Gateways for Glutamate Neuroprotection in Parkinson’s Disease (PD): Essential Role of EAAT3 and NCX1 Revealed in an In Vitro Model of PD
Abstract
:1. Introduction
2. Materials and Methods
2.1. Cell Culture and Treatments
2.2. Generation of HNE-Induced α-Syn Oligomers
2.3. Silencing of NCX1 and EAAT3
2.4. Cell Viability
2.5. Mitochondrial Activity
2.6. ATP Assay
2.7. Detection of Mitochondrial ROS Formation
2.8. Analysis of Cytoplasmic and Mitochondrial Ca2+ Levels
2.9. Drug and Chemicals
2.10. Data Analysis
3. Results
3.1. Effect of α-Syn and α-Syn Plus Rot on Cell Viability
3.2. Glutamate Recovery of Cell Injury, ROS Overproduction and ATP Synthesis Reduction Induced by α-Syn Plus Rot
3.3. Involvement of EAAT3 and NCX1 in the Glutamate Neuroprotection Against α-Syn Plus Rot Toxicity
3.4. Effect of Glutamate on α-Syn Plus Rot-Induced Cytoplasmic and Mitochondrial Ca2+ Increase: The Central Role of EAAT3 and NCX1
4. Discussion
Author Contributions
Funding
Acknowledgments
Conflicts of Interest
References
- Powers, R.; Lei, S.; Anandhan, A.; Marshall, D.D.; Worley, B.; Cerny, R.L.; Dodds, E.D.; Huang, Y.; Panayiotidis, M.I.; Pappa, A.; et al. Metabolic Investigations of the Molecular Mechanisms Associated with Parkinson’s Disease. Metabolites 2017, 7, 22. [Google Scholar] [CrossRef] [Green Version]
- Ramalingam, M.; Huh, Y.J.; Lee, Y.I. The Impairments of alpha-Synuclein and Mechanistic Target of Rapamycin in Rotenone-Induced SH-SY5Y Cells and Mice Model of Parkinson’s Disease. Front. Neurosci. 2019, 13, 1028. [Google Scholar] [CrossRef] [Green Version]
- Sherer, T.B.; Betarbet, R.; Testa, C.M.; Seo, B.B.; Richardson, J.R.; Kim, J.H.; Miller, G.W.; Yagi, T.; Matsuno-Yagi, A.; Greenamyre, J.T. Mechanism of toxicity in rotenone models of Parkinson’s disease. J. Neurosci. 2003, 23, 10756–10764. [Google Scholar] [CrossRef] [PubMed]
- Subramaniam, S.R.; Chesselet, M.F. Mitochondrial dysfunction and oxidative stress in Parkinson’s disease. Prog. Neurobiol. 2013, 106–107, 17–32. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Abramov, A.Y.; Gegg, M.; Grunewald, A.; Wood, N.W.; Klein, C.; Schapira, A.H. Bioenergetic consequences of PINK1 mutations in Parkinson disease. PLoS ONE 2011, 6, e25622. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Dawson, T.M.; Dawson, V.L. The role of parkin in familial and sporadic Parkinson’s disease. Mov. Disord. Off. J. Mov. Disord. Soc. 2010, 25 (Suppl. 1), S32–S39. [Google Scholar] [CrossRef] [PubMed]
- Liu, Y.; Ma, X.; Fujioka, H.; Liu, J.; Chen, S.; Zhu, X. DJ-1 regulates the integrity and function of ER-mitochondria association through interaction with IP3R3-Grp75-VDAC1. Proc. Natl. Acad. Sci. USA 2019, 116, 25322–25328. [Google Scholar] [CrossRef] [PubMed]
- Ludtmann, M.H.R.; Kostic, M.; Horne, A.; Gandhi, S.; Sekler, I.; Abramov, A.Y. LRRK2 deficiency induced mitochondrial Ca2+ efflux inhibition can be rescued by Na+/Ca2+/Li+ exchanger upregulation. Cell Death Dis. 2019, 10, 265. [Google Scholar] [CrossRef]
