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Originally published In Press as doi:10.1074/jbc.M402759200 on July 23, 2004

J. Biol. Chem., Vol. 279, Issue 41, 43035-43045, October 8, 2004
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A Mechanism of Sulfite Neurotoxicity

DIRECT INHIBITION OF GLUTAMATE DEHYDROGENASE*

Xin Zhang, Annette Shoba Vincent, Barry Halliwell, and Kim Ping Wong{ddagger}

From the Department of Biochemistry, Faculty of Medicine, National University of Singapore, 8 Medical Drive, Singapore 117597, Singapore

Received for publication, March 11, 2004 , and in revised form, July 22, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Exposure of Neuro-2a and PC12 cells to micromolar concentrations of sulfite caused an increase in reactive oxygen species and a decrease in ATP. Likewise, the biosynthesis of ATP in intact rat brain mitochondria from the oxidation of glutamate was inhibited by micromolar sulfite. Glutamate-driven respiration increased the mitochondrial membrane potential (MMP), and this was abolished by sulfite but the MMP generated by oxidation of malate and succinate was not affected. The increased rate of production of NADH from exogenous NAD+ and glutamate added to rat brain mitochondrial extracts was inhibited by sulfite, and mitochondria preincubated with sulfite failed to reduce NAD+. Glutamate dehydrogenase (GDH) in rat brain mitochondrial extract was inhibited dose-dependently by sulfite as was the activity of a purified enzyme. An increase in the Km (glutamate) and a decrease in Vmax resulting in an attenuation in Vmax/Km (glutamate) at 100 µM sulfite suggest a mixed type of inhibition. However, uncompetitive inhibition was noted with decreases in both Km (NAD+) and Vmax, whereas Vmax/Km (NAD+) remained relatively constant. We propose that GDH is one target of action of sulfite, leading to a decrease in {alpha}-ketoglutarate and a diminished flux through the tricarboxylic acid cycle accompanied by a decrease in NADH through the mitochondrial electron transport chain, a decreased MMP, and a decrease in ATP synthesis. Because glutamate is a major metabolite in the brain, inhibition of GDH by sulfite could contribute to the severe phenotype of sulfite oxidase deficiency in human infants.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Sulfite is formed from sulfur dioxide, an environmental pollutant. It is also generated endogenously by the metabolism of sulfur-containing amino acids such as methionine and cysteine and from sulfate in response to bacterial lipopolysaccharide (1). The most common exogenous source is sulfiting agents used as preservatives in dried fruits and vegetables (2) and in wine where millimolar concentrations have been reported (3). Sulfite is also used as a stabilizer in many drugs administered to patients (4). Interestingly, its presence in a dexamethasone preparation increased the neurotoxicity of excitotoxic agents (5) suggesting that it could have an effect on neuronal cells. Humans can oxidize sulfite () to sulfate () by sulfite oxidase (SO),1 a mitochondrial enzyme. However, the SO activity in human liver was reported to be only 10% that of rat liver (6). Although SO is expressed in human lung (7), its activity was reported to be low (8), which may be why some asthmatic subjects react adversely to sulfite in food or atmospheric sulfur dioxide (9). The comprehensive literature over a 33-year period also shows a relationship between sulfite as food additives and asthma (10).

Sulfite oxidase is a dimeric metallohemoprotein with molybdenum and protoheme as prosthetic groups (11, 12). The catalytic molybdenum centers of SO from various species of animals appear to be identical as examined by electron paramagnetic resonance (13, 14). However, SO activity from human liver was almost 10 to 20 times lower than levels found in rat and chicken liver, and it was suggested that the decreased reactivity of the human enzyme could be due to nonfunctional molybdenum centers (6). A deficiency of SO in humans could be due to a mutation in the SO gene or in any of the several genes encoding the synthesis of molybdopterins (15–18). The associated severe neurological dysfunction characterized by dislocation of ocular lenses, mental retardation, and attenuated growth of the brain suggests that neuronal cells are highly susceptible to sulfite toxicity. Indeed, SO activity measured in whole brain of some laboratory animals was consistently low compared with other tissues (19, 20). Measurement of the expression of SO in human tissues in our laboratory concurred with this observation (21). Four missense mutations were characterized in cell lines from patients with isolated sulfite oxidase deficiency (22). A substitution of G to A of the cDNA of liver SO resulted in an Arg-to-Gln substitution at amino acid residue 190 (23). Another arginine residue (Arg-160) was recently reported to be essential for the binding of sulfite near the molybdenum cofactors in human SO (24), whereas the residue Tyr-343 was proposed to mediate the substrate specificity and catalytic activity of the molybdoprotein (25). Despite the advances made in the molecular biology of SO, there is little information on the mechanism by which accumulation of sulfite affects neuronal function. Cell death was observed in CSM 14.1.4 (a rat neuronal cell line) following exposure to 5 mM sulfite (26). However, the mechanism of toxicity was not elucidated, although free radicals were implicated as increased toxicity of sulfite was observed when intracellular reduced glutathione was compromised (27). One electron oxidation of sulfite would produce a sulfite radical (), capable of damaging DNA, lipids, and proteins (28, 29). In this study, we examined the effects of on rat brain mitochondria and Neuro-2a and PC12 cells and attempted to elucidate its mechanism of action following our earlier observation that micromolar concentrations of produced an increase in reactive oxygen species (ROS) in Madin-Darby canine kidney (MDCK) and opossum kidney (OK) cells. The sulfite-mediated oxidative stress was accompanied by a depletion of intracellular ATP, and this was thought to be due to its inhibitory action on mitochondrial glutamate dehydrogenase (30).


