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Oxidative Medicine and Cellular Longevity
Volume 2013 (2013), Article ID 607610, 8 pages
Neurotoxic Effects of trans-Glutaconic Acid in Rats
1Laboratório de Erros Inatos do Metabolismo, Programa de Pós-graduação em Ciências da Saúde, Unidade Acadêmica de Ciências da Saúde, Universidade do Extremo Sul Catarinense, 88806-000 Criciúma, SC, Brazil
2Núcleo de Excelência em Neurociências Aplicadas de Santa Catarina (NENASC), Programa de Pós-Graduação em Ciências da Saúde, Unidade Acadêmica de Ciências da Saúde, Universidade do Extremo Sul Catarinense, 88806-000 Criciúma, SC, Brazil
3Departamento de Bioquímica, Instituto de Ciências Básicas da Saúde, Universidade Federal do Rio Grande do Sul, 90035-003 Porto Alegre, RS, Brazil
Received 9 January 2013; Revised 3 March 2013; Accepted 4 March 2013
Academic Editor: Emilio Luiz Streck
Copyright © 2013 Patrícia F. Schuck et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
trans-Glutaconic acid (tGA) is an unsaturated C5-dicarboxylic acid which may be found accumulated in glutaric aciduria type I, whose pathophysiology is still uncertain. In the present work it was investigated the in vitro effect of increasing tGA concentrations on neurochemical and oxidative stress parameters in rat cerebral cortex. We observed that Na+, K+-ATPase activity was reduced by tGA, but not creatine kinase, respiratory chain complex IV, and ATP synthase activities. On the other hand, tGA significantly increased lipid peroxidation (thiobarbituric acid-reactive species levels and spontaneous chemiluminescence), as well as protein oxidative damage (oxidation of sulfhydryl groups). tGA also significantly decreased nonenzymatic antioxidant defenses (TRAP and reduced glutathione levels). Our data suggest that tGA may be neurotoxic in rat brain.
Glutaryl-CoA dehydrogenase deficiency (OMIM: number 231670), also known as glutaric aciduria type I, is an autosomal recessive metabolic disorder due to a blockade in the catabolic pathway of the amino acids lysine, hydroxylysine, and tryptophan. This disease was first described by Goodman et al.  and is biochemically characterized by tissue accumulation of predominantly glutaric acid and, to a lesser degree, of 3-hydroxyglutaric and trans-glutaconic (tGA) acids [2, 3]. Clinically, affected patients present macrocephaly, progressive dystonia, and dyskinesia, symptoms that are apparent within the first year of life [4, 5]. Degeneration of caudate and putamen following encephalopathic crises and frontotemporal atrophy at birth is also commonly seen in these patients [6, 7]. Children presenting residual glutaryl-CoA dehydrogenase activity of up to 30% (low excretors) present low or undetectable excretion of glutaric acid, but the neurological signs may be found similar in these individuals, regardless of the amount of glutaric acid excreted [8–11].
The primary cause of neurological alterations in glutaric acidemia type I is still not defined. It has been demonstrated that brain tissue exposure to the main accumulating metabolites, glutaric, and 3-hydroxyglutaric acids results in excitotoxicity [12–15], oxidative stress induction [16–18], and/or disruption of energy homeostasis [12, 19–24]. Furthermore, it has been recently showed that lysine administration to a knockout mice model of glutaric aciduria type I (Gcdh−/−) results in oxidative damage and energy impairment in the brain of these animals, as well as in compromised neurodevelopmental and cognitive behavior [25–28]. On the other hand, very little is known regarding the direct toxicity of tGA. In this scenario, and it has been demonstrated that tGA is able to inhibits glutamate decarboxylase activity in rat and rabbit brains, human mitochondrial NAD(P)(+)-dependent malic enzyme, and dopamine beta-monooxygenase activities in purified preparations, as well as inducing mitochondrial permeability transition in rat liver mitochondrial preparations, provoking apoptosis in immature oligodendrocytes, and reducing cell viability of primary cultures of mice cerebral neocortical neurons [29–34].
Therefore, the aim of the present work was to investigate the in vitro effects of tGA on various neurochemical parameters in cerebral cortex of young rats. The investigated parameters were the values of Na+, K+-ATPase, creatine kinase (CK), complex IV and ATP synthase activities, carbonyl and sulfhydryl content, thiobarbituric acid-reactive substances (TBA-RS) levels, spontaneous chemiluminescence, reduced glutathione (GSH) concentrations, and total nonenzymatic antioxidant capacity of the tissue (TRAP) in cerebral cortex of young rats.
2. Material and Methods
All chemicals were purchased from Sigma (St. Louis, MO, USA). tGA was dissolved in the day of the experiments in the incubation medium used for each technique, and the solution had its pH adjusted to 7.4. The final concentrations of tGA in the medium ranged from 0.01 to 1.0 mM.
Parallel experiments were always carried out with negative controls (blanks) in the presence or absence of tGA and also with or without cortical supernatants in order to detect artifacts caused by this organic acid in the assays. Therefore, any interference of tGA on the reactions used to measure the biochemical parameters would be identified.
