Neuroprotective Action of Cycloheximide Involves Induction of Bcl-2 and Antioxidant Pathways

Dissociated cell cultures of fetal rat hippocampus were established from 18-d embryos and maintained as described previously (Mattson et al., 1989, 1995). Cells were plated in 35-mm-diam plastic or glass-well dishes on a polyethyleneimine substrate in 0.8 ml of medium consisting of MEM with Earle's salts supplemented with heat-inactivated FBS (5%; vol/vol), 1 mM l-glutamine, 1 mM pyruvate, 20 mM KCl, and 26 mM sodium bicarbonate (pH 7.2). Cells were grown at a density of 80–120 cells per mm² culture surface, and experiments were performed in 8–10-d-old cultures, a time period during which neurons express both N-methyl-d-aspartate (NMDA) and non-NMDA glutamate receptors, and are vulnerable to excitotoxicity (Mattson et al., 1993c), Aβ toxicity (Mattson et al., 1993a), and FeSO₄ toxicity (Zhang et al., 1993). Cycloheximide, glutamate, buthionine sulfoximine, and FeSO₄ were purchased from Sigma Chemical Co. (St. Louis, MO) and prepared as 200–500 × stocks in saline (pH 7.2). Aβ25-35 (lot ZM500) was purchased from Bachem California (Torrance, CA) and stored in lyophilized form, and 1 mM stocks were prepared by dissolving the peptide in sterile distilled water 2–4 h before use. Oligodeoxynucleotides (ODNs) were purchased from IDT Inc. (Coralville, IA). The sequence of the Bcl-2 antisense ODN was 5′-TCCCGGCTTGCGCCAT-3′. Three different control ODNs were used: sense ODN; missense ODN, 5′-TCGCGGCATGCCCCAT-3′; and nonsense ODN, 5′-CTGTCGCGCTCGACTC3′. Immediately before experimental treatment, the culture maintenance medium was replaced with Locke's solution that had the following composition (mM): 154 NaCl; 5.6 KCl; 2.3 CaCl₂; 1.0 MgCl₂; 3.6 NaHCO₃; 5 Hepes; 10 glucose.

These methods are detailed in our previous studies (Mattson et al., 1989, 1995). Briefly, viable neurons were counted in premarked microscope fields (×10 objective) before experimental treatment and 20–24 h after treatment. Most neurons that died during the exposure period to glutamate, FeSO₄, or Aβ were absent at the 20-h time point, and viability of the remaining neurons was assessed by morphological criteria. Neurons with intact neurites of uniform diameter and a soma with a smooth appearance were considered viable. Neurons with fragmented neurites and a vacuolated and/or swollen soma were considered nonviable.

These methods were those described previously (Martin et al., 1992). Cultures (incubated in methionine- and cysteine-free medium) were pretreated with vehicle or CHX, and then [³⁵S]methionine/cysteine (ICN Radiochemicals, Irvine, CA; sp act >1,000 Ci/mmol) was added to a final concentration of 25 μCi/ml (in the continued presence of CHX). 30 min later, cultures were washed once with medium, and 2 ml of a solution containing 0.5% SDS, 1 mM EDTA, 10 mM Tris (pH 7.5) was added. 40 μg of BSA was added to each sample, and protein was precipitated with 10% TCA on ice. Precipitated protein was collected by filtration onto 0.45-μm nitrocellulose filters. Filters were washed twice with cold 10% TCA and dried, and radioactivity was quantified in a liquid scintillation counter.

Analyses of gene expression were performed as described previously (Estus et al., 1994). Briefly, polyA+ RNA was isolated from cells after the indicated treatments by using an oligo-dT–cellulose-based purification kit (QuickPrep Micro kit; Pharmacia Fine Chemicals, Piscataway, NJ) and concentrated by coprecipitation with glycogen, all as directed by the manufacturer. Half of the mRNA was converted to cDNA by reverse transcription (RT) by using random hexamers (16 μM) to prime Moloney murine leukemia virus reverse transcriptase (Superscript; Life Technologies, Grand Island, NY). The 30-μl reaction contained 50 mM Tris (pH 8.3), 40 mM KCl, 6 mM MgCl₂, 1 mM DTT, 500 μM each dATP, dTTP, dCTP, dGTP, and 20 U RNasin (Promega, Madison, WI). After 10 min at 20°C, the samples were incubated for 50 min at 42°C, and the reaction was then terminated by heating to 94°C for 5 min.

