Address correspondence and reprint requests to Dr. A. Almeida at Departamento de Bioquímica y Biología Molecular, Universidad de Salamanca, Edificio Departamental, Plaza Doctores de la Reina, s/n, 37007 Salamanca, Spain. E-mail: email@example.com
Abstract: The possible neuroprotective effect of D-glucose against glutamate-mediated neurotoxicity was studied in rat cortical neurons in primary culture. Brief (5-min) exposure of neurons to glutamate (100 μM) increased delayed (24-h) necrosis and apoptosis by 3- and 1.8-fold, respectively. Glutamate-mediated neurotoxicity was accompanied by a D-(-)-2-amino-5-phosphonopentanoate (100 μM) and Nω-nitro-L-arginine methyl ester (1 mM)-inhibitable, time-dependent ATP depletion (55% at 24 h), confirming the involvement of NMDA receptor stimulation followed by nitric oxide synthesis in this process. Furthermore, the presence of D-glucose (20 mM), but not its inactive enantiomer, L-glucose, fully prevented glutamate-mediated delayed ATP depletion, necrosis, and apoptosis. Succinate- cytochrome c reductase activity, but not the activities of NADH-coenzyme Q1 reductase or cytochrome c oxidase, was inhibited by 32% by glutamate treatment, an effect that was abolished by incubation with D-glucose. Lactate accumulation in the culture medium was unmodified by any of these treatments, ruling out the possible involvement of the glycolysis pathway in either glutamate neurotoxicity or D-glucose neuroprotection. In contrast, D-glucose, but not L-glucose, abolished glutamate-mediated glutathione oxidation and NADPH depletion. Our results suggest that NADPH production from D-glucose accounts for glutathione regeneration and protection from mitochondrial dysfunction. This supports the notion that the activity of the pentose phosphate pathway may be an important factor in protecting neurons against glutamate neurotoxicity.
Glutamate is the main excitatory neurotransmitter in the mammalian CNS, but overstimulation of its postsynaptic receptors, mainly the NMDA subtype, causes neurodegeneration (Choi, 1987). Thus, an NMDA receptor-mediated increase in free cytosolic Ca2+ concentrations stimulates neuronal nitric oxide (.NO) synthase (nNOS) activity (Garthwaite et al., 1988), leading to neurotoxicity (Schulz et al., 1995). The mechanism of this toxicity appears to be associated with energy depletion, possibly because of the .NO and/or peroxynitrite (ONOO-)-induced DNA damage leading to poly(ADP-ribose) synthetase activation (Zhang et al., 1994), lipid peroxidation (Radi et al., 1991b), and mitochondrial dysfunction (for review, see Bolaños et al., 1997).
Although the exact mechanism responsible for .NO-mediated mitochondrial dysfunction and cell death is still a matter of debate, an increasing body of evidence now suggests that the intracellular concentration of glutathione (GSH) may be a key factor in determining cellular vulnerability (Murphy et al., 1989; Almeida et al., 1998; Clementi et al., 1998). Therefore, up-regulation of GSH synthesis and/or GSH regeneration from oxidized glutathione (GSSG) may be important mechanisms for protecting neural cells against .NO-mediated toxicity. In this sense, we have recently shown that astrocytes are able to up-regulate NADPH concentrations through the transcriptional induction of glucose-6-phosphate dehydrogenase, the rate-limiting step in the pentose phosphate pathway (García-Nogales et al., 1999). This mechanism appears to contribute to GSH regeneration from GSSG at the expense of NADPH under excessive endogenous .NO biosynthesis. In addition, direct activation of the pentose phosphate pathway by exogenous H2O2 (Ben-Yoseph et al., 1996) or cumene hydroperoxide (Kussmaul et al., 1999) has been suggested to contribute to neuronal resistance to oxidative stress.
In this work, we have studied the ability of neurons in primary culture to prevent cell death upon excess endogenous .NO biosynthesis. We show that the neuronal utilization of D-glucose is neuroprotective against .NO-mediated mitochondrial damage, necrosis, and apoptosis caused by glutamate receptor stimulation. Moreover, the neuroprotective role for D-glucose must be associated with its consumption mainly through the pentose phosphate pathway because it prevents glutamate-mediated NADPH depletion and glutathione oxidation.