Thirty-day-old male Wistar rats obtained from the Central Animal House of the Departamento de Bioquímica, Federal University of Rio Grande do Sul (UFRGS), were used. Rats were kept with dams until weaning at 21 days of age. The animals had free access to water and to a standard commercial chow and were maintained on a 12 : 12 h light/dark cycle in an air-conditioned constant temperature (22 1°C) colony room. The “Principles of Laboratory Animal Care” (NIH publication number 80-23, revised 1996) were followed in all experiments and the Ethics Committee for Animal Research of UFRGS, Porto Alegre, Brazil, approved the experimental protocol. All efforts were made to minimize the number of animals used and their suffering.
2.3. Tissue Preparation and Incubation for Oxidative Stress Parameters
On the day of the experiments the animals were killed by decapitation without anesthesia, and the brain was rapidly excised on a Petri dish placed on ice. The olfactory bulbs, brain stem, medulla, cerebellum, hippocampus, corpus callosum, and striatum were discarded, and the cerebral cortex was peeled away from the subcortical structures, weighed and homogenized in 10 volumes (1 : 10, w/v) of 20 mM sodium phosphate buffer, pH 7.4 containing 140 mM KCl. Homogenates were centrifuged at 750 ×g for 10 min at 4°C to discard nuclei and cell debris . The pellet was discarded, and the supernatant, a suspension of mixed and preserved organelles, including mitochondria, was separated and incubated in 20 mM sodium phosphate buffer, pH 7.4 containing 140 mM KCl at 37°C for one hour with tGA at concentrations of 0.01, 0.1, or 1 mM. Controls did not contain this metabolite in the incubation medium. Immediately after incubation, aliquots were taken to measure the values of carbonyl and sulfhydryl content, TBA-RS levels, spontaneous chemiluminescence, GSH concentrations, and TRAP.
2.4. Preparation of Synaptic Plasma Membrane from Rat Cerebral Cortex
Cerebral cortex was homogenized in 10 volumes of 0.32 mM sucrose solution containing 5.0 mM HEPES and 1.0 mM EDTA. The homogenate was preincubated at 37°C for 1 h in the absence or presence of 0.01, 0.1, or 1 mM tGA. Membranes were prepared afterwards according to the method of Jones and Matus  using a discontinuous sucrose density gradient consisting of successive layers of 0.3, 0.8, and 1.0 mM. After centrifugation at 69,000 ×g for 2 h, the fraction at the 0.8–1.0 mM sucrose interface was taken as the membrane enzyme preparation.
2.5. Tissue Preparations for Determination of Respiratory Chain Complex IV and Creatine Kinase Activities
For the determination of the activities of the respiratory chain complex IV, cerebral cortex was homogenized (1 : 20, w/v) in SETH buffer (250 mM sucrose, 2.0 mM EDTA, 10 mM Trizma base, and 50 UI·mL−1 heparin), pH 7.4. The homogenates were centrifuged at 800 ×g for 10 min, and the supernatants were kept at −70°C until being used for enzyme activity determination. For total creatine kinase activity determination, the cerebral cortex was homogenized (1 : 10 w/v) in isosmotic saline solution .
2.6. Mitochondrial Preparations
Mitochondrial fractions prepared according to Cassina and Radi  were used for the determination of ATP synthase activity, TBA-RS levels, and sulfhydryl content.
2.7. Determination of Na+, K+-ATPase Activity
Na+, K+-ATPase activity was evaluated according to Tsakiris and Deliconstantinos . Released inorganic phosphate (Pi) was measured by the method of Chan et al. . Enzyme-specific activity was expressed as nmol Pi released·min−1·mg protein−1.
2.8. Determination of Bioenergetics Parameters
The activity of respiratory chain complex IV was determined according to Rustin et al.  slightly modified, as described in details in a previous report . ATP synthase activity was measured in mitochondrial preparations from cerebral cortex, according to Rustin et al. . Complex IV and ATP synthase activities were calculated as nmol·min−1·mg protein−1.
2.9. Determination of Protein Carbonyl Formation Content
Protein carbonyl content, a marker of oxidized proteins, was measured spectrophotometrically according to Reznick and Packer . The results were calculated as nmol of carbonyls groups·mg of protein−1, using the extinction coefficient of 22,000 × 106 nmol·mL−1 for aliphatic hydrazones.
2.10. Sulfhydryl (Thiol) Group Oxidation
This assay was performed according to Aksenov and Markesbery . The protein-bound sulfhydryl content is inversely correlated to oxidative damage to proteins. Results were reported as nmol TNB·mg of protein−1.