For PCR amplification of specific cDNAs, stock reactions (50 μl) were prepared on ice and contained 50 μM dCTP, 100 μM each dGTP, dATP, and dTTP, 15 μCi α-[³²P]dCTP (3,000 Ci/mmol), 1.5 mM MgCl₂, 50 mM KCl, 10 mM Tris (pH 9.0), 0.1% Triton X-100, 1 μM each primer, 1 U of Taq polymerase, and 3% of the cDNA synthesized in the RT reaction. The sequences of the primers used in this study were: NFM sense primer: 5′ ACG CTG GAC TCG CTG GGC AA 3′, NFM antisense primer: 5′ GCG AGC GCG CTG CGC TTG TA 3′ (156-bp product); c-jun sense primer: 5′ ACT CAG TTC TTG TGC CCC AA 3′, c-jun antisense primer: 5′ CGC ACG AAG CCT TCG GCG AA 3′ (64-bp product); c-fos sense primer: 5′ AAT AAG ATG GCT GCA GCC AA, c-fos antisense primer: 5′ TTG GCA ATC TCG GTC TGC AA 3′ (116-bp product); bcl-2 sense primer: 5′-CTGTACGGCCCCAGCATGCG-3′, bcl-2 antisense primer: 5′-GCTTTGTTTCATGGTACATC-3′ (210-bp product). The stock solutions were separated into three equal aliquots that were covered with a drop of mineral oil and subjected to various numbers of PCR cycles to determine the minimum number of cycles necessary to detect the PCR product. The typical reaction conditions were 1 min at 95°C, 1 min at 55°C, and 2 min at 72°C. The results depicted here represent 16–22 cycles of amplification. After amplification, the cDNAs were separated by electrophoresis on 12% polyacrylamide gels, visualized by autoradiography of the dried gels, and quantified with a PhosphorImager (Molecular Dynamics, Sunnyvale, CA). The identity of the PCR product amplified by each primer pair was confirmed typically either by direct sequencing (fmole sequencing kit; Promega), or by subcloning the amplified cDNAs into pBluescript (Stratagene, La Jolla, CA) and then sequencing the inserts.

These methods were similar to those described previously (Mattson et al., 1993c; Cheng et al., 1994). Briefly, cells were fixed in 4% paraformaldehyde, with membranes permeabilized by exposure for 5 min to 0.2% Triton X-100 in PBS, and placed in blocking serum (0.5% horse serum). Cells were then exposed to primary antibody overnight at 4°C, followed by incubation for 1 h with biotinylated anti–mouse IgG, a 30-min incubation in the presence of ABC reagent (Vector Labs, Inc., Burlingame, CA), and a 5-min exposure to DAB tetrahydrochloride. Parallel control cultures were processed identically except that the primary antibody was deleted from the procedure. The primary antibodies were affinity-purified rabbit anti– c-Fos (1:500) and anti–c-Jun (1:1,000) (Oncogene Science, Inc., Minecola, NY), and mouse monoclonal anti–Bcl-2 (1:500; Santa Cruz Biotechnology, Santa Cruz, CA). Immunostained cultures were visualized under bright-field optics, and relative levels of immunoreactivity of neuronal cell bodies were scored according to the following scale: 0, no immunoreactivity; 1, weak; 2, moderate; 3, intense; 4, very intense. 100 neurons were scored per culture. All cultures were scored without knowledge of their treatment history. For Western blot analysis, solublized proteins were separated by SDS-PAGE (12% gel), transferred to a nitrocellulose sheet, and immunoreacted overnight with Bcl-2 antibody in the presence of blocking serum (1:500). The blot was then exposed for 1 h to HRP-conjugated secondary antibody (1:10,000; Jackson ImmunoResearch Laboratories Inc., West Grove, PA) and immunoreactive protein was visualized using a chemiluminescence-based detection kit according to the manufacturer's protocol (ECL kit; Amersham Corp., Arlington Heights, IL).