MATERIALS AND METHODS
Dulbecco's modified Eagle's medium (DMEM), horse serum, cytosine arabinoside, L-glutamate, glycine, Nω-nitro-L-arginine monomethyl ester (NAME), ubiquinone-5 (coenzyme Q1; CoQ1); 2-vinylpyridine, 1-isobutyl-3-methylxanthine, D-(-)-2-amino-5-phosphonopentanoate (APV), and 4′,6-diamino-2-phenylindole (DAPI) were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.). DMEM was always supplemented with penicillin (100 U/ml), streptomycin (100 μg/ml), and amphotericin B (0.25 μg/ml) (Sigma). Fetal calf serum and cytochrome c were purchased from Boehringer Mannheim (Heidelberg, Germany). Cytochrome c was reduced with sodium ascorbate (Sigma) just before use and passed through Sephadex G-25M (PD-10 columns; Amersham Pharmacia Biotech, Uppsala, Sweden) to remove excess ascorbate. Other substrates, enzymes, and coenzymes were purchased from Sigma, Boehringer, or Merck (Darmstadt, Germany). Plastic tissue culture dishes were purchased from Nunc (Denmark) or Linbro (McLean, VA, U.S.A.).
Albino Wistar rats fed ad libitum on stock laboratory diet were used for the experiments. Rats were maintained at ≈22°C with a 12-h light/dark cycle. Virgin females weighing 210-250 g were caged overnight with males, and conception was confirmed the next morning by the presence of spermatozoa in vaginal smears.
Cerebral cortex neurons in primary culture were prepared from fetal rats at 16-17 days of gestation (Almeida et al., 1998). Dissociated cell suspensions were plated at a density of 2.5 × 105 cells/cm2 in either 2-, 4-, 9.6-, or 28.2-cm2 plastic Petri dishes coated with poly-D-lysine in DMEM supplemented with 10% fetal calf serum. Cells were incubated at 37°C in a humidified atmosphere containing 5% CO2/95% air. Fortyeight hours after plating, the medium was replaced with DMEM supplemented with 5% horse serum and 20 mM D-glucose. On day 4 of culture, cytosine arabinoside was added at a final concentration of 10 μM to prevent nonneuronal proliferation, and neurons were used on day 9.
Exposure of neurons to glutamate
On day 9 in culture, the medium was removed, neurons were washed once with prewarmed (37°C) buffered Hanks' solution (5.26 mM KCl, 0.43 mM KH2PO4, 132.4 mM NaCl, 4.09 mM NaHCO3, 0.33 mM Na2HPO4, 5.44 mM glucose, 2 mM CaCl2, and 20 mM HEPES, pH 7.4), and preincubated for 5 min at 37°C in the absence (control) or presence of APV (100 μM) or NAME (1 mM) in buffered Hanks' solution. Immediately after, L-glutamate was added at a final concentration of 100 μM (plus 10 μM glycine), and cells were further incubated for 5 min at 37°C. After this incubation, the buffer was aspirated, replaced with DMEM alone (which contains 5.5 mM D-glucose), DMEM supplemented with 20 mM D-glucose, or DMEM supplemented with 20 mM L-glucose, and the cells were further incubated for 3 or 24 h at 37°C.
Determination of neuronal necrosis and apoptosis
Necrosis was assessed by examination of trypan blue-staining cells (Koh and Choi, 1988). In brief, 24 h after glutamate exposure, neuronal cultures were washed with warm (37°C) phosphate-buffered saline (PBS; 136 mM NaCl, 2.7 mM KCl, 7.8 mM Na2HPO4, 1.7 mM KH2PO4, pH 7.4) and incubated with 0.2% trypan blue in PBS for 2 min at room temperature. Microphotographs (20× magnification; Diaphot, Nikon) were taken for each experimental condition, and viable and necrotic (stained) cells were counted. Apoptosis was assessed by staining the nuclei of cells with DAPI, a membrane-permeable fluorescent dye that binds DNA and allows the quantification of apoptotic neurons, that is, neurons displaying fragmented or condensed nuclei (Mailly et al., 1999). In brief, 24 h after glutamate exposure, neuronal cultures were washed with warm (37°C) PBS and fixed with 4% (wt/vol) paraformaldehyde in PBS for 30 min at room temperature. After washing with PBS, cells were exposed to 3 μM DAPI for 10 min at room temperature and then washed twice with PBS. Neurons were scored for chromatin condensation by fluorescence microscopy (fluorescein filter 330- to 380-nm excitation; 30× magnification). Total and apoptotic nuclei were counted. In all cases, ∼600-1,000 cells were counted per well by an observer author blind to the protocol design. At least three different cell cultures utilizing four separate wells were employed, so that a minimum of 7,200-12,000 neurons were counted for each data point.
