and the ubiquity of cellular targets for ethanol (Harris, 1999; Samson & Harris, 1992; Weight, 1992), we cannot exclude a role for other targets, including other ion channels, in the cell death we observe. Previously, long-term ethanol treatment has been reported to have mixed effects on voltage-gated Ca 2⫹ currents. Several studies have demonstrated up-regulation of Ca 2⫹ influx in PC12 cells (Catlin et al., 1999; Messing et al., 1986). Thus, it is likely that differences between these results and our observations in primary cultures of hippocampal cells result from phenotypic differences in the response of different cell types to chronic ethanol exposure. In cerebellar neurons, changes in Ca 2⫹ influx have also been reported with chronic exposure to ethanol. In these studies, effects on Ca 2⫹ influx were complicated. Net down-regulation (Zou et al., 1995) and up-regulation (Gruol & Parsons, 1994) of high-voltage activated Ca 2⫹ currents have been reported for cerebellar neurons. Further study in hippocampal cells will be necessary to determine the intracellular pathways involved in ethanolinduced depression of Ca 2⫹ influx. ©

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408

ACKNOWLEDGMENTS The authors thank Ann Benz for help with preparation of cultures and John Olney, Nuri Farber, Lou Muglia, Yuki Izumi, and David Wozniak for discussion. This study was supported by NIH Grants GM47969, AA12951, and MH45493 (CFZ), an Alzheimer’s Disease Research Center Pilot Grant, the Klingenstein Fund, a NARSAD Young Investigator Award, NS40488, and AA12952 (SM). KM was supported by DA07261. Some of these data were published in abstract form.

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