These methods were similar to those described previously (Goodman and Mattson, 1994). Briefly, cells were incubated for 50 min in the presence of 50 μM 2,7-dichlorofluorescin diacetate (DCF; Molecular Probes, Inc., Eugene, OR), and then were washed three times (2 ml per wash) in HBSS containing 10 mM Hepes and 10 mM glucose. Cells were imaged using a confocal laser scanning microscope (Sarastro 2000; Molecular Dynamics) coupled to an inverted microscope (Nikon, Inc., Garden City, NY). Cells were located under visible light and a single image was acquired by scanning with the laser (488-nm excitation and 510-nm barrier filter). The laser intensity and photodetector gain were held constant to allow quantitative comparisons of relative fluorescence intensity of cells between treatment groups. Values of relative DCF fluorescence (average pixel intensity per cell) were obtained using the Imagespace software supplied by the manufacturer (Molecular Dynamics).

Activities of Cu/Zn-SOD, Mn-SOD, and catalase in cultured cells were quantified by methods described previously (Mattson et al., 1995). Cells from 60-mm culture dishes were pelleted by low speed centrifugation and homogenized in 2.5 ml of a nitrogen-purged buffer consisting of 10 mM Hepes, 137 mM NaCl, 4.6 mM KCl, 1.1 mM KH₂PO₄, 0.6 mM MgSO₄ (pH 7.4). The homogenate was centrifuged for 1 h at 100,000 g at 4°C, and the supernatant was used for enzyme assays. For the SOD activity assay, 200 μl of the supernatant was added to 1.48 ml of 68 mM NaH₂PO₄ containing 1.35 mM EDTA (pH 7.8), 100 μl of 4 mM xanthine, and 170 μl of 3.53 mM epinephrine (pH 11.5). After a 5-min incubation at 30°C, 50 μl of xanthine oxidase was added to the cuvette, and the absorbance was followed for 3 min at 320 nm. To control for variability among lots, xanthine oxidase was diluted until an assay mixture without SOD yielded 0.03 A change per min. Mn-SOD activity was taken as the SOD activity remaining after addition of potassium cyanide (200 mM). One unit of SOD activity was defined as the amount that reduced the absorbance change by 50%, and results were expressed as units per mg protein. For the catalase activity assay, cell homogenate supernatant (1.5 ml) was mixed with 18 μl absolute ethanol and incubated on ice for 8 min, and 7.8 μl of 10% Triton X-100 and 2.5 ml of 10 mM NaH₂PO₄ buffer (pH 7.4) were added. Aliquots (0.6 ml) of the solution were added to 20-ml test tubes along with 3.57 ml of cold 6 mM hydrogen peroxide and the tube was vortexed. After 3 min, 0.714 ml of 6 N H₂SO₄ was added and the mixture was vortexed. 5 ml of 0.01 N KMnO₄ was added to the solution, the tube was vortexed, and the absorbance was measured at 480 nm within 60 s using a Spec 21 UV-Vis spectrophotometer (Shimadzu, Inc., Kyoto, Japan). Quantitation was based on the comparison of the tissue samples with a calibration curve of known peroxide concentrations. The activity of the enzyme was expressed as units per mg protein with 1 unit = 1 μmol of H₂O₂ catalyzed per min.