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—The following were from Sigma: ATP, ADP, Ap5A, succinate, glutamate, malate, {alpha}-ketoglutarate, CCCP (carbonylcyanide-m-chlorophenylhydrazone), NAD+, NADH, FL-ASC Bioluminescent Somatic cell assay kit, and glutamate dehydrogenase (type II from bovine liver, EC 1.4.1.3 [EC] , with specific activity of 40 units/mg of protein). Dichlorofluorescein diacetate (DCFDA), JC-1, and MitoTracker Red (CMX-rosamine) were from Molecular Probes. Sodium sulfite was from Merck, Germany. MTT was obtained from BDH Laboratory Supplies, Poole, UK. Neuro-2a, PC12, and human fetal liver cells were kindly provided, respectively, by A/P HP Too and Prof. K. Jeyaseelan of the Dept. of Biochemistry, and Prof. K. H. Sit of the Dept. of Anatomy of the National University of Singapore. HepG2 cells were purchased from American Type Culture Collection, Manassas, VA. DEPMPO was from Oxis Health Products Inc., Portland, OR.

Cell Culture—Neuro-2a, PC12, HepG2, and human fetal liver cells were cultured in Dulbecco's modified Eagle's medium containing 100 units/ml each of penicillin G and streptomycin and 0.25 µg of amphotericin B supplemented with 10% fetal bovine serum in a humidified incubator with 5% CO2 at 37 °C.

Measurement of ROS and ATP in Neuronal and Hepatic Cells—ROS were assayed using DCFDA (31). The measurement of intracellular ATP was based on the luciferin-luciferase reaction. Both methods have been reported in our previous study (30).

Preparation of Intact Rat Brain Mitochondria—The procedure as reported (32) was essentially a combination of differential centrifugation (33) and purification by Percoll density gradient centrifugation (34). Routinely, two adult Wistar rats of about 150 g each were killed by cervical dislocation, followed by decapitation. The forebrains (weighing ~2.5 g) were immediately excised and cut into small pieces with scissors in ice-cold isolation buffer containing 20 mM Tris-HCl, 250 mM sucrose, 40 mM KCl, 2 mM EGTA, and bovine serum albumin at 1 mg/ml, pH 7.2, using a hand-held glass homogenizer with six upward and downward strokes. The crude homogenate was centrifuged at 2,000 x g for 3 min to remove cell debris and nuclei. The supernatant was re-centrifuged at 12,000 x g for 11 min. The mitochondrial pellet was re-suspended in 3 ml of 15% (v/v) Percoll. This suspension was overlain on two preformed layers consisting of 3.5 ml each of 23 and 40% (v/v) Percoll. The gradient was established by centrifugation at 30,700 x g for 5 min. The distinct creamy layer near the interface of 15 and 23% Percoll (v/v) was collected and re-suspended in 0.5 ml of the same ice-cold isolation buffer. Isolation buffer (0.5 ml) was added to the pellet obtained after two centrifugations at 12,000 x g for 11 min to give a protein concentration of 10–15 mg/ml as measured by the Bradford method (35).

Staining of Isolated Mitochondria—MitoTracker Red (or CMX-rosamine) was used to stain isolated mitochondria (36). A stock solution of 0.5 mM CMX-rosamine was dissolved in Me2SO and stored at –20 °C. This was diluted with distilled water to 10 µM for use. Isolated brain mitochondria containing about 200 µg protein were washed in 1 ml PBS and centrifuged for 5 min at 10,000 x g. The pellet was re-suspended in 1 ml of PBS. To this was added 3 µl of the diluted CMX-rosamine solution and incubated at 4 °C for 15 min followed by centrifugation as before. The mitochondrial pellet was re-suspended in 50 µl PBS and transferred onto a glass slide for visualization by confocal microscopy (Becton Dickinson, Franklin Lakes, NJ, USA). Excitation was by an argon laser beam at 543 nm at 1 mW and the fluorescence emission was detected through a 560 nm long-pass filter.