- Pang, S.Y.; Ho, P.W.; Liu, H.F.; Leung, C.T.; Li, L.; Chang, E.E.S.; Ramsden, D.B.; Ho, S.L. The interplay of aging, genetics and environmental factors in the pathogenesis of Parkinson’s disease. Transl. Neurodegener. 2019, 8, 23. [Google Scholar] [CrossRef]
- Verma, M.; Callio, J.; Otero, P.A.; Sekler, I.; Wills, Z.P.; Chu, C.T. Mitochondrial Calcium Dysregulation Contributes to Dendrite Degeneration Mediated by PD/LBD-Associated LRRK2 Mutants. J. Neurosci. Off. J. Soc. Neurosci. 2017, 37, 11151–11165. [Google Scholar]
- Abramov, A.Y.; Angelova, P.R. Cellular mechanisms of complex I-associated pathology. Biochem. Soc. Trans. 2019, 47, 1963–1969. [Google Scholar] [CrossRef] [PubMed]
- Chen, C.; Turnbull, D.M.; Reeve, A.K. Mitochondrial Dysfunction in Parkinson’s Disease-Cause or Consequence? Biology 2019, 8, 38. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Dey, K.; Bazala, M.A.; Kuznicki, J. Targeting mitochondrial calcium pathways as a potential treatment against Parkinson’s disease. Cell Calcium 2020, 89, 102216. [Google Scholar] [CrossRef] [PubMed]
- Keane, P.C.; Kurzawa, M.; Blain, P.G.; Morris, C.M. Mitochondrial dysfunction in Parkinson’s disease. Parkinson’s Dis. 2011, 2011, 716871. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Rocha, E.M.; de Miranda, B.; Sanders, L.H. Alpha-synuclein: Pathology, mitochondrial dysfunction and neuroinflammation in Parkinson’s disease. Neurobiol. Dis. 2018, 109, 249–257. [Google Scholar] [CrossRef]
- Schapira, A.H. Mitochondrial dysfunction in Parkinson’s disease. Cell Death Differ. 2007, 14, 1261–1266. [Google Scholar] [CrossRef]
- Scorziello, A.; Borzacchiello, D.; Sisalli, M.J.; di Martino, R.; Morelli, M.; Feliciello, A. Mitochondrial Homeostasis and Signaling in Parkinson’s Disease. Front. Aging Neurosci. 2020, 12, 100. [Google Scholar] [CrossRef] [Green Version]
- Langston, J.W.; Ballard, P.; Tetrud, J.W.; Irwin, I. Chronic Parkinsonism in humans due to a product of meperidine-analog synthesis. Science 1983, 219, 979–980. [Google Scholar] [CrossRef] [Green Version]
- Mizuno, Y.; Ohta, S.; Tanaka, M.; Takamiya, S.; Suzuki, K.; Sato, T.; Oya, H.; Ozawa, T.; Kagawa, Y. Deficiencies in complex I subunits of the respiratory chain in Parkinson’s disease. Biochem. Biophys. Res. Commun. 1989, 163, 1450–1455. [Google Scholar] [CrossRef]
- Schapira, A.H.; Cooper, J.M.; Dexter, D.; Jenner, P.; Clark, J.B.; Marsden, C.D. Mitochondrial complex I deficiency in Parkinson’s disease. Lancet 1989, 1, 1269. [Google Scholar] [CrossRef]
- Winklhofer, K.F.; Haass, C. Mitochondrial dysfunction in Parkinson’s disease. Biochim. Biophys. Acta 2010, 1802, 29–44. [Google Scholar] [CrossRef] [PubMed]
- Bastioli, G.; Piccirillo, S.; Castaldo, P.; Magi, S.; Tozzi, A.; Amoroso, S.; Calabresi, P. Selective inhibition of mitochondrial sodium-calcium exchanger protects striatal neurons from alpha-synuclein plus rotenone induced toxicity. Cell Death dis. 2019, 10, 80. [Google Scholar] [CrossRef] [PubMed]