Hippocampal neurons express NMDA and AMPA/kainate receptors and are vulnerable to glutamate-induced excitotoxicity mediated by calcium influx (Choi, 1987; Mattson et al., 1989) and generation of reactive oxygen species (Lafon-Cazal et al., 1993; Dugan et al., 1995; Mattson et al., 1995). Cultured hippocampal neurons are also vulnerable to killing by a variety of oxidative insults including exposure to FeSO₄ (Zhang et al., 1993) and to Aβ (see Yankner, 1996; for review Mark et al., 1996); Aβ- induced death manifests as apoptosis (Loo et al., 1993). Exposure of cultures to 10 μM glutamate, 5 μM FeSO₄, or 5 μM Aβ resulted in killing 60–70% of the neurons (Fig. 1). Pretreatment of cultures with increasing concentrations of CHX for 16 h before exposure to glutamate, FeSO₄, or Aβ resulted in biphasic concentration-dependent increase in neuronal survival. CHX concentrations of 50–100 nM significantly increased neuronal survival such that 80–90% of the neurons survived exposure to each of the three insults (Fig. 1). Concentrations of CHX of ⩾1 μM were ineffective in protecting neurons, and concentrations >1 μM caused significant neuronal death during a 24-h exposure period (Fig. 1 D).

Since the concentrations of CHX that were neuroprotective were below those expected to inhibit protein synthesis maximally, we examined the time course and concentration dependence of CHX effects on incorporation of radiolabeled amino acids into TCA-precipitable protein (Fig. 2). Parallel cultures were exposed for increasing times (1, 4, 12, and 24 h) to either 100 nM or 10 μM CHX. In cultures exposed to 100 nM CHX, levels of protein synthesis were reduced to ∼60% of control levels within 1 h of exposure, recovered to ∼80% of control levels by 4 and 12 h, and then decreased somewhat at 24 h (Fig. 2,A). In contrast, levels of protein synthesis in cultures exposed to 10 μM CHX were reduced to 10% of control levels within 1 h of exposure and remained at this low level through 24 h (Fig. 2 A). In a separate experiment, the concentration dependence of inhibition of protein synthesis by CHX was examined in more detail. Levels of protein synthesis were reduced to 60–75% of control levels in cultures exposed to 10–300 nM CHX for 1 h. CHX concentrations of ⩾1 μM caused a >90% inhibition of protein synthesis.

A characteristic set of genes is rapidly induced when neurons are subjected to toxic insults. Perhaps the most widely studied is the immediated early gene c-fos, which is induced at both the mRNA and protein levels in neural cells subjected to ischemia (Aden et al., 1994; Dragunow et al., 1994), excitotoxins (Ghosh et al., 1994), oxidative insults (Naveilhan et al., 1994; Goldstone et al., 1995), and trophic factor withdrawal (Mesner et al., 1995). We previously used RT-PCR approaches to examine expression of immediate early genes after NGF withdrawal from cultured sympathetic neurons (Estus et al., 1994), and after exposure to Aβ in cultured rat cortical neurons (Estus et al., 1995). To gain insight into the neuroprotective mechanism of CHX, we determined the effects of CHX on levels of mRNA encoding c-Fos, c-Jun, and the antiapoptotic gene Bcl-2. CHX induced a concentration-dependent increase in levels of bcl-2, c-fos, and c-jun mRNAs (Fig. 3). In the case of bcl-2 mRNA, a twofold increase occurred in cells exposed to 30 nM CHX, and the peak increase of approximately fivefold occurred in cultures exposed to 100 nM CHX. Higher concentrations of CHX were ineffective in increasing bcl-2 mRNA levels. In the cases of c-fos and c-jun mRNAs, the concentration–response curves were also biphasic and exhibited similar profiles with peak increases occurring in cultures exposed to 100–300 nM CHX (Fig. 3). Similar results were obtained in two additional concentration–response experiments using separate cultures. The time course of changes in bcl-2, c-fos and c-jun mRNAs after exposure to 100 nM CHX is shown in Fig. 3 C. The mRNA levels for each gene increased within 10 min of exposure to CHX, peaked by 30 min, and remained elevated for at least 8 h.