JC-1 is a cationic carbocyanine dye whose uptake into mitochondria is driven by the transmembrane potential (37). A stock solution of 2 mM JC-1 prepared in Me2SO was diluted to 20 µM for use. Into an Eppendorf tube were added 10 µl of rat brain mitochondria, 100 µl of respiratory buffer (0.1 M KCl, 12.5 mM K2PO4, 10 mM Tris-HCl, pH 7.6), and 2 µl of 20 µM JC-1 solution. An aliquot of the sample was then transferred immediately onto a glass slide for visualization by confocal microscopy with excitation by an argon laser beam at 485 nm. The green fluorescence emission was detected through a long-pass filter (BA505–525), and the emission of the red aggregates through another long-pass filter (BA560). Subsequently, 5 µl of a 1.88 M succinate solution was introduced to energize the mitochondria, and one drop of the reaction mixture was again examined as before for both JC-1 monomers and aggregates.

Measurement of Mitochondrial Membrane Potential—The mitochondrial membrane potential (MMP) was measured on a PerkinElmer Life Sciences LS50B luminescence spectrometer (Buckinghamshire, UK). The cuvette was maintained at 37 °C by an external water bath. 2 ml of respiratory buffer was introduced into a cuvette followed by 10 µl of isolated brain mitochondrial extract (containing between 0.1 and 0.15 mg of protein) and 10 µlof20 µM JC-1. The contents in the cuvette were continuously stirred with a magnet. Two modes of monitoring were selected, namely the scan program for wavelength-dependent kinetics and the dual-wavelength program for time-dependent kinetics. Generally, the former was suitable for capturing rapid changes within a few minutes by continuously scanning the green fluorescence of the monomers and the red fluorescence of the J-aggregates from 520 to 620 nm. Subsequent changes were recorded at 1-min intervals. On the other hand, the dual-wavelength program provides a continuous record for longer times (30 min or more), and the excitation/emission wavelengths were 485 nm/535 nm for the green monomers and 485 nm/595 nm for the red aggregates of JC-1. Following an initial period of incubation, glutamate, malate, or succinate was introduced to energize the mitochondria followed by the addition of sulfite or CCCP. The concentrations of substrates, sulfite, and CCCP are as stated in individual sets of experiments.

ATP Biosynthesis by Isolated Brain Mitochondria—The assay procedure reported previously (30) was employed with one modification: P1,P5-Di(adenosine-5')pentaphosphate (Ap5A), an inhibitor of adenylate kinase (38), was included to account for its contribution to ATP biosynthesis. To 1.5 ml of respiratory buffer was added 125 µM ADP and 25 µM Ap5A. Each of the following substrates, namely glutamate, malate, {alpha}-ketoglutarate, and succinate was added at 5 mM concentration in the presence or absence of 100 µM sulfite. To mimic a more physiological cellular environment where a mixture of respiratory substrates is normally present, the four respiratory substrates were also introduced together at concentrations that bear a ratio similar to their concentrations reported in the whole rat/mouse brain. The respective values for glutamate, {alpha}-ketoglutarate, malate, and succinate in the brain expressed in micromoles/g fresh wt tissue were 10.4, 0.19, 0.26, and 0.69 (39); these substrates were therefore added at 5, 0.09, 0.13, and 0.33 mM concentrations. In the reaction mixture containing succinate, 25 µM rotenone was included to inhibit complex I to prevent the accumulation of oxaloacetate (from the oxidation of malate), which is a known competitive inhibitor of succinate dehydrogenase. The reaction was carried out in four replicates with 10 µl of fresh isolated mitochondria containing about 0.1 mg of protein at room temperature (26 °C) for 15 min. It was stopped by boiling for 3 min, and the reaction mixtures were assayed immediately or after storage at –80 °C for ATP.

Fresh or thawed samples were centrifuged at 15,000 rpm for 10 min in a Microfuge (Beckman) at 4 °C. The supernatant was removed, and suitable dilutions of 10 or 50 times were made depending on the concentration of ATP analyzed so that results could be extrapolated from a standard curve of 5–100 pmol of ATP, a range that produced a linear response. 25-µl aliquots of the diluted supernatant were used for ATP determination following the procedure provided in the Sigma FL-ASC Bioluminescent Somatic cell assay kit. The FL-AAM (ATP assay mix) was diluted 250 times in FL-AAB solution (ATP assay mix dilution buffer), and 100 µl of this diluted FL-AAM was then added within 1 min. The chemiluminescence emitted by the luciferin-luciferase reaction was measured using the single photon mode of counting in a liquid scintillation counter (Beckman LS6500) (30, 40).

Measurement of Activities of GDH and MDH in Brain Mitochondrial Extract—NADH formed by GDH and MDH was measured fluorometrically on a PerkinElmer Life Sciences LS50B luminescence spectrometer with excitation/emission of 352/464 nm. The mitochondrial extract was from preparations of intact mitochondria that had been kept frozen at –80 °C. It was re-suspended in a sucrose-containing medium: 0.25 M mannitol, 7.5 mM sucrose, 5 mM HEPES, 2.5 mM potassium phosphate, 5 mM MgSO4, and 1 mM EGTA, pH 7.4. Aliquots of mitochondrial extract containing about 0.3 mg of protein were used in a final volume of 2 ml of sucrose-containing medium for measuring GDH and 0.003 mg of protein for MDH. Kinetic data were obtained at 37 °C with 5 mM glutamate and 1 mM NAD+ in the presence or absence of 25–100 µM sulfite. Sequential addition of glutamate and NAD+ was made with sulfite introduced either at the beginning or the end of the sequence. Appropriate controls were carried out to ensure that the reagents: glutamate, malate, NAD+, and sulfite did not interfere with the measurement of the fluorescence of NADH.