2.11. Determination of TBA-RS Levels
TBA-RS was determined according to the method of Esterbauer and Cheeseman . A calibration curve was performed using 1,1,3,3-tetramethoxypropane, and each curve point was subjected to the same treatment as supernatants. Some experiments were performed in the absence or presence of reduced glutathione (GSH; 100 μM), melatonin (100 μM), the nitric oxide synthase inhibitor Nω-nitro-L-arginine methyl ester (L-NAME; 500 μM), trolox (5 μM), or a combination of catalase (Cat; 10 mU·mL−1) plus superoxide dismutase (SOD; 10 mU·mL−1). TBA-RS values were calculated as nmol of TBA-RS·mg protein−1.
2.12. Spontaneous Chemiluminescence
Samples were assayed for spontaneous chemiluminescence in a dark room by the method of Gonzalez-Flecha et al. . Results were calculated as cpm·mg protein−1.
2.13. Determination of TRAP
TRAP, representing the total nonenzymatic antioxidant capacity of the tissue, was determined by measuring the chemiluminescence intensity of luminol induced by 2,2′-azo-bis-(2-amidinopropane) (ABAP) according to the method of Lissi et al. . TRAP values were expressed as nmol of trolox·mg of protein−1.
2.14. GSH Levels Quantification
GSH levels were measured according to Browne and Armstrong . Some experiments were performed in the presence or absence of melatonin (100 μM), L-NAME (500 μM), trolox (5 μM), or Cat (10 mU·mL−1) plus SOD (10 mU·mL−1). Calibration curve was prepared with standard GSH (0.01–1 mM), and the concentrations were calculated as nmol·mg protein−1.
The oxidation of a commercial solution of GSH (200 μM) was also tested by exposing this solution to 1 mM tGA for one hour in a medium devoid of brain supernatants. After tGA exposition, 7.4 mM o-phthaldialdehyde was added to the vials, and the mixture was incubated at room temperature during 15 minutes.
2.15. Protein Determination
Protein was measured by the method of Lowry et al.  using bovine serum albumin as standard.
2.16. Statistical Analysis
Results are presented as mean standard deviation. Assays were performed in duplicate, and the mean was used for statistical analysis. Data was analysed using one-way analysis of variance (ANOVA) followed by the post hoc Duncan multiple range test when was significant. Differences between groups were rated significant at . All analyses were carried out in an IBM-compatible PC computer using the Statistical Package for the Social Sciences (SPSS) software.
tGA inhibits Na+, K+-ATPase activity in synaptic plasma membranes from rat cerebral cortex.
Initially, we tested the influence of tGA on Na+, K+-ATPase activity in synaptic plasma membranes prepared from cerebral cortex. Figure 1 shows that the incubation of tGA with purified synaptic membrane preparations caused a significant inhibition of Na+, K+-ATPase activity.
3.1. tGA Does Not Interfere on Important Parameters of Brain Bioenergetics
We also investigated the effect of tGA on important parameters of brain bioenergetics. It is observed in Table 1 that the presence of up to 1.0 mM tGA in the incubation medium does not disturb creatine kinase, respiratory chain complex IV, and ATP synthase activities in rat cerebral cortex.
3.2. Protein Oxidative Damage Is Induced by tGA
We then evaluated the effect of tGA on carbonyl formation and sulfhydryl oxidation in cortical homogenates in order to evaluate protein oxidative damage. We found that tGA provoked a significant decrease of sulfhydryl content at 0.1 and 1.0 mM in cortical homogenates (Figure 2(a)), although it did not affect protein carbonyl content. Furthermore, sulfhydryl content was not altered by tGA in brain mitochondrial preparations (Table 2).
3.3. tGA Induces Lipid Peroxidation
The influence of tGA on the lipid peroxidation parameters TBA-RS levels and spontaneous chemiluminescence was investigated. Figure 2(b) shows that tGA at 1 mM significantly increased TBA-RS levels in cortical homogenates. Similar results were obtained for spontaneous chemiluminescence.
We also tested the effect of the antioxidants GSH (100 μM), melatonin (100 μM), L-NAME (200 μM), trolox (5 μM), or a combination of Cat plus SOD (10 mU mL−1 each) on tGA-elicited increase of TBA-RS levels (Figure 3). These antioxidants were coincubated with 1.0 mM of the organic acid and brain cortical homogenates for 60 min, after which the levels of TBA-RS were measured. We observed that GSH and melatonin fully prevented tGA-induced increased lipid peroxidation, whereas the combination of Cat plus SOD partially prevented such effects. Trolox and L-NAME did not prevent tGA effect.
It was also observed that tGA did not affect TBA-RS levels in brain mitochondrial preparations (Table 2).
3.4. Nonenzymatic Antioxidant Defenses Are Decreased by tGA
We assessed the in vitro effect of tGA on cortical GSH levels and TRAP. We found that this organic acid significantly decreased GSH levels and TRAP at the highest tested concentration (1.0 mM) (Figure 4). Furthermore, tGA-induced effect was totally prevented by the free-radical scavengers melatonin and trolox and by the combination of Cat plus SOD, but not by L-NAME (Figure 3).