- Bir, A.; Sen, O.; Anand, S.; Khemka, V.K.; Banerjee, P.; Cappai, R.; Sahoo, A.; Chakrabart, S. alpha-Synuclein-induced mitochondrial dysfunction in isolated preparation and intact cells: Implications in the pathogenesis of Parkinson’s disease. J. Neurochem. 2014, 131, 868–877. [Google Scholar] [CrossRef] [PubMed]
- Chinta, S.J.; Mallajosyula, J.K.; Rane, A.; Andersen, J.K. Mitochondrial alpha-synuclein accumulation impairs complex I function in dopaminergic neurons and results in increased mitophagy in vivo. Neurosci. Lett. 2010, 486, 235–239. [Google Scholar] [CrossRef] [Green Version]
- Devi, L.; Raghavendran, V.; Prabhu, B.M.; Avadhani, N.G.; Anandatheerthavarada, H.K. Mitochondrial import and accumulation of alpha-synuclein impair complex I in human dopaminergic neuronal cultures and Parkinson disease brain. J. Biol. Chem. 2008, 283, 9089–9100. [Google Scholar] [CrossRef] [Green Version]
- Martinez, J.H.; Fuentes, F.; Vanasco, V.; Alvarez, S.; Alaimo, A.; Cassina, A.; Leskow, F.C.; Velazquez, F. Alpha-synuclein mitochondrial interaction leads to irreversible translocation and complex I impairment. Arch. Biochem. Biophys. 2018, 651, 1–12. [Google Scholar] [CrossRef] [Green Version]
- Reeve, A.K.; Ludtmann, M.H.; Angelova, P.R.; Simcox, E.M.; Horrocks, M.H.; Klenerman, D.; Gandhi, S.; Turnbull, D.M.; Abramov, A.Y. Aggregated alpha-synuclein and complex I deficiency: Exploration of their relationship in differentiated neurons. Cell Death Dis. 2015, 6, e1820. [Google Scholar] [CrossRef]
- Mirzaei, H.; Schieler, J.L.; Rochet, J.C.; Regnier, F. Identification of rotenone-induced modifications in alpha-synuclein using affinity pull-down and tandem mass spectrometry. Anal. Chem. 2006, 78, 2422–2431. [Google Scholar] [CrossRef]
- Yuan, Y.H.; Yan, W.F.; Sun, J.D.; Huang, J.Y.; Mu, Z.; Chen, N.H. The molecular mechanism of rotenone-induced alpha-synuclein aggregation: Emphasizing the role of the calcium/GSK3beta pathway. Toxicol. Lett. 2015, 233, 163–171. [Google Scholar] [CrossRef]
- Billingsley, K.J.; Barbosa, I.A.; Bandres-Ciga, S.; Quinn, J.P.; Bubb, V.J.; Deshpande, C.; Botia, J.A.; Reynolds, R.H.; Zhang, D.; Simpson, M.A.; et al. Mitochondria function associated genes contribute to Parkinson’s Disease risk and later age at onset. NPJ Parkinson’s Dis. 2019, 5, 8. [Google Scholar] [CrossRef] [Green Version]
- Larsen, S.B.; Hanss, Z.; Kruger, R. The genetic architecture of mitochondrial dysfunction in Parkinson’s disease. Cell Tissue Res. 2018, 373, 21–37. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Beal, M.F. Bioenergetic approaches for neuroprotection in Parkinson’s disease. Ann. Neurol. 2003, 53 (Suppl. 3), S39–S47. [Google Scholar] [CrossRef] [PubMed]
- Quansah, E.; Peelaerts, W.; Langston, J.W.; Simon, D.K.; Colca, J.; Brundin, P. Targeting energy metabolism via the mitochondrial pyruvate carrier as a novel approach to attenuate neurodegeneration. Mol. Neurodegener. 2018, 13, 28. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Norwitz, N.G.; Hu, M.T.; Clarke, K. The Mechanisms by Which the Ketone Body D-beta-Hydroxybutyrate May Improve the Multiple Cellular Pathologies of Parkinson’s Disease. Front. Nutr. 2019, 6, 63. [Google Scholar] [CrossRef]