Effect of Sulfite on GDH—The reaction mixture contained 7.5 units of a commercial preparation of GDH (Sigma, Type II from bovine liver with specific activity of 40 units/mg of protein), 5 mM glutamate, and 1 mM NAD+ in a final volume of 2 ml of sucrose-containing medium at pH 7.4 as described above. NADH produced in the presence or absence of 25–100 µM sulfite was monitored at 340 nm on a Beckman spectrophotometer (Model DU640B) for 10 min.

Kinetics of the GDH Reaction—0.5 unit of pure GDH was employed in a total volume of 2 ml of sucrose-containing medium as described above. NADH produced was measured continuously for 5 min at 37 °C using a PerkinElmer Life Sciences LS55 luminescence spectrometer. Two sets of experiments were performed in the absence and presence of 25, 50, and 100 µM sulfite using (a) 0.31–7.5 mM glutamate and 1 mM NAD+ and (b) 0.063–1.5 mM NAD+ and 5 mM glutamate. The Km and Vmax values were generated using the GraphPad software.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Sulfite-induced Changes in Neuronal and Hepatic Cells— The production of ROS, measured by the oxidation of DCFDA, was evident following exposure of Neuro-2a and PC12 cells to micromolar concentrations of sulfite for 30 min (Fig. 1). Intracellular ATP measured 3 h later was significantly depleted by 100 µM sulfite, with about 50–60% decrease in both cell lines (Fig. 2). However, cell viability determined by the MTT assay was not compromised (data not shown). Similar observations were made in HepG2 and human fetal liver cells with respect to ROS and ATP, although the decrease in ATP was less pronounced in hepatic compared with neuronal cells.



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  		FIG. 1.
Increase of reactive oxygen species (ROS) by sulfite. ROS was assayed by the increase in fluorescence of DCF following exposure of confluent cells (30,000 cells in a 96-well plate) in the presence and absence of freshly prepared 5–500 µM sulfite for 30 min. Details of the assay have been reported (30). The following cell lines were used: Neuro-2a, PC12, HepG2, and human fetal liver cells. Readings were taken using a microplate reader, and values are expressed as means ± S.D. for n = 4. p values indicate statistically significant increases of ROS as follows: *, p ≤ 0.05; **, p ≤ 0.01; and #, p ≤ 0.005.

 



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  		FIG. 2.
Sulfite-induced depletion of cellular ATP. Confluent cells (300,000 per well in 24 well plates) were exposed to freshly prepared sulfite solutions of 5 to 500 µM for 3 h. A decrease in intracellular ATP measured by the luciferin-luciferase assay as reported (30) was observed in Neuro-2a, PC12, HepG2 and human fetal liver cells. Values represent means ± S.D. for n = 4, and p values indicate statistically significant decreases in cellular ATP as follows: *p < 0.05, ** p < 0.01, # p < 0.005.

 
Quality of Mitochondrial Preparations—Three different procedures showed that mitochondria prepared from rat brain by differential centrifugation and purification by Percoll density gradient contained intact mitochondria suitable for studies of bioenergetics. Uptake of MitoTracker Red (or CMX-rosamine) into intact mitochondria was demonstrated by the intense red fluorescence of the probe (Fig. 3). With JC-1, another specific mitochondrial probe, its uptake as green fluorescent monomers was apparent under basal conditions without added exogenous substrate (Fig. 4a), with less intense red fluorescence of the J aggregates (Fig. 4b). Upon subsequent addition of succinate whose oxidation energized the mitochondria, uptake of JC-1 monomers (Fig. 4c) with an increase in MMP was reflected by the increased intensity of red fluorescence of the J aggregates (Fig. 4d). Isolated rat brain mitochondria also exhibited ADP-independent (state 4) and ADP-dependent (state 3) respiration on addition of succinate (data not shown).



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  		FIG. 3.
Confocal microscopy of isolated rat brain mitochondria. Isolated rat brain mitochondria (200 µg of protein/ml) were exposed to 0.03 µM CMX-rosamine at 4 °C for 15 min. The re-suspended mitochondrial pellet in 50 µl of PBS was transferred to a glass slide for confocal microscopic examination. Uptake of CMX-rosamine by intact mitochondria was shown by the intense fluorescence of MitoTracker Red.

 



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  		FIG. 4.
Energization of intact mitochondria by succinate. Isolated rat brain mitochondria (containing 100 µg of protein) in 100 µl of respiratory buffer was incubated with 0.36 µM JC-1 solution at 4 °C for 15 min. Uptake of JC-1 in the absence of exogenous respiratory substrate showed the green fluorescence of JC-1 monomers (a) with some red fluorescence of the J-aggregates (b). Mitochondria energized by exogenous 80 mM succinate also showed uptake of green monomers (c) and formation of more J-aggregates of higher fluorescence intensity (d).