We then investigated whether tGA-induced GSH levels decrease was secondary to a direct oxidative attack. We therefore exposed a commercial solution of GSH (200 μM) to 1.0 mM tGA for 60 min in the absence of cortical supernatants. It was observed that this organic acid per se did not oxidize highly purified GSH (Table 2).
tGA, or (E)-pentene-1,5-dioic acid, is an unsaturated C5-dicarboxylic acid that is found in patients affected by glutaric aciduria type I. Its role on this organic aciduria pathogenesis is under debate, since there are patients affected by glutaric aciduria type I who do not present tGA accumulation . On the other hand, several tGA toxic effects have already been reported [29–34].
Herein we present data demonstrating that Na+, K+-ATPase activity, a crucial enzyme for maintaining ion homeostasis and membrane potential in cells , was inhibited by tGA at 0.1 and 1.0 mM concentrations. Inhibition of Na+, K+-ATPase activity has been described in various diseases, including cerebral ischemia [52, 53], neurodegenerative , and metabolic disorders [55, 56]. Furthermore, studies have indicated that Na+, K+-ATPase inhibition may result in cellular death by activating apoptotic cascades and neuronal damage probably due to amplification of potassium homeostasis impairment .
Considering that this enzyme is present at high concentrations in brain cellular membranes and consumes about 40–50% of the ATP generated , our next step was to evaluate the effects of tGA on some important bioenergetics parameters in order to identify whether ATP deficit could contribute to Na+, K+-ATPase activity inhibition caused by this organic acid. It was observed that tGA does not affect the activities of respiratory chain complex IV, creatine kinase, and ATP synthase. However, at this point, it should be mentioned that effects of tGA on glycolysis Krebs cycle enzyme activities, and the other respiratory chain complexes activities cannot be ruled out.
It was further investigated the in vitro effect of tGA on oxidative stress parameters, since Na+, K+-ATPase activity is highly dependent of critical sulfhydryl groups present in its catalytic site, rendering this enzyme activity highly susceptible to oxidative damage [59, 60]. We showed that tGA in vitro induced oxidative damage to proteins, as observed by sulfhydryl content decrease, and to lipids, since it increased TBA-RS levels and spontaneous chemiluminescence. Interestingly, carbonyl content was not altered by the presence of tGA in the incubation medium, suggesting that particularly sulfhydryl groups are prone to protein oxidative damage by this organic acid. On the other hand, light emitted in the spontaneous chemiluminescence assay mainly arises from oxidized lipids due to an increase in reactive oxygen or nitrogen species production, and TBA-RS levels reflects the amount of malondialdehyde formation, an end product of membrane fatty acid peroxidation . In this scenario, it is tempting to speculate that tGA oxidized lipids from cell and organelle membranes, which could lead to an impairment of membrane fluidity. Consequently, integral proteins such as Na+, K+-ATPase could also be affected, having their functioning impaired.
Furthermore, it was observed that GSH and melatonin fully prevented tGA-induced increased lipid peroxidation, since these compounds prevented TBA-RS increase elicited by this organic acid, whereas the combination of Cat plus SOD only partially prevented such effects. On the other hand, trolox (a hydrophilic vitamin E analogue) and L-NAME did not prevent tGA effect. It is speculated that lipid oxidation induced by tGA occurs due to an increase of reactive oxygen species.
We then assessed nonenzymatic antioxidant defenses by measuring GSH levels and TRAP and observed that both parameters were decreased by tGA in vitro. Since these parameters are suitable to evaluate the capacity of a tissue to prevent and respond to oxidative damage [35, 61, 62], it is likely that tGA impairs rat cortical antioxidant defenses. Moreover, tGA-induced effect on GSH levels was totally prevented by the free-radical scavengers melatonin and trolox and by the combination of Cat plus SOD, but not by L-NAME, suggesting that mainly peroxyl, alkoxyl, and hydroxyl radicals were involved in the reduction of GSH levels provoked by tGA. Considering that tGA was not able to oxidize a commercial GSH solution in the absence of brain supernatants in the incubation medium, it is unlikely that this organic acid is per se a direct oxidant agent, corroborating the idea that it probably provoked lipid and protein oxidative damage by increasing free radical generation.
Since the alterations elicited by tGA on TBA-RS and sulfhydryl content in whole homogenates, which contain the whole cell machinery, were not observed when this organic acid was supplemented to mitochondrial preparations, it is feasible that oxidative stress induced by tGA was probably mediated by cytosolic mechanisms. For instance, some putative mechanisms could involve xanthine oxidase, cytosolic NADPH oxidase, lysosomes, peroxisomes, and others .
Oxidative stress is an imbalance between the total tissue antioxidant (enzymatic and nonenzymatic) defenses and reactive species generation in cell leading to a deleterious cell condition . In this study, we showed evidences that tGA induces oxidative stress in cerebral cortex in vitro, since it increased oxidative damage and decreased antioxidant defenses. It should be emphasized that the brain is highly susceptible to damage induced by increased reactive species generation, since it has low cerebral antioxidant defenses as compared to other tissues . Furthermore, it may be also speculated that the inhibition of Na+, K+-ATPase activity elicited by tGA reported in the present study could be secondary to increased oxidation of critical sulfhydryl groups at the catalytic site of the enzyme.