- Zielinski, L.P.; Smith, A.C.; Smith, A.G.; Robinson, A.J. Metabolic flexibility of mitochondrial respiratory chain disorders predicted by computer modelling. Mitochondrion 2016, 31, 45–55. [Google Scholar] [CrossRef]
- Kim, A.Y.; Baik, E.J. Glutamate Dehydrogenase as a Neuroprotective Target Against Neurodegeneration. Neurochem. Res. 2019, 44, 147–153. [Google Scholar] [CrossRef]
- Ehinger, J.K.; Piel, S.; Ford, R.; Karlsson, M.; Sjovall, F.; Frostner, E.A.; Morota, S.; Taylor, R.W.; Turnbull, D.M.; Cornell, C.; et al. Cell-permeable succinate prodrugs bypass mitochondrial complex I deficiency. Nat. Commun. 2016, 7, 12317. [Google Scholar] [CrossRef] [Green Version]
- Magi, S.; Arcangeli, S.; Castaldo, P.; Nasti, A.A.; Berrino, L.; Piegari, E.; Bernardini, R.; Amoroso, S.; Lariccia, V. Glutamate-induced ATP synthesis: Relationship between plasma membrane Na+/Ca2+ exchanger and excitatory amino acid transporters in brain and heart cell models. Mol. Pharmacol. 2013, 84, 603–614. [Google Scholar] [CrossRef] [Green Version]
- Maiolino, M.; Castaldo, P.; Lariccia, V.; Piccirillo, S.; Amoroso, S.; Magi, S. Essential role of the Na+-Ca2+ exchanger (NCX) in glutamate-enhanced cell survival in cardiac cells exposed to hypoxia/reoxygenation. Sci. Rep. 2017, 7, 13073. [Google Scholar] [CrossRef]
- Piccirillo, S.; Castaldo, P.; Macri, M.L.; Amoroso, S.; Magi, S. Glutamate as a potential “survival factor” in an in vitro model of neuronal hypoxia/reoxygenation injury: Leading role of the Na+/Ca2+ exchanger. Cell Death Dis. 2018, 9, 731. [Google Scholar] [CrossRef]
- Magi, S.; Lariccia, V.; Castaldo, P.; Arcangeli, S.; Nasti, A.A.; Giordano, A.; Amoroso, S. Physical and functional interaction of NCX1 and EAAC1 transporters leading to glutamate-enhanced ATP production in brain mitochondria. PLoS ONE 2012, 7, e34015. [Google Scholar] [CrossRef] [PubMed]
- Danbolt, N.C. Glutamate uptake. Prog. Neurobiol. 2001, 65, 1–105. [Google Scholar] [CrossRef]
- Magi, S.; Piccirillo, S.; Amoroso, S.; Lariccia, V. Excitatory Amino Acid Transporters (EAATs): Glutamate Transport and Beyond. Int. J. Mol. Sci. 2019, 20, 5674. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Blaustein, M.P.; Lederer, W.J. Sodium/calcium exchange: Its physiological implications. Physiol. Rev. 1999, 79, 763–854. [Google Scholar] [CrossRef] [PubMed]
- Philipson, K.D.; Nicoll, D.A. Sodium-calcium exchange: A molecular perspective. Annu. Rev. Physiol. 2000, 62, 111–133. [Google Scholar] [CrossRef]
- Gobbi, P.; Castaldo, P.; Minelli, A.; Salucci, S.; Magi, S.; Corcione, E.; Amoroso, S. Mitochondrial localization of Na+/Ca2+ exchangers NCX1-3 in neurons and astrocytes of adult rat brain in situ. Pharmacol. Res. 2007, 56, 556–565. [Google Scholar] [CrossRef]
- Lytton, J. Na+/Ca2+ exchangers: Three mammalian gene families control Ca2+ transport. Biochem. J. 2007, 406, 365–382. [Google Scholar] [CrossRef]