Enzyme activity determinations
For the determination of mitochondrial respiratory chain complex activities, neurons plated on 28.2-cm2 Petri dishes were used. Following glutamate exposure and the 24-h incubation period, neuronal cultures were washed with ice-cold PBS and the surviving cells were collected by trypsinization and resuspended in 300 μl of 0.1 M potassium phosphate buffer (pH 7.0). Cell suspensions (containing ∼3-4 mg of protein/ml) were frozen and thawed three times to ensure cell lysis. Enzyme activities were determined in the cell lysates using a Hitachi U2000 spectrophotometer (Hitachi, Tokyo, Japan). NADH-CoQ1 reductase (complex I; EC 22.214.171.124) activity was measured as described by Ragan et al. (1987). The activity of succinate-cytochrome c reductase (complex II-III) was determined following the method of King (1967). Cytochrome c oxidase (complex IV; EC 126.96.36.199) activity was determined as described by Wharton and Tzagoloff (1967). All enzyme activities were expressed as nanomoles per minute per milligram of protein, except for cytochrome c oxidase, which was expressed as the first-order rate constant (k per minute per milligram of protein).
For ATP determinations, neurons plated in 2-cm2 wells and treated as above were rapidly washed with ice-cold PBS, scraped off with 2 × 0.5 ml of 0.3 M HClO4, and neutralized with 0.5 ml of 2 M KHCO3 at pH 6.5. The perchlorate precipitate was removed by centrifugation, and ATP was determined in the supernatants by chemiluminescence using a commercially available kit (Sigma) following the manufacturer's instructions.
For cyclic GMP determinations, neurons plated in 9.6-cm2 wells were treated as above, except that the phosphodiesterase inhibitor 1-isobutyl-3-methylxanthine (1 mM) was included throughout the experiments. After a 5-min incubation period, the buffer was aspirated, cells were scraped off with 2 × 0.5 ml of ice-cold ethanol, dried, and used for cyclic GMP determination using a commercially available radioimmunoassay kit (Amersham Pharmacia Biotech), following the manufacturer's instructions.
For glutathione determinations, neurons plated in 9.6-cm2 well plates and treated as above were washed with ice-cold PBS and immediately collected by scraping off with 0.5 ml of 1% (wt/vol) sulfosalicylic acid. Cell lysates were centrifuged at 13,000 g for 5 min at 4°C, and the supernatants used for glutathione determinations on the same day. GSx (the amount of GSH plus two times the amount of GSSG) and GSSG were measured exactly as previously described (Dringen and Hamprecht, 1996; García-Nogales et al., 1999). GSH and GSSG concentrations were expressed as nanomoles per milligram of protein.
For lactate determinations, neurons plated in 2-cm2 wells were treated as above, and the media were collected for lactate determination as described by Gutmann and Wahlefeld (1974).
NADPH concentrations were measured as previously reported (García-Nogales et al., 1999). In brief, neurons plated in 4-cm2 wells were treated as above, washed with ice-cold PBS, and collected in 250 μl of 0.5 M KOH in 50% (vol/vol) ethanol. Aliquots (200 μl) of the cell lysates were neutralized (pH 7.8) with 200 μl of 0.5 M triethanolamine/0.5 M potassium phosphate and centrifuged at 13,000 g for 2 min at 4°C. A 50-μl aliquot of the supernatant was immediately used for NADPH determination by chemiluminescence as previously described (Wulff, 1985), with the exception that NADH was oxidized by incubation of the samples with 0.5 mU/μl lactate dehydrogenase and 1 mM pyruvate (Klingerberg, 1985).
Proteins were determined either in the cell suspensions, in lysates, or in parallel cell culture incubations after solubilization with 0.1 M NaOH. Protein concentrations were determined by the method of Lowry et al. (1951) using bovine serum albumin as standard.
Measurements from individual cultures were performed in triplicate, and the results are expressed as the mean ± SEM values for the number of culture preparations indicated in the legends of the figures and tables. Statistical analysis of the results was determined by one-way analysis of variance followed by the least significant difference multiple range test. In all cases, p < 0.05 was considered significant.