Taken together, our results provide evidences that tGA is toxic to brain cells in vitro, by causing alterations in cell ion balance, and probably neurotransmission, as well as oxidative stress in rat cerebral cortex. We cannot at this point establish whether our data have a pathophysiological significance for glutaric aciduria type I. tGA accumulates in urine for excretion [65, 66], and excretion of this acid may become prominent, exceeding that of 3-hydroxyglutaric acid, during episodes of ketosis [4, 67]. Some studies indicated that the diagnostic relevance of tGA is limited , since tGA excretion in urine may be inconsistently found. However, it should be highlighted that even glutaric acid, known to be the major accumulating in glutaric aciduria type I, may be excreted at discrete or even normal concentrations (low excretor patients), as above mentioned.
Considering that the main signs and symptoms of affected patients are neurological , and in case the present findings are confirmed in vivo in animal experiments and also in tissues of patients accumulating tGA, it may be speculated that tGA toxicity could collaborate to the brain damage characteristic of glutaric aciduria type I affected patients.
Conflict of Interests
The authors declare that there are no possible conflict of interests.
This work was supported by Grants from CNPq, FAPERGS, and PROPESQ/UFRGS.
- S. I. Goodman, S. P. Markey, P. G. Moe, B. S. Miles, and C. C. Teng, “Glutaric aciduria; a “new” disorder of amino acid metabolism,” Biochemical Medicine, vol. 12, no. 1, pp. 12–21, 1975.
- I. Baric, L. Wagner, P. Feyh, M. Liesert, W. Buckel, and G. F. Hoffmann, “Sensitivity and speciticity of free and total glutaric acid and 3-hydroxyglutaric acid measurements by stable-isotope dilution assays for the diagnosis of glutaric aciduria type I,” Journal of Inherited Metabolic Disease, vol. 22, no. 8, pp. 867–881, 1999.
- S. I. Goodman and F. E. Frerman, “Organic acidemias due to defects in lysine oxidation: 2-ketoadipic academia and glutaric acidemia,” in The Metabolic and Molecular Bases of Inherited Disease, C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle, Eds., pp. 2195–2204, McGraw-Hill, New York, NY, USA, 8th edition, 2001.
- S. I. Goodman, M. D. Norenberg, and R. H. Shikes, “Glutaric aciduria: biochemical and morphologic considerations,” Journal of Pediatrics, vol. 90, no. 5, pp. 746–750, 1977.
- G. F. Hoffmann and J. Zschocke, “Glutaric aciduria type I: from clinical, biochemical and molecular diversity to successful therapy,” Journal of Inherited Metabolic Disease, vol. 22, no. 4, pp. 381–391, 1999.
- J. Brismar and P. T. Ozand, “CT and MR of the brain in glutaric acidemia type I: a review of 59 published cases and a report of 5 new patients,” American Journal of Neuroradiology, vol. 16, no. 4, pp. 675–683, 1995.
- P. Jafari, O. Braissant, L. Bonafe, and D. Ballhausen, “The unsolved puzzle of neuropathogenesis in glutaric aciduria type I,” Molecular Genetics and Metabolism, vol. 104, pp. 425–437, 2011.
- C. Busquets, B. Merinero, E. Christensen et al., “Glutaryl-CoA dehydrogenase deficiency in Spain: evidence of two groups of patients, genetically, and biochemically distinct,” Pediatric Research, vol. 48, no. 3, pp. 315–322, 2000.
- E. Christensen, A. Ribes, B. Merinero, and J. Zschocke, “Correlation of genotype and phenotype in glutaryl-CoA dehydrogenase deficiency,” Journal of Inherited Metabolic Disease, vol. 27, no. 6, pp. 861–868, 2004.
- R. C. Gallagher, T. M. Cowan, S. I. Goodman, and G. M. Enns, “Glutaryl-CoA dehydrogenase deficiency and newborn screening: retrospective analysis of a low excretor provides further evidence that some cases may be missed,” Molecular Genetics and Metabolism, vol. 86, no. 3, pp. 417–420, 2005.
- S. Kölker, E. Christensen, J. V. Leonard et al., “Diagnosis and management of glutaric aciduria type I—revised recommendations,” Journal of Inherited Metabolic Disease, vol. 34, no. 3, pp. 677–694, 2011.
- S. Kölker, D. M. Koeller, S. Sauer et al., “Excitotoxicity and bioenergetics in glutaryl-CoA dehydrogenase deficiency,” Journal of Inherited Metabolic Disease, vol. 27, no. 6, pp. 805–812, 2004.
- M. Wajner, S. Kölker, D. O. Souza, G. F. Hoffmann, and C. F. de Mello, “Modulation of glutamatergic and GABAergic neurotransmission in glutaryl-CoA dehydrogenase deficiency,” Journal of Inherited Metabolic Disease, vol. 27, no. 6, pp. 825–828, 2004.