- Minelli, A.; Castaldo, P.; Gobbi, P.; Salucci, S.; Magi, S.; Amoroso, S. Cellular and subcellular localization of Na+-Ca2+ exchanger protein isoforms, NCX1, NCX2, and NCX3 in cerebral cortex and hippocampus of adult rat. Cell Calcium 2007, 41, 221–234. [Google Scholar] [CrossRef]
- Quednau, B.D.; Nicoll, D.A.; Philipson, K.D. Tissue specificity and alternative splicing of the Na+/Ca2+ exchanger isoforms NCX1, NCX2, and NCX3 in rat. Am. J. Physiol. 1997, 272, C1250–C1261. [Google Scholar] [CrossRef]
- Magi, S.; Piccirillo, S.; Preziuso, A.; Amoroso, S.; Lariccia, V. Mitochondrial localization of NCXs: Balancing calcium and energy homeostasis. Cell Calcium 2020, 86, 102162. [Google Scholar] [CrossRef]
- Piccirillo, S.; Magi, S.; Castaldo, P.; Preziuso, A.; Lariccia, V.; Amoroso, S. NCX and EAAT transporters in ischemia: At the crossroad between glutamate metabolism and cell survival. Cell Calcium 2020, 86, 102160. [Google Scholar] [CrossRef] [PubMed]
- Roberts, R.F.; Wade-Martins, R.; Alegre-Abarrategui, J. Direct visualization of alpha-synuclein oligomers reveals previously undetected pathology in Parkinson’s disease brain. Brain J. Neurol. 2015, 138, 1642–1657. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Staiano, R.I.; Granata, F.; Secondo, A.; Petraroli, A.; Loffredo, S.; Frattini, A.; Annunziato, L.; Marone, G.; Triggiani, M. Expression and function of Na+/Ca2+ exchangers 1 and 3 in human macrophages and monocytes. Eur. J. Immunol. 2009, 39, 1405–1418. [Google Scholar] [CrossRef] [PubMed]
- van Meerloo, J.; Kaspers, G.J.; Cloos, J. Cell sensitivity assays: The MTT assay. Methods Mol. Biol. 2011, 731, 237–245. [Google Scholar]
- Esteras, N.; Rohrer, J.D.; Hardy, J.; Wray, S.; Abramov, A.Y. Mitochondrial hyperpolarization in iPSC-derived neurons from patients of FTDP-17 with 10+16 MAPT mutation leads to oxidative stress and neurodegeneration. Redox Biol. 2017, 12, 410–422. [Google Scholar] [CrossRef]
- Diaz-Corrales, F.J.; Asanuma, M.; Miyazaki, I.; Miyoshi, K.; Ogawa, N. Rotenone induces aggregation of gamma-tubulin protein and subsequent disorganization of the centrosome: Relevance to formation of inclusion bodies and neurodegeneration. Neuroscience 2005, 133, 117–135. [Google Scholar] [CrossRef] [Green Version]
- Sherer, T.B.; Betarbet, R.; Stout, A.K.; Lund, S.; Baptista, M.; Panov, A.V.; Cookson, M.R.; Greenamyre, J.T. An in vitro model of Parkinson’s disease: Linking mitochondrial impairment to altered alpha-synuclein metabolism and oxidative damage. J. Neurosci. Off. J. Soc. Neurosci. 2002, 22, 7006–7015. [Google Scholar]
- Dias, V.; Junn, E.; Mouradian, M.M. The role of oxidative stress in Parkinson’s disease. J. Parkinson’s Dis. 2013, 3, 461–491. [Google Scholar] [CrossRef] [Green Version]
- Shimamoto, K.; Lebrun, B.; Yasuda-Kamatani, Y.; Sakaitani, M.; Shigeri, Y.; Yumoto, N.; Nakajima, T. DL-threo-beta-benzyloxyaspartate, a potent blocker of excitatory amino acid transporters. Mol. Pharmacol. 1998, 53, 195–201. [Google Scholar] [CrossRef] [Green Version]