To provide insight into whether the increase of immediate early gene mRNA levels resulted from increased transcription or stabilization of mRNA, we examined levels of c-fos heteronuclear RNA (hnRNA) in cultures exposed to increasing concentrations of CHX. At the CHX concentrations that were most effective in protecting neurons against excitotoxicity and oxidative insults (100–300 nM), CHX induced a clear increase in the level of c-fos hnRNA (Fig. 4). Lower and higher concentrations of CHX had little or no effect on c-fos hnRNA levels. Because the concentrations of CHX effective in protecting neurons against glutamate toxicity and oxidative insults were below concentrations that inhibit protein synthesis maximally, and because c-fos and bcl-2 are induced by neurotrophic factors (Marsh et al., 1993; Downen et al., 1993) that can protect neurons against some of the same insults examined in the present study (Mattson et al., 1993b,c, 1995), we determined the effects of CHX on levels of Bcl-2, c-Fos, and c-Jun proteins. Cultures were exposed for 4 h to increasing concentrations of CHX, and then were immunostained with antibodies to Bcl-2, c-Fos, and c-Jun. Relative levels of neuronal immunoreactivity were quantified (see Materials and Methods). Photomicrographs of control and CHXtreated cells immunostained with bcl-2 and c-fos antibodies are shown in Fig. 5,A. CHX induced concentration- dependent increases in levels of Bcl-2, c-Fos, and c-Jun, with maximum increases occurring in neurons exposed to 100 nM CHX (Fig. 5 B). Higher concentrations of CHX did not increase levels of any of the three proteins.

Since neuroprotective concentrations of CHX induced increased levels of bcl-2 mRNA and protein, and since bcl-2 is known to have antioxidant functions (Hockenbery et al., 1993; Kane et al., 1993), we determined whether pretreatment of hippocampal cultures with CHX would reduce levels of oxidative stress in neurons exposed to FeSO₄. Cultures were pretreated with vehicle or 100 nM CHX, and then exposed to FeSO₄ for 20 min. Levels of peroxides were quantified using the DCF probe, and confocal laser scanning microscopy (Goodman and Mattson, 1994). FeSO₄ induced a large increase in peroxide levels (Fig. 6, A and B). The FeSO₄-induced increase of peroxide was attenuated in neurons pretreated with 100 nM CHX (Fig. 6 B), but not with higher or lower concentrations of CHX (10 nM and 1 μM; data not shown), demonstrating that a neuroprotective concentration of CHX reduces oxidative stress.

If the mechanism whereby CHX protected neurons involved enhancement of antioxidant pathways, then depletion of antioxidants should prevent protection by CHX. Buthionine sulfoximine (BSO) is an agent that causes glutathione depletion by irreversibly inhibiting γ-glutamylcysteine synthase, an enzyme required for glutathione synthesis (Griffith and Meister, 1979). Hippocampal cultures were pretreated with vehicle or 100 nM CHX for 16 h, and then were exposed for 20 h to glutamate alone or in combination with 150 μM BSO. BSO significantly reduced the excitoprotective effect of CHX (Fig. 6 C). BSO alone was not toxic during a 20-h exposure period.

Previous studies used antisense approaches to demonstrate roles for Bcl-2 in prevention of cell death in several cell death paradigms (Kitada et al., 1994; Allsopp et al., 1995; Keith et al., 1995). In preliminary studies we showed that exposure of cultured hippocampal neurons to 10–50 μM of a Bcl-2 antisense ODN for 16 h resulted in a decrease in levels of Bcl-2 protein as determined by immunocytochemistry (Fig. 7,A) and Western blot analysis (Fig. 7,B). The ability of 100 nM CHX to protect neurons against glutamate toxicity in Bcl-2 antisense ODN-pretreated cultures and control (missense ODN- or vehicle-pretreated cultures) was then determined. Pretreatment with Bcl-2 antisense ODN significantly reduced the neuroprotective action of CHX, whereas the control ODN had no effect on the neuroprotective efficacy of CHX (Fig. 8). 20–35% of the neurons survived exposure to glutamate in the absence of CHX and in cultures pretreated with Bcl-2 antisense ODN plus CHX, whereas 50–65% of the neurons survived in cultures pretreated with CHX alone or CHX plus control ODN. Bcl-2 antisense and missense ODNs (Fig. 8), and sense and nonsense ODNs (data not shown), alone had no effect on neuronal survival during a 48-h exposure period.