- R. B. Rosa, K. B. Dalcin, A. L. Schmidt et al., “Evidence that glutaric acid reduces glutamate uptake by cerebral cortex of infant rats,” Life Sciences, vol. 81, no. 25-26, pp. 1668–1676, 2007.
- D. V. Magni, A. F. Furian, M. S. Oliveira et al., “Kinetic characterization of l-[3H]glutamate uptake inhibition and increase oxidative damage induced by glutaric acid in striatal synaptosomes of rats,” International Journal of Developmental Neuroscience, vol. 27, no. 1, pp. 65–72, 2009.
- F. de Oliveira Marques, M. E. K. Hagen, C. D. Pederzolli et al., “Glutaric acid induces oxidative stress in brain of young rats,” Brain Research, vol. 964, no. 1, pp. 153–158, 2003.
- A. Latini, K. Scussiato, G. Leipnitz, C. S. Dutra-Filho, and M. Wajner, “Promotion of oxidative stress by 3-hydroxyglutaric acid in rat striatum,” Journal of Inherited Metabolic Disease, vol. 28, no. 1, pp. 57–67, 2005.
- A. Latini, G. C. Ferreira, K. Scussiato et al., “Induction of oxidative stress by chronic and acute glutaric acid administration to rats,” Cellular and Molecular Neurobiology, vol. 27, no. 4, pp. 423–438, 2007.
- C. G. Silva, A. R. Silva, C. Ruschel et al., “Inhibition of energy production in vitro by glutaric acid in cerebral cortex of young rats,” Metabolic Brain Disease, vol. 15, no. 2, pp. 123–131, 2000.
- A. M. Das, T. Lücke, and K. Ullrich, “Glutaric aciduria I: creatine supplementation restores creatinephosphate levels in mixed cortex cells from rat incubated with 3-hydroxyglutarate,” Molecular Genetics and Metabolism, vol. 78, no. 2, pp. 108–111, 2003.
- C. da Ferreira G, C. M. Viegas, P. F. Schuck et al., “Glutaric acid moderately compromises energy metabolism in rat brain,” International Journal of Developmental Neuroscience, vol. 23, pp. 687–693, 2005.
- G. C. Ferreira, A. Tonin, P. F. Schuck et al., “Evidence for a synergistic action of glutaric and 3-hydroxyglutaric acids disturbing rat brain energy metabolism,” International Journal of Developmental Neuroscience, vol. 25, pp. 391–398, 2007.
- C. Ferreira Gda, P. F. Schuck, C. M. Viegas et al., “Energy metabolism is compromised in skeletal muscle of rats chronically-treated with glutaric acid,” Metabolic Brain Disease, vol. 22, pp. 1111–1123, 2007.
- S. W. Sauer, “Biochemistry and bioenergetics of glutaryl-CoA dehydrogenase deficiency,” Journal of Inherited Metabolic Disease, vol. 30, no. 5, pp. 673–680, 2007.
- A. U. Amaral, C. Cecatto, B. Seminotti et al., “Marked reduction of Na+, K+-ATPase and creatine kinase activities induced by acute lysine administration in glutaryl-CoA dehydrogenase deficient mice,” Molecular Genetics and Metabolism, vol. 107, no. 1-2, pp. 81–86, 2012.
- E. N. Busanello, L. Pettenuzzo, P. H. Botton et al., “Neurodevelopmental and cognitive behavior of glutaryl-CoA dehydrogenase deficient knockout mice,” Life Sciences, vol. 92, no. 2, pp. 137–142, 2013.
- B. Seminotti, A. U. Amaral, M. S. da Rosa et al., “Disruption of brain redox homeostasis in glutaryl-CoA dehydrogenase deficient mice treated with high dietary lysine supplementation,” Molecular Genetics and Metabolism, vol. 108, pp. 30–39, 2013.
- B. Seminotti, M. S. da Rosa, C. G. Fernandes et al., “Induction of oxidative stress in brain of glutaryl-CoA dehydrogenase deficient mice by acute lysine administration,” Molecular Genetics and Metabolism, vol. 106, pp. 31–38, 2012.
- O. Stokke, S. I. Goodman, and P. G. Moe, “Inhibition of brain glutamate decarboxylase by glutarate, glutaconate, and β hydroxyglutarate: explanation of the symptoms in glutaric aciduria?” Clinica Chimica Acta, vol. 66, no. 3, pp. 411–415, 1976.
- C. M. Palmeira, M. I. Rana, C. B. Frederick, and K. B. Wallace, “Induction of the mitochondrial permeability transition in vitro by short-chain carboxylic acids,” Biochemical and Biophysical Research Communications, vol. 272, no. 2, pp. 431–435, 2000.
- D. S. Wimalasena, S. P. Jayatillake, D. C. Haines, and K. Wimalasena, “Plausible molecular mechanism for activation by fumarate and electron transfer of the dopamine β-mono-oxygenase reaction,” Biochemical Journal, vol. 367, no. 1, pp. 77–85, 2002.