 
Inhibition of ATP Biosynthesis from Oxidation of Glutamate by Sulfite—Little or no ATP was biosynthesized in the absence of exogenous substrate and ADP (Table I). When fortified with 125 µM ADP, mitochondria produced ATP, and this ADP-dependent endogenous respiration was inhibited significantly by Ap5A, a specific inhibitor of adenylate kinase (33). Thus without addition of exogenous respiratory substrate, much of the resulting ATP appeared to be via adenylate kinase. Addition of 5 mM of respiratory substrates such as succinate, glutamate, malate, or {alpha}-ketoglutarate fueled electron transport to produce more ATP. This respiratory substrate-driven biosynthesis of ATP was not significantly affected by Ap5A (Table I). A possible explanation is that in the presence of a highly energized electron transport chain driven by the added respiratory substrates, most of the ADP added was efficiently harnessed by F0F1-ATPase. However, Ap5A was added in subsequent measurements unless otherwise stated. The biosynthesis of ATP from glutamate and ADP, in the presence of Ap5A, was significantly inhibited by 10–100 µM sulfite in a dose-dependent manner (Table II), with almost 50% inhibition at 100 µM sulfite; inhibition was not observed with malate, {alpha}-ketoglutarate, or succinate plus rotenone (Table II). In a separate experiment where glutamate was introduced together with the other three respiratory substrates at concentrations that reflect their relative concentrations in the normal whole rat/mouse brain (39), the inhibition by sulfite on ATP biosynthesis was not evident (Table III). The decrease in ATP biosynthesis from glutamate oxidation by 5–100 µM sulfite was not affected by DEPMPO, a spin-trapping agent of free sulfite radicals (41), as shown in Fig. 5.


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  		TABLE I
Biosynthesis of ATP in isolated rat brain mitochondria Intact rat brain mitochondria (0.1 mg of protein/ml) were incubated in the absence and presence of respiratory substrates, 125 µM ADP and 25 µM Ap5A (denoted by footnote a), for 15 min at 26 °C. ATP was determined by the luciferin-luciferase assay described under "Experimental Procedures." Values are expressed in means ± S.D. (n = 4).

 


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  		TABLE II
Effect of sulfite on mitochondrial GDH activity Intact rat brain mitochondria (0.1 mg protein/ml) were incubated with respiratory substrates in the absence or presence of 10 to 100 µM sulfite for 15 min at 26 °C, followed by boiling for 3 min. With succinate, 25 µM rotenone was also included, and this is denoted by footnote a. ATP biosynthesized from 125 µM ADP in the presence 25 µM Ap5A (to inhibit adenylate kinase) was determined by the luciferin-luciferase assay described under "Experimental Procedures." Values represent means ± S.D. (n = 4). The p values were determined from Student's t test.

 


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  		TABLE III
Generation of ATP from oxidation of respiratory substrates In a separate experiment carried out under conditions similar to those described in Table II, Ap5A and rotenone were omitted to mimic a more physiological intracellular environment. The respiratory substrates were added singly (at 5 mM each) or together at concentrations given in parentheses: glutamate (5 mM), succinate (0.33 mM), malate (0.13 mM), and {alpha}-ketoglutarate (0.09 mM), which reflect their relative concentrations in normal whole rat/mouse brain (39). Values are means ± S.D. (n = 4).

 



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  		FIG. 5.
Effect of DEPMPO on mitochondrial function. Isolated rat brain mitochondria (100 µg of protein/ml) were incubated with 5 mM glutamate and 125 µM ADP in the absence ({blacksquare}) and presence ({diamondsuit}) of 100 µM DEPMPO. The dose-dependent decrease in ATP biosynthesis produced by 10–100 µM sulfite was not significantly different in the two sets of data, suggesting that sulfite radicals are probably not implicated in sulfite-induced mitochondrial impairment.

 
Loss of MMP by Sulfite—The uptake of JC-1 as green monomers (535 nm) by intact rat brain mitochondria under basal (without added respiratory substrate) conditions was apparent, and formation of J-aggregates (595 nm) also occurred (Fig. 6A). The addition of 5 mM glutamate energized the mitochondria with an immediate increase in the J-aggregate peak and a reciprocal decrease in the monomer peak (Fig. 6A). Successive traces monitored at 1-min intervals showed a progressive increase of the J-aggregates (595 nm) with a corresponding decrease of the green monomers (535 nm). When an apparent equilibrium was reached with no further visible changes, 100 µM sulfite was added. A collapse of the MMP was evident, and the decrease of the J-aggregates continued for several more minutes (Fig. 6B). Likewise, isolated mitochondria were energized by 5 mM malate (Fig. 6C) and 5 mM succinate (Fig. 6E), but sulfite failed to dissipate the established MMP in both instances (Fig. 6, D and F), suggesting that sulfite is unlikely to act directly on the inner mitochondrial membrane. In a separate experiment using 8.33 mM succinate, sulfite again showed no action on the pre-established MMP but de-energization of mitochondria was observed by a subsequent addition of 4 µM CCCP, a classic uncoupler. The decrease in MMP started almost instantaneously, and MMP continued to fall as shown in subsequent 1-min scans (Fig. 6G).