Several different neurotrophic factors that protect neurons against the kinds of oxidative insults examined in the present study have been shown to induce the expression of one or more antioxidant enzymes in several types of neurons including hippocampal neurons (Jackson et al., 1994; Mattson et al., 1995). Since CHX suppressed accumulation of H₂O₂ and toxicity of oxidative insults, we quantified activities of the antioxidant enzymes catalase, Cu/Zn-SOD, and Mn-SOD in control hippocampal cultures and in cultures pretreated for 24 h with increasing concentrations of CHX. Activities of all three antioxidant enzymes were significantly increased in cultures treated with 100–500 nM CHX (Fig. 9). Lower or higher concentrations of CHX did not affect antioxidant enzyme activities.

The ability of CHX to protect cultured hippocampal neurons against the toxicities of glutamate, FeSO₄, and Aβ is in general agreement with previous reports in various cell culture models. CHX protected cultured retinal ganglion cells against NMDA toxicity (Dreyer et al., 1995), protected PC12 cells against glutamate toxicity (Serghini et al., 1994), and protected cultured cortical neurons against oxidative injury (Ratan et al., 1994b) and Aβ toxicity (Takashima et al., 1993). In such prior studies, a single concentration of CHX was usually used and concentration–effect analyses were not performed. Our data clearly show that the neuroprotective actions of CHX in hippocampal cultures are concentration dependent with significant effects being observed within the range of 50–500 nM CHX. It was reported that only intermediate doses of CHX protect neurons against ischemic injury in adult rats in vivo (Tortosa et al., 1994), a finding consistent with our cell culture data. Moreover, Oguchi et al. (1994) directly demonstrated that low concentrations of CHX that have a negligible effect on protein synthesis can induce a large increase in inducible nitric oxide synthase mRNA levels in cultured macrophages. Several lines of evidence in the present study suggest that the neuroprotective actions of CHX were not the result of inhibition of protein synthesis. First, the most effective concentrations of CHX (50–100 nM) were considerably lower than the concentrations required to inhibit protein synthesis maximally in most cell types and, in our cultures, caused only a 40% reduction in incorporation of radiolabeled amino acids into TCA-precipitable protein. Second, higher micromolar concentrations of CHX that inhibited protein synthesis maximally were ineffective in protecting neurons against the excitotoxic and oxidative insults. Third, neuroprotective concentrations of CHX induced increases in levels of Bcl-2, c-Fos, and c-Jun protein (and their encoding mRNAs); increased production of these proteins argues against a general inhibition of protein synthesis at those CHX concentrations. Finally, neuroprotective concentrations of CHX induced increased activity levels of the antioxidant enzymes catalase, Cu/ZnSOD, and Mn-SOD.