- T. M. Lund, E. Christensen, A. S. Kristensen, A. Schousboe, and A. M. Lund, “On the neurotoxicity of glutaric, 3-hydroxyglutaric, and trans-glutaconic acids in glutaric acidemia type 1,” Journal of Neuroscience Research, vol. 77, no. 1, pp. 143–147, 2004.
- B. Gerstner, A. Gratopp, M. Marcinkowski, M. Sifringer, M. Obladen, and C. Bührer, “Glutaric acid and its metabolites cause apoptosis in immature oligodendrocytes: a novel mechanism of white matter degeneration in glutaryl-CoA dehydrogenase deficiency,” Pediatric Research, vol. 57, no. 6, pp. 771–776, 2005.
- K. L. Su, K. Y. Chang, and H. C. Hung, “Effects of structural analogues of the substrate and allosteric regulator of the human mitochondrial NAD(P)+-dependent malic enzyme,” Bioorganic and Medicinal Chemistry, vol. 17, no. 15, pp. 5414–5419, 2009.
- P. Evelson, M. Travacio, M. Repetto, J. Escobar, S. Llesuy, and E. A. Lissi, “Evaluation of total reactive antioxidant potential (TRAP) of tissue homogenates and their cytosols,” Archives of Biochemistry and Biophysics, vol. 388, no. 2, pp. 261–266, 2001.
- D. H. Jones and A. I. Matus, “Isolation of synaptic plasma membrane from brain by combined flotation sedimentation density gradient centrifugation,” Biochimica et Biophysica Acta, vol. 356, no. 3, pp. 276–287, 1974.
- P. F. Schuck, G. Leipnitz, C. A. J. Ribeiro et al., “Inhibition of creatine kinase activity in vitro by ethylmalonic acid in cerebral cortex of young rats,” Neurochemical Research, vol. 27, no. 12, pp. 1633–1639, 2002.
- A. Cassina and R. Radi, “Differential inhibitory action of nitric oxide and peroxynitrite on mitochondrial electron transport,” Archives of Biochemistry and Biophysics, vol. 328, no. 2, pp. 309–316, 1996.
- S. Tsakiris and G. Deliconstantinos, “Influence of phosphatidylserine on (Na+ K+)-stimulated ATPase and acetylcholinesterase activities of dog brain synaptosomal plasma membranes,” Biochemical Journal, vol. 220, no. 1, pp. 301–307, 1984.
- K. M. Chan, D. Delfert, and K. D. Junger, “A direct colorimetric assay for Ca2+-stimulated ATPase activity,” Analytical Biochemistry, vol. 157, no. 2, pp. 375–380, 1986.
- P. Rustin, D. Chretien, T. Bourgeron et al., “Biochemical and molecular investigations in respiratory chain deficiencies,” Clinica Chimica Acta, vol. 228, no. 1, pp. 35–51, 1994.
- C. G. da Silva, C. A. J. Ribeiro, G. Leipnitz et al., “Inhibition of cytochrome c oxidase activity in rat cerebral cortex and human skeletal muscle by D-2-hydroxyglutaric acid in vitro,” Biochimica et Biophysica Acta, vol. 1586, no. 1, pp. 81–91, 2002.
- B. P. Hughes, “A method for the estimation of serum creatine kinase and its use in comparing creatine kinase and aldolase activity in normal and pathological sera,” Clinica Chimica Acta, vol. 7, no. 5, pp. 597–603, 1962.
- A. Z. Reznick and L. Packer, “Oxidative damage to proteins: spectrophotometric method for carbonyl assay,” Methods in Enzymology, vol. 233, pp. 357–363, 1994.
- M. Y. Aksenov and W. R. Markesbery, “Changes in thiol content and expression of glutathione redox system genes in the hippocampus and cerebellum in Alzheimer's disease,” Neuroscience Letters, vol. 302, no. 2-3, pp. 141–145, 2001.
- H. Esterbauer and K. H. Cheeseman, “Determination of aldehydic lipid peroxidation products: malonaldehyde and 4-hydroxynonenal,” Methods in Enzymology, vol. 186, pp. 407–421, 1990.
- B. Gonzalez Flecha, S. Llesuy, and A. Boveris, “Hydroperoxide-initiated chemiluminescence: an assay for oxidative stress in biopsies of heart, liver, and muscle,” Free Radical Biology and Medicine, vol. 10, no. 2, pp. 93–100, 1991.
- E. Lissi, C. Pascual, and M. D. Del Castillo, “Luminol luminescence induced by 2,-Azo-bis(2-amidinopropane) thermolysis,” Free Radical Research Communications, vol. 17, no. 5, pp. 299–311, 1992.
- R. W. Browne and D. Armstrong, “Reduced glutathione and glutathione disulfide,” Methods in Molecular Biology, vol. 108, pp. 347–352, 1998.