A brief exposure of neurons to glutamate leads to NMDA receptor-mediated .NO formation and delayed energy depletion and neurotoxicity
In this study, the exposure of neurons to glutamate (100 μM) was performed for only 5 min. To confirm that this short-term treatment was sufficient to activate NMDA receptor and nNOS activity, we measured the intracellular concentrations of cyclic GMP, a well-known index of endogenous .NO formation (Garthwaite et al., 1988). As shown in Table 1, a significant increase in neuronal cyclic GMP concentrations was observed after 5 min of glutamate exposure. Furthermore, the glutamate-mediated increase in cyclic GMP level was antagonized with either APV (100 μM), a competitive antagonist of NMDA glutamate-subtype receptors, or NAME (1 mM), an inhibitor of NOS (Table 1). It should be noted that cyclic GMP values in glutamate plus NAME-treated cells were below the control values, indicating the importance for endogenous nNOS activity of these neurons even under control conditions.
Table 1. Glutamate-mediated cyclic GMP formation in neurons in primary culture
Glutamate + APV
Glutamate + NAME
Neurons were exposed to 100 μM glutamate for 5 min at 37°C in either the absence or the presence of 100 μM APV or 1 mM NAME. Cyclic GMP concentrations were measured as described in Materials and Methods. Values are means ± SEM from four or five different culture preparations.
aSignificantly different when compared with the control group.
bSignificantly different when compared with the glutamate group.
The short-term effect of glutamate on .NO biosynthesis was accompanied by a subsequent time-dependent depletion of ATP concentrations. Thus, as shown in Fig. 1, ATP concentrations were decreased by 14% after 3 h and by 55.8% after 24 h of incubation in DMEM without glutamate; these effects were prevented by preincubation with either APV or NAME. Moreover, glutamate treatment caused delayed neurotoxicity, as assessed by appropriate dye staining and counting of either necrotic cells or apoptotic nuclei (see in Fig. 2 the results of counting by an observer author blind to the protocol design). Control cultures already showed a measurable proportion (as compared with the total number of cells) of necrotic (15.3 ± 5.3%) and apoptotic (7.9 ± 1.1%) cells. However, glutamate exposure significantly increased these proportions to 48.0 ± 3.8 and 14.4 ± 1.6% for necrotic and apoptotic cells, respectively (Fig. 2).
D-Glucose prevents glutamate-mediated mitochondrial damage, energy depletion, and neurotoxicity
We were prompted to investigate whether D-glucose (20 mM) was neuroprotective against glutamate neurotoxicity. For this purpose, neurons were first stimulated for 5 min with glutamate, washed, and incubated for 3 or 24 h in DMEM supplemented (or not) with D-glucose (20 mM). Therefore, DMEM supplemented with 20 mM glucose actually contained 25.5 mM D-glucose, a concentration that has been used by others for the long-term maintenance of neuronal cultures (Dawson et al., 1993). Control cells never received glutamate and were incubated in DMEM for the 3- and 24-h periods. As shown in Fig. 1, the presence of D-glucose for 24 h abolished the decrease in ATP concentrations caused by glutamate treatment, an effect that was not observed after 3 h. The presence of L-glucose (20 mM), the biologically inactive enantiomer of D-glucose, did not prevent ATP depletion at 3 or 24 h. Moreover, D-glucose, but not L-glucose, treatment fully prevented the increase in the number of necrotic and apoptotic cells as compared with glutamate-exposed cells (Fig. 2).
To address the possibility that the restoration of ATP concentrations by D-glucose could be accounted for by a compensated increased glycolytic rate, we evaluated the possible accumulation of lactate in the culture medium. However, as shown in Table 2, lactate concentrations remained unchanged under all the circumstances studied, ruling out the possible contribution of this pathway to the observed maintenance of ATP levels by D-glucose. On the other hand, a 5-min exposure of glutamate to neurons inhibited succinate-cytochrome c reductase activity by 32% after 24 h, without affecting those of NADH-CoQ1 reductase or cytochrome c reductase (Table 3), confirming our previous observations (Almeida et al., 1998). Interestingly, the presence of D-glucose during the 24-h period following glutamate exposure completely restored the inhibition of this mitochondrial complex activity (Table 3).
Table 2. Lactate released to culture medium after exposure of neurons in primary culture to glutamate
Lactate released (μmol/mg of protein)
Neurons were exposed to 100 μM glutamate for 5 min at 37°C, and the cells were incubated in DMEM, DMEM + 20 mM D-glucose, or DMEM + 20 mM L-glucose at 37°C for a further 3 or 24 h. Lactate concentrations were determined in the medium as described in Materials and Methods. Values are means ± SEM from three different culture preparations.