It is increasingly recognized that apoptosis and necrosis often represent different manifestations of cell death that, nevertheless, share common underlying mechanisms. Indeed, the same initiator of cell death can kill the same cell type by either apoptosis or necrosis depending upon intensity/duration parameters of the insult (Ankarcrona et al., 1995). In the present study CHX protected hippocampal neurons against three different insults that are known to engender both apoptosis and necrosis. Our previous studies (Mattson et al., 1989; Mark et al., 1995) and unpublished data from studies of rat hippocampal cell cultures (M.P. Mattson and R.J. Mark) indicate that (at the concentrations used in the present study) glutamate and FeSO₄ kill neurons mainly by necrosis, whereas Aβ induces mainly apoptotic death (see also Loo et al., 1993). The mechanisms of toxicities of these three insults have been well characterized in previous studies. Glutamate induces calcium influx through NMDA receptors and voltage- dependent channels, and the elevated calcium levels induce production of superoxide anion radical (Lafon-Cazal et al., 1993) and H₂O₂ (Mattson et al., 1995), probably by disrupting mitochondrial transmembrane potential (Mattson et al., 1993d). FeSO₄ kills neurons by inducing hydroxyl radical production and lipid peroxidation; activation of NMDA receptors also contributes to FeSO₄ neurotoxicity in hippocampal cell cultures (Zhang et al., 1993). Aβ induces membrane oxidation (Butterfield et al., 1994; Behl et al., 1994), which impairs function of ion- motive ATPases (Mark et al., 1995, 1997a) and glucose (Mark et al., 1997b) and glutamate (Keller et al., 1997) transporters, resulting in membrane depolarization and calcium influx; both free radicals and calcium appear to contribute to the neurotoxicity of Aβ (Mark et al., 1996). Our data indicate that the mechanism whereby CHX protects neurons against these different insults is by enhancing antioxidant defense systems. As evidence, we found that neuroprotective concentrations of CHX induce increased levels of Bcl-2. Moreover, exposure of hippocampal cultures to Bcl-2 antisense oligonucleotides reduced Bcl-2 levels and abrogated the neuroprotective effects of CHX. These findings suggest that the increase in Bcl-2 induced by neuroprotective concentrations of CHX was mechanistically linked to neuroprotection. Many different studies of both apoptotic and necrotic cell death paradigms have shown that Bcl-2 can protect cells against oxidative insults (Hockenbery et al., 1993; Fukunaga-Johnson et al., 1995; Kane et al., 1995). Measurements of levels of reactive oxygen species including H₂O₂ have shown that expression of Bcl-2 is correlated with reduced levels of oxidative stress in cells exposed to oxidative insults (Kane et al., 1993). Consistent with a role for Bcl-2 in the neuroprotective actions of CHX, we found that pretreatment of hippocampal neurons with neuroprotective concentrations of CHX resulted in a significant attenuation of FeSO₄-induced accumulation of cellular H₂O₂. In addition, we found that activity levels of three different antioxidant enzymes were increased in CHX-treated cultures. It is known that oxidative stress itself can induce expression of immediate early gene products and antioxidant enzymes (Goldstone et al., 1995). However, we did not detect an increase in levels of H₂O₂ in cultured hippocampal neurons exposed to neuroprotective concentrations of CHX, suggesting that CHX was likely acting by a more direct effect on expression of the antioxidant enzymes. Another potential mechanism by which CHX might increase resistance to oxidative insults is suggested by the work of Ratan et al. (1994b) who showed that CHX can enhance cellular antioxidant pathways by shunting cysteine from protein synthesis to glutathione. Consistent with the latter mechanism, we found that BSO, which depletes cellular glutathione levels, significantly reduced the excitoprotective efficacy of CHX. The cumulative data therefore suggest that there may be several mechanisms, each involving enhancement of antioxidant pathways, whereby CHX protects neurons against excitotoxic and oxidative insults. Our data suggest that, in the cell death paradigms examined in the present study, the neuroprotective mechanism of CHX involves induction of cytoprotective gene products involved in antioxidant pathways rather than suppression of killer gene products. However, because the neuroprotective concentrations of CHX did cause a modest 20% decrease in overall protein synthesis, we cannot completely rule out the possibility that a selective inhibition of production of death genes occurred under those conditions.

The similarities between the neuroprotective actions of CHX demonstrated in the present study and the neuroprotective actions of neurotrophic factors documented in prior studies (for review see Mattson et al., 1993b) are striking. Pretreatment of hippocampal cell cultures with bFGF, NGF, and BDNF protected neurons against glutamate toxicity and suppressed accumulation of reactive oxygen species (Mattson et al., 1989; Cheng and Mattson, 1994; Mattson et al., 1995). bFGF and NGF also protected cultured hippocampal neurons against FeSO₄ toxicity (Zhang et al., 1993). bFGF (Mattson et al., 1993b) and BDNF (Mattson, M.P., unpublished data) protected cultured hippocampal neurons against Aβ toxicity. Each of these neurotrophic factors has been shown to induce the expression of one or more gene products linked to resistance to oxidative insults. Examples include: bFGF induced Cu/Zn-SOD and glutathione reductase in hippocampal cell cultures and protected neurons against glutamate toxicity (Mattson et al., 1995); NGF induced catalase expression in PC12 cells and protected them against H₂O₂ toxicity (Jackson et al., 1994); BDNF induced glutathione peroxidase in striatal neurons and protected them against metabolic/excitotoxic insults (Spina et al., 1992); and PDGF induced a doubling of catalase and Cu/Zn-SOD activity levels in hippocampal cultures and protected neurons in those cultures against FeSO₄- and glucose deprivation–induced injury (Cheng et al., 1995). None of the latter studies examined the effects of the trophic factors on Bcl-2 levels. However, recent data suggest that BDNF can induce bcl-2 expression in neurons that are dependent upon BDNF for survival, and that Bcl-2 may mediate the survival response to BDNF (Allsopp et al., 1995). Since we found that Bcl-2 antisense blocked the neuroprotective effects of CHX, the contribution of CHXinduced increases in antioxidant enzyme activities to the neuroprotective action of CHX remains to be established.