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  		FIG. 6.
Membrane potential in rat brain mitochondria. Isolated rat brain mitochondria (50 µg of protein/ml) were incubated with 0.1 µM JC-1 in 2 ml of respiratory buffer at 37 °C in a cuvette with a magnetic stirrer. The green fluorescence of the monomers and the red fluorescence of the J-aggregates were scanned continuously from 520 to 620 nm in a PerkinElmer Life Sciences LS50B luminescence spectrometer. Subsequent scans were recorded at 1-min intervals following addition of 5 mM glutamate. A progressive increase in fluorescence at 595 nm was observed (A). The MMP so established was dissipated upon the addition of 100 µM sulfite (B), and the collapse of the MMP at this wavelength was observed progressively in subsequent scans. Likewise, energization of mitochondria by 5 mM malate (C) and 5 mM succinate (E) was apparent, but the MMP so established was not affected by sulfite (D and F). In a separate experiment (g), MMP generated by succinate was again not affected by sulfite, but it was dissipated by 4 µM CCCP, a classic uncoupler.

 
Similar to the observations above but recorded by the time-dependent dual-wavelength scan, isolated rat brain mitochondria took up the JC-1 probe as green fluorescent monomers (upper trace) and formed red fluorescent J-aggregates (lower trace) progressively for about 10 min (Fig. 7A). The reciprocal relationship between the two species was consistently shown in all traces. Energization of the mitochondria, reflected by an increase in the J-aggregate formation, was evident upon addition of 5 mM each of glutamate (Fig. 7A), malate (Fig. 7C), and succinate (Fig. 7E). Preincubation of mitochondria with sulfite abolished completely the energizing action of glutamate (Fig. 7B) while increasing the MMP shown in Fig. 7A, but partially that with malate as substrate (Fig. 7D), and it had little effect when succinate was the respiratory substrate (Fig. 7F).



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  		FIG. 7.
Generation of MMP by respiratory substrates. The experimental conditions were as described in Fig. 6, but the recordings were carried out using the dual-wavelength program with excitation/emission of 485 nm/535 nm for the green monomers (upper trace) and 485 nm/595 nm for the red J-aggregates (lower trace). Isolated rat brain mitochondria (50 µg/ml) were energized by 5 mM each of glutamate (A), malate (C), and succinate (E). Preincubation of mitochondria with sulfite abolished the ability of mitochondria to generate an increase in MMP by glutamate (B), in contrast to that observed in a. The inhibitory effect of sulfite was partial with malate (D) and not apparent with succinate (F).

 
Effect of Sulfite on NAD-linked Oxidation of Glutamate and Malate—The oxidation of glutamate and malate by their respective NAD-linked dehydrogenases in rat brain mitochondrial extract was monitored by measuring the fluorescence of NADH. In the presence of 1 mM NAD+ and 5 mM glutamate or malate, NADH was generated, and this was inhibited completely in the GDH reaction by 100 µM sulfite (Fig. 8A) but partially with malate as substrate (Fig. 8D). A dose-dependent inhibition was apparent from 25–100 µM sulfite in the GDH reaction (Fig. 8B). Preincubation of the mitochondrial extract with sulfite abolished the GDH reaction fully (Fig. 8C) but inhibited the MDH activity partially (Fig. 8E). Controls with NAD+, glutamate, and sulfite added to a solution of NADH showed that they did not interfere in the measurement of fluorescence of NADH (data not shown).



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  		FIG. 8.
Inhibition of activities of GDH and MDH by sulfite. Aliquots of rat brain mitochondrial extract containing 0.3 mg of protein (for GDH assay) and 0.003 mg of protein (for MDH assay) were added to a final volume of 2 ml of sucrose-containing medium. A 100 times diluted enzyme extract was necessary, because the MDH activity was about 70 times higher than that of GDH.2 The oxidation of 5 mM each of glutamate (A–C) or malate (D and E) was recorded when 1 mM NAD+ was introduced. NADH formed was measured fluorometrically on a PerkinElmer Life Sciences LS50B luminescence spectrometer with excitation/emission of 352 nm/464 nm. Complete inhibition was observed with 100 µM sulfite (A and B) with progressive inhibition by sulfite at 25 and 50 µM (B). Preincubation of mitochondria with 100 µM sulfite abolished the GDH activity (C). Partial inhibition of MDH was observed with sulfite added after or prior to the progress of the reaction (D and E, respectively).

 
Inhibition of GDH by Sulfite—The activity of pure GDH was inhibited by sulfite in a dose-dependent manner from 25–100 µM (Fig. 9); this was measured spectrophotometrically. Kinetic data obtained by the fluorometric analysis of NADH are shown in Table IV with Km and Vmax values in the presence and absence of sulfite.