1.3 ± 0.1
7.7 ± 0.2
1.2 ± 0.1
7.5 ± 0.3
Glutamate + D-glucose
1.5 ± 0.1
7.3 ± 0.3
Glutamate + L-glucose
1.3 ± 0.1
6.9 ± 0.4
Table 3. Effect of glutamate exposure on activity of mitochondrial respiratory chain complexes in neurons in primary culture
NADH-CoQ1 reductase (nmol/min/mg of protein)
Succinate-cytochrome c reductase (nmol/min/mg of protein)
Cytochrome c oxidase (k/min/mg of protein)
Neurons were exposed to 100 μM glutamate for 5 min at 37°C, and the cells were incubated in DMEM or DMEM + 20 mM D-glucose at 37°C for a further 24 h. Enzyme activities were determined in the cells as described in Materials and Methods. k is the first rate constant for cytochrome c oxidation. Values are expressed as means ± SEM from three different culture preparations.
aSignificantly different when compared with the control group.
bSignificantly different when compared with the glutamate group.
D-Glucose prevents glutamate-mediated glutathione oxidation and NADPH depletion
As glycolysis cannot account for the maintenance of ATP levels in the presence of D-glucose, even though a protective effect against mitochondrial complex damage was observed, the possible effect of D-glucose in preventing disturbances in antioxidant metabolism could be advanced. To investigate this issue, we determined the intracellular concentrations of GSH and GSSG. As shown in Fig. 3, even though GSH concentrations were unchanged, those of GSSG increased significantly by 16% after glutamate treatment. This effect was prevented by D-glucose but not by L-glucose, strongly suggesting that glucose metabolism, rather than its possible role as a free radical scavenger, is a necessary factor in this effect. Accordingly, the possible metabolism of D-glucose through the pentose phosphate pathway could be proposed. We therefore determined the intracellular concentrations of NADPH as one product of this pathway. As shown in Fig. 4, glutamate treatment was accompanied by a significant 20% decrease in NADPH concentrations after 24 h of incubation. Interestingly, incubation of the cells in the presence of D-glucose, but not L-glucose, fully prevented NADPH depletion (Fig. 4).
The primary cultured neurons used in this work seemed to be mature, because exposure to a low glutamate concentration rapidly increased cyclic GMP synthesis and caused delayed neurotoxicity and ATP depletion. These phenomena were completely abolished by either NMDA receptor antagonism or nitric oxide synthesis inhibition, suggesting that glutamate-mediated neurotoxicity and ATP depletion would be due to NMDA receptor activation followed by nitric oxide synthesis. Accordingly, this neuronal system appears to be appropriate for the study of possible endogenous mechanisms aimed at preventing glutamate receptor-mediated neurotoxicity.
Glutamate-mediated neurotoxicity was reflected mainly in necrosis, as trypan blue-stained cells increased by ∼3-fold, whereas apoptotic cells (showing condensed or fragmented DAPI-stained nuclei) increased by only 1.8-fold as compared with control cells. These results partially confirm those reported by Ankarcrona et al. (1995), who showed that glutamate neurotoxicity occurs through necrosis or apoptosis depending on whether mitochondrial function is damaged (after a “severe” glutamate insult) or undamaged (after a “mild” glutamate insult), respectively. Under our conditions, “mild” glutamate treatment led to “severe” mitochondrial dysfunction, as judged by the delayed 32% inhibition of succinate-cytochrome c reductase and the 55% decrease in ATP concentrations. Despite this, a slight but significant increase in apoptosis was observed. However, it should be mentioned that necrosis was the main neurotoxic mechanism, in keeping with the idea of Ankarcrona et al. (1995).