While it is well documented that CHX can increase levels of immediate early gene mRNAs (Greenberg and Ziff, 1984), our demonstrations of increased levels of immediate early gene proteins and antioxidant enzyme activities are novel findings. The mechanism whereby CHX induces increased levels of immediate early gene products and antioxidant enzymes may involve stimulation of transcription and/or stabilization of mRNAs. We found that neuroprotective concentrations of CHX induced an increase in c-fos hnRNA, indicating that, at least for this gene, CHX can induce transcription. Consistent with our findings are previous data showing that CHX can induce activation of transcription factors (Greenberg and Ziff, 1984; Subramaniam et al., 1989; Zinck et al., 1995). On the other hand, CHX has also been shown to reduce degradation of certain mRNAs (Edwards and Mahadevan, 1992) and could, in that way, increase mRNA levels. Such actions of CHX at concentrations below those that inhibit protein synthesis could, in theory, result in increased protein synthesis. Indeed, our data suggest that levels of c-Fos, c-Jun, and Bcl-2 were increased in cultures exposed to neuroprotective concentrations of CHX.

Taken together with previous findings, the present data suggest that CHX can suppress cell death by two quite different mechanisms, one involving suppression of production of presumptive death genes, and the other involving induction of expression of cytoprotective gene products. Some prior studies have provided strong evidence that CHX can suppress cell death by inhibiting protein synthesis. For example, in sympathetic neurons where CHX acts at higher concentrations, CHX may suppress cell death by inhibiting synthesis of killer gene products (Martin et al., 1988, 1992). However, in most studies that have used CHX to block cell death, the extent to which the cytoprotective concentrations of CHX suppressed protein synthesis was not determined. Moreover, in in vivo studies of cerebral ischemia and other insults that have documented neuroprotective actions of CHX, no information was obtained concerning levels of protein synthesis in the “saved” neurons (e.g., Goto et al., 1990; Linnik et al., 1993; Svendsen et al., 1994; Tortosa et al., 1994). The collective data from the present study demonstrate that CHX protects cortical neurons against insults relevant to the pathogenesis of ischemic and excitotoxic brain injury by inducing production of antioxidant gene products. Therefore, we suggest that the fact that CHX protects neurons against a particular insult does not justify the conclusion that death genes are involved in the cell death process. It will be necessary to rigorously evaluate the mechanism of neuroprotection by CHX in each paradigm and, ultimately, to identify specific life gene products that are induced, or death gene products that are suppressed.

We thank J. Begley, S. Bose, and R. Pelphrey for technical assistance.

This work was supported by grants to M.P. Mattson from the National Institutes of Health (NIH) (NS29001 and NS30583), the Metropolitan Life Foundation, and the Kentucky Spinal Cord and Head Injury Trust, and by grants to S. Estus from the NIH and the Alzheimer's Association.

Aβ

amyloid β-peptide

BDNF

brainderived neurotrophic factor

bFGF

basic FGF

BSO

buthionine sulfoximine

CHX

cycloheximide

DCF

2,7-dichlorofluorescein diacetate

hnRNA

heteronuclear RNA

NMDA

N-methyl-d-aspartate

ODN

oligodeoxynucleotide

RT

reverse transcription

SOD

superoxide dismutase