- O. H. Lowry, N. J. Rosebrough, A. L. Farr, and R. J. Randall, “Protein measurement with the Folin phenol reagent,” The Journal of Biological Chemistry, vol. 193, no. 1, pp. 265–275, 1951.
- M. Ericinska and I. A. Silver, “Silver, ions and energy in mammalian brain,” Progress in Neurobiology, vol. 16, pp. 37–71, 1994.
- A. T. S. Wyse, E. L. Streck, S. V. T. Barros, A. M. Brusque, A. I. Zugno, and M. Wajner, “Methylmalonate administration decreases Na+, K+-ATPase activity in cerebral cortex of rats,” NeuroReport, vol. 11, no. 10, pp. 2331–2334, 2000.
- A. T. de Souza Wyse, E. L. Streck, P. Worm, A. Wajner, F. Ritter, and C. A. Netto, “Preconditioning prevents the inhibition of NA+,K+-ATPase activity after brain ischemia,” Neurochemical Research, vol. 25, no. 7, pp. 971–975, 2000.
- S. P. Yu, “Na+, K+-ATPase: the new face of an old player in pathogenesis and apoptotic/hybrid cell death,” Biochemical Pharmacology, vol. 66, no. 8, pp. 1601–1609, 2003.
- M. R. Fighera, L. F. F. Royes, A. F. Furian et al., “GM1 ganglioside prevents seizures, Na+,K+-ATPase activity inhibition and oxidative stress induced by glutaric acid and pentylenetetrazole,” Neurobiology of Disease, vol. 22, no. 3, pp. 611–623, 2006.
- P. F. Schuck, D. R. De Assis, C. M. Viegas et al., “Ethylmalonic acid modulates Na+,K+-ATPase activity and mRNA levels in rat cerebral cortex,” Synapse, vol. 67, no. 3, pp. 111–117, 2013.
- X. Q. Wang, A. Y. Xiao, C. Sheline et al., “Apoptotic insults impair Na+, K+-ATPase activity as a mechanism of neuronal death mediated by concurrent ATP deficiency and oxidant stress,” Journal of Cell Science, vol. 116, no. 10, pp. 2099–2110, 2003.
- G. J. Lees, “Inhibition of sodium-potassium-ATPase: a potentially ubiquitous mechanism contributing to central nervous system neuropathology,” Brain Research Reviews, vol. 16, no. 3, pp. 283–300, 1991.
- H. Rauchová, Z. Drahota, and J. Koudelová, “The role of membrane fluidity changes and thiobarbituric acid-reactive substances production in the inhibition of cerebral cortex Na+/K+-ATPase activity,” Physiological Research, vol. 48, no. 1, pp. 73–78, 1999.
- J. F. Zhou, W. Zhou, S. M. Zhang, Y. E. R. Luo, and H. H. Chen, “Oxidative stress and free radical damage in patients with acute dipterex poisoning,” Biomedical and Environmental Sciences, vol. 17, no. 2, pp. 223–233, 2004.
- B. Halliwell and J. M. C. Gutteridge, “Detection of free radicals and others reactive species: trapping and fingerprinting,” in Free Radicals in Biology and Medicine., B. Halliwell and J. M. C. Gutteridge, Eds., pp. 351–425, Oxford University Press, Oxford, UK, 1999.
- E. Lissi, M. Salim-Hanna, C. Pascual, and M. D. Del Castillo, “Evaluation of total antioxidant potential (TRAP) and total antioxidant reactivity from luminol-enhanced chemiluminescence measurements,” Free Radical Biology and Medicine, vol. 18, no. 2, pp. 153–158, 1995.
- B. Halliwell and J. M. C. Gutteridge, “Oxygen is a toxic gas—an introduction to oxygen toxicity and reactive species,” in Free Radicals in Biology and Medicine, B. Halliwell, Gutteridge, and J. M. C, Eds., pp. 1–29, Oxford University Press, Oxford, UK, 2007.
- B. Halliwell and J. M. C. Cutteridge, “Oxygen radicals and the nervous system,” Trends in Neurosciences, vol. 8, no. 1, pp. 22–26, 1985.
- K. A. Strauss, E. G. Puffenberger, D. L. Robinson, and D. H. Morton, “Type I glutaric aciduria, part 1: natural history of 77 patients,” American Journal of Medical Genetics, vol. 121, no. 1, pp. 38–52, 2003.
- G. L. Hedlund, N. Longo, and M. Pasquali, “Glutaric acidemia type 1,” American Journal of Medical Genetics, vol. 142, no. 2, pp. 86–94, 2006.
- N. Gregersen and N. J. Brandt, “Ketotic episodes in glutaryl-CoA dehydrogenase deficiency (glutaric aciduria),” Pediatric Research, vol. 13, no. 9, pp. 977–981, 1979.
- S. Kölker, E. Christensen, J. V. Leonard et al., “Guideline for the diagnosis and management of glutaryl-CoA dehydrogenase deficiency (glutaric aciduria type I),” Journal of Inherited Metabolic Disease, vol. 30, no. 1, pp. 5–22, 2007.