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  		FIG. 9.
Dose-dependent inhibition of GDH by sulfite. GDH activity in a commercial preparation was measured spectrophotometrically. 7.5 units of GDH (Sigma Type II from bovine liver of specific activity of 40 units/mg of protein) were employed in an assay mixture containing 5 mM glutamate and 1 mM NAD. A dose-dependent inhibition of GDH activity by 25–100 µM sulfite was observed.

 


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  		TABLE IV
Kinetics of sulfite action on glutamate dehydrogenase 0.5 unit of pure GDH was incubated in the absence and presence of 25, 50, and 100 µM sulfite at 37 oC in a final volume of 2 ml of sucrose-containing buffer. NADH produced was monitored at excitation/emission 352 nm/464 nm in a PerkinElmer Life Sciences LS55 luminescence spectrometer for 5 min. From the kinetic data of Vmax/Km generated from GraphPad software, a mixed type of inhibition was observed when measured with 0.31–7.5 mM glutamate and 1.5 mM NAD+, whereas uncompetitive inhibition was apparent with 0.063–1.5 mM NAD+ and 5 mM glutamate.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Clinical studies have alluded to an energy deficit in cases of human sulfite oxidase (SO) deficiency (42). In other cases of molybdenum cofactor deficiency, which would compromise SO activity, data from magnetic resonance imaging also suggested hypoxic-ischemic encephalopathy (43). It was proposed that neuronal injury was caused by mitochondrial damage by sulfite, which disrupts mitochondrial membrane integrity leading to decreased ATP or energy failure (44). From these noninvasive studies, it was concluded that the underlying cause was a dysfunction of brain mitochondria. Our studies provide evidence that sulfite induced a significant and dose-dependent decrease in ATP synthesis in rat brain mitochondria respiring on glutamate, a major complex I substrate in brain (Table II). This sulfite-mediated depletion of ATP was also shown in Neuro-2a, PC12, HepG2, and human fetal liver cells (Fig. 2). Similar observations have been reported with rat kidney mitochondria and MDCKII and OK cells (30). However, when other respiratory substrates, namely {alpha}-ketoglutarate, malate, and succinate, were added at concentrations that mimic their relative concentrations in the whole rat/mouse brain, the inhibitory effect of sulfite on ATP production was not shown (Table III). It would appear that, under these experimental conditions, ATP generated from these other substrates could compensate for the inhibition on GDH. However, we do not know the relative levels of these various metabolites in brain mitochondria or in human brain generally. If the human brain is more dependent on glutamate than the other respiratory substrates (which is likely under sulfite-mediated stress situations to be discussed later), the inhibitory effect of sulfite may manifest in compromised neuroenergetics. It must also be emphasized that accumulation of sulfite is more likely to occur in human brain than in rat brain whose SO activity was reported to be 10 times higher (6).

Besides a decrease in ATP, an oxidative stress state was apparent following exposure of Neuro-2a, PC12, HepG2, and human fetal liver cells to sulfite (Fig. 1). Cells in culture are possibly under a state of "culture shock" (45), and this was observed by an increase in the fluorescence of DCF in the controls without sulfite. However, sulfite further increased the ROS production (Fig. 1). Although a decrease in ATP and an increase in ROS following exposure to exogenous sulfite were observed in both neuronal and non-neuronal cells, under normal physiological in vivo situations, accumulation of sulfite is more likely to occur in neuronal cells compared with hepatic and renal cells whose high SO activity (19, 20) would facilitate its oxidation to sulfate more efficiently. This could possibly contribute to the more pronounced decrease in ATP in neuronal compared with hepatic cells (Fig. 2). The increase in oxidative stress and decreased ATP suggested that the target of sulfite action could be the mitochondrion. Using isolated rat brain mitochondria, oxidation of glutamate with phosphorylation of ADP was found to be significantly inhibited by sulfite in a dose-dependent manner (Table II) with as much as 50% decrease in the presence of 100 µM sulfite, i.e. mitochondria are a direct target. This sulfite-mediated decrease in ATP was not affected by the presence of DEPMPO (Fig. 5), which seemed to suggest that reactive sulfite radicals are probably not involved, because DEPMPO is a spin trap able to form adducts with sulfite radicals (41). The biosynthesis of ATP from the oxidation of malate, {alpha}-ketoglutarate, or succinate was, however, not affected by sulfite. From our measurements of three parameters of mitochondrial function, namely ATP biosynthesis, MMP, and NAD-linked dehydrogenase activities, GDH was found to be vulnerable to the action of sulfite. With an inhibition of GDH (Figs. 8, A–C) and the associated loss of the MMP (Figs. 6B and 7B), ATP production from the oxidation of glutamate was decreased significantly (Tables II and III). Although MDH activity was also inhibited by sulfite, the effect was only evident when a substantially diluted (100 times) preparation of mitochondria was used. Even so, only partial inhibition was observed (Fig. 8, D and E). The biosynthesis of ATP from the oxidation of succinate or {alpha}-ketoglutarate was not compromised (Table II), suggesting that respiratory complexes I–IV are functional. It was therefore concluded that sulfite targets specifically GDH. Kinetic data on pure GDH showed a decrease in Vmax and an increase in Km for glutamate in the presence of 25–100 µM sulfite resulting in a progressive decrease in Vmax/Km (Table IV) suggestive of a mixed type of inhibition. On the other hand, when the concentrations of NAD+ were varied, uncompetitive inhibition with parallel decreases in Km for NAD+ and Vmax were obtained, and the Vmax/Km remained relatively constant (Table IV). Presumably, sulfite could bind to the NAD+·GDH binary complex or to the NAD+·GDH·glutamate ternary complex.