To study possible endogenous pathways protecting against glutamate neurotoxicity, we decided to increase the concentration of D-glucose in the DMEM immediately after glutamate treatment. The presence of D-glucose (20 mM) completely abolished the glutamate-mediated increase in necrotic and apoptotic cells and elicited full protection against neuronal ATP depletion. In contrast to D-glucose, the presence of the inactive enantiomer, L-glucose, which is not recognized by hexose transporters, was not able to prevent glutamate-mediated neurotoxicity or ATP depletion. Lizasoain et al. (1996) have previously shown that D-glucose is able to directly scavenge ONOO- in submitochondrial particles, suggesting that in the intact cell, this mechanism might exert a protective effect against ONOO--mediated mitochondrial dysfunction. However, under our conditions, L-glucose, a compound that should scavenge ONOO- in a similar fashion to D-glucose, was not neuroprotective. In this context, it is well known that the addition of membrane-impermeant superoxide dismutase to the extracellular medium prevents glutamate neurotoxicity in primary neurons (Dawson et al., 1991), suggesting that intercellular trafficking of the superoxide anion could account for this type of neurotoxicity (Dawson et al., 1991; Lafon-Cazal et al., 1993). Therefore, it is conceivable that, under our conditions, the lack of protection shown by L-glucose would not be due to a putative lack of free radical-scavenging ability but rather to its inability to enter the cell. Furthermore, the presence of 2-deoxy-D-glucose (20 mM), a compound that does enter the cell, is phosphorylated by hexokinase, but is not further metabolized, was also unable to prevent glutamate-mediated neurotoxicity and energy depletion (results not shown). These results strongly suggest that the neuro-protective action of D-glucose may be indirect and mediated by its uptake and metabolism within the neuron.
Interestingly, the presence of 20 mM D-glucose restored ATP concentrations after 24 h, but not after 3 h, of glutamate exposure, in keeping with the idea that the long-term neuroprotective effect of D-glucose is due to its intracellular uptake and further metabolism. In this context, neurons express both low (Km = 20 mM) and high (Km = 3.5 mM) -affinity glucose transport systems, namely, GLUT1 and GLUT3, respectively (Bell et al., 1993). It is therefore conceivable that, at least under our conditions, D-glucose may be readily taken up by neurons. To elucidate whether D-glucose was utilized in the glycolytic pathway, we measured the concentration of lactate in the culture medium, as its accumulation has been widely regarded as an index of glycolytic rate (Pauwels et al., 1985). This possibility is plausible because mitochondrial inhibition is associated with increased glucose consumption through “anaerobic” glycolysis in several cell types, including neurons (Pauwels et al., 1985). Moreover, a putative increase in the glycolytic rate would account for the observed ATP compensation and recovery from glutamate-mediated cell death. However, lactate concentrations, measured in the culture medium 3 or 24 h after glutamate exposure, were unchanged under all the circumstances studied. This result rules out (1) the possibility that glutamate-mediated ATP depletion would be due to glycolytic inhibition and (2) the notion that the neuroprotective effect of D-glucose would be due to a putative compensated increased glycolysis. This is in agreement with the observation that neurons, unlike astrocytes, do not switch to “anaerobic” glycolysis after antimycin-mediated (Pauwels et al., 1985) or ONOO--mediated (Bolaños et al., 1995) mitochondrial inhibition.
The presence of D-glucose fully prevented glutamate-mediated inhibition of succinate-cytochrome c reductase activity, which might account for its neuroprotective effect. Moreover, the presence of D-glucose, but not L-glucose, abolished the glutamate-mediated increase in GSH oxidation to GSSG. Finally, glutamate treatment was accompanied by NADPH depletion, an effect that was fully prevented by D-glucose but not by L-glucose. As NADPH is a necessary cofactor for GSH regeneration from GSSG by glutathione reductase, our results are consistent with the idea that the neuroprotective effect of D-glucose would be due to its metabolism through the pentose phosphate pathway. This strongly agrees with recent observations suggesting that the availability of hexose for the pentose phosphate pathway may be an important source of NADPH in astrocytes exposed to either cumene hydroperoxide (Kussmaul et al., 1999) or endogenous [UNK]NO (García-Nogales et al., 1999).
In conclusion, this work shows that D-glucose is neuroprotective against glutamate neurotoxicity in neurons in primary culture. This is consistent with the idea that D-glucose is metabolized through the pentose phosphate pathway, hence providing sufficient NADPH for GSH regeneration from GSSG. As GSH reacts with [UNK]NO and/or ONOO- to form GSSG (Radi et al., 1991a), the compensatory effect of D-glucose to regenerate GSH would serve as an endogenous mechanism aimed at preventing glutamate-mediated mitochondrial dysfunction and neurotoxicity. Whether this mechanism is neuroprotective in the intact brain is not known. However, it is interesting to note that postmortem brain samples from Alzheimer's disease patients show decreased glucose transporter GLUT1 and GLUT3 expression (Maher et al., 1994), a fact that could limit glucose utilization in the degenerating neurons.