Under normal physiological conditions, the main source of energy for brain function is glucose (46) of which 60–80% of its oxidation takes place in glutamatergic neurons to support the glutamate/glutamine cycle (47). Glutamine is an abundant amino acid in the brain, and it can be converted to glutamate, which is present in severalfold higher concentrations than the Krebs cycle metabolites; it therefore contributes significantly to mitochondrial respiratory function. Inhibition of GDH by an accumulation of sulfite in the brain of subjects with human SO deficiency would conceivably cause an energy crisis as shown in MR imaging (42–44). With inhibition of GDH by sulfite, the supply of {alpha}-ketoglutarate would be curtailed, because in the brain the GDH reaction operates in the direction of oxidative deamination to form {alpha}-ketoglutarate (48). Other tricarboxylic acid cycle intermediates such as malate and succinate would be correspondingly decreased because of a diminished flux through the tricarboxylic acid cycle. Thus under the sulfite-mediated state, their ability to compensate for GDH inhibition in terms of ATP production (as shown in Table III) would not prevail. Besides the depletion of {alpha}-ketoglutarate, the rate-limiting enzyme of the tricarboxylic acid cycle, namely {alpha}-ketoglutarate dehydrogenase, has been reported to be inhibited in isolated synaptosomes by hydrogen peroxide (49, 50). This enzyme is generally vulnerable to oxidative stress, a phenomenon that contributes to multiple neurodegenerative diseases, including Alzheimer's disease and Parkinson's disease (51). ROS generated by sulfite in Neuro-2a and PC12 cells could act likewise. Thus the combined inhibition of GDH and possibly {alpha}-ketoglutarate dehydrogenase would lead to an accumulation of glutamate relative to the above mentioned tricarboxylic acid intermediates. The association of a high level of glutamate with brain dysfunction is conceptually attractive because glutamate itself is neuroexcitotoxic. Sulfite present in a dexamethasone preparation was shown to increase the neurotoxicity of excitotoxic agents (5). Glutamate is also the precursor of GABA, an inhibitory neurotransmitter. Interestingly, a loss of GDH activity was reported in a multiple system atrophy condition called olivopontocerebellar atropy where GDH activity was decreased to a greater extent than other mitochondrial enzymes (52), suggesting that GDH is more sensitive to insults. Our study showed that it is, however, not inhibited by hydrogen peroxide from 10–220 µM concentration.2 GDH has been found to be widely distributed in numerous areas of the human brain (53), and it is tempting to conclude that its inhibition by sulfite is responsible for the widespread abnormal ischemic patterns in MR imaging of a patient with SO deficiency (54). We propose that the energy deficiency is the result of an inhibition of GDH with decrease in {alpha}-ketoglutarate and other tricarboxylic acid cycle intermediates, an overall decrease in NADH flux through the mitochondrial electron transport chain, a compromised MMP, and a decrease in ATP biosynthesis. These parameters were demonstrated in this study. The impairment of mitochondrial function could be severe, or it could predispose the brain to damage by other insults leading to early death of afflicted infants with this condition (15–17). ATP is also required for the synthesis of 3'-phosphoadenosine-5'-phosphosulfate or "active sulfate." Thus, a decrease in sulfate and ATP would block synthesis of sulfated lipids and other sulfated macromolecules in the brain.


   FOOTNOTES
 
* This work was supported by research grants from National University of Singapore (R-183-000-033-112) and National Medical Research Council (0655/2002). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed. Tel.: 65-6874-3244; Fax: 65-6779-1453; E-mail: bchsitkp{at}nus.edu.sg.

1 The abbreviations used are: SO, sulfite oxidase; ROS, reactive oxygen species; Ap5A, P1,P5-di(adenosine-5')pentaphosphate; CCCP, carbonylcyanide-m-chlorophenylhydrazone; DEPMPO, 5-diethoxyphosphoryl-5-methyl-1-pyrroline-n-oxide; GDH, glutamate dehydrogenase; JC-1, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide; MDH, malate dehydrogenase; MDCK, Madin-Darby canine kidney cells; MMP, mitochondrial membrane potential; MTT, thiazolyl blue or 3-(4,5-dimethylthiazolyl-2)-2,5-diphenyltetrazolium bromide; OK, opossum kidney cells. Back

2 X. Zhang, A. S. Vincent, B. Halliwell, and K. P. Wong, unpublished observation. Back



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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