- Iwamoto, T.; Inoue, Y.; Ito, K.; Sakaue, T.; Kita, S.; Katsuragi, T. The exchanger inhibitory peptide region-dependent inhibition of Na+/Ca2+ exchange by SN-6 [2-[4-(4-nitrobenzyloxy)benzyl]thiazolidine-4-carboxylic acid ethyl ester], a novel benzyloxyphenyl derivative. Molecular Pharmacol. 2004, 66, 45–55. [Google Scholar] [CrossRef] [Green Version]
- Abeti, R.; Abramov, A.Y. Mitochondrial Ca2+ in neurodegenerative disorders. Pharmacol. Res. 2015, 99, 377–381. [Google Scholar] [CrossRef] [PubMed]
- Angelova, P.R.; Ludtmann, M.H.; Horrocks, M.H.; Negoda, A.; Cremades, N.; Klenerman, D.; Dobson, C.M.; Wood, N.W.; Pavlov, E.V.; Gandhi, S.; et al. Ca2+ is a key factor in alpha-synuclein-induced neurotoxicity. J. Cell Sci. 2016, 129, 1792–1801. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Freestone, P.S.; Chung, K.K.; Guatteo, E.; Mercuri, N.B.; Nicholson, L.F.; Lipski, J. Acute action of rotenone on nigral dopaminergic neurons—Involvement of reactive oxygen species and disruption of Ca2+ homeostasis. Eur. J. Neurosci. 2009, 30, 1849–1859. [Google Scholar] [CrossRef] [PubMed]
- Luth, E.S.; Stavrovskaya, I.G.; Bartels, T.; Kristal, B.S.; Selkoe, D.J. Soluble, prefibrillar alpha-synuclein oligomers promote complex I-dependent, Ca2+-induced mitochondrial dysfunction. J. Biol. Chem. 2014, 289, 21490–21507. [Google Scholar] [CrossRef] [Green Version]
- Macdonald, R.; Barnes, K.; Hastings, C.; Mortiboys, H. Mitochondrial abnormalities in Parkinson’s disease and Alzheimer’s disease: Can mitochondria be targeted therapeutically? Biochem. Soc. Trans. 2018, 46, 891–909. [Google Scholar] [CrossRef] [PubMed]
- Nunnari, J.; Suomalainen, A. Mitochondria: In sickness and in health. Cell 2012, 148, 1145–1159. [Google Scholar] [CrossRef] [Green Version]
- Janetzky, B.; Hauck, S.; Youdim, M.B.; Riederer, P.; Jellinger, K.; Pantucek, F.; Zochling, R.; Boissl, K.W.; Reichmann, H. Unaltered aconitase activity, but decreased complex I activity in substantia nigra pars compacta of patients with Parkinson’s disease. Neurosci. Lett. 1994, 169, 126–128. [Google Scholar] [CrossRef]
- Mann, V.M.; Cooper, J.M.; Daniel, S.E.; Srai, K.; Jenner, P.; Marsden, C.D.; Schapira, A.H. Complex I, iron, and ferritin in Parkinson’s disease substantia nigra. Ann. Neurol. 1994, 36, 876–881. [Google Scholar] [CrossRef]
- Schapira, A.H.; Cooper, J.M.; Dexter, D.; Clark, J.B.; Jenner, P.; Marsden, C.D. Mitochondrial complex I deficiency in Parkinson’s disease. J. Neurochemistry 1990, 54, 823–827. [Google Scholar] [CrossRef]
- Greenamyre, J.T.; Sherer, T.B.; Betarbet, R.; Panov, A.V. Complex I and Parkinson’s disease. IUBMB Life 2001, 52, 135–141. [Google Scholar] [CrossRef]
- Betarbet, R.; Sherer, T.B.; MacKenzie, G.; Garcia-Osuna, M.; Panov, A.V.; Greenamyre, J.T. Chronic systemic pesticide exposure reproduces features of Parkinson’s disease. Nat. Neurosci. 2000, 3, 1301–1306. [Google Scholar] [CrossRef] [PubMed]
- Melachroinou, K.; Xilouri, M.; Emmanouilidou, E.; Masgrau, R.; Papazafiri, P.; Stefanis, L.; Vekrellis, K. Deregulation of calcium homeostasis mediates secreted alpha-synuclein-induced neurotoxicity. Neurobiol. Aging 2013, 34, 2853–2865. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Wang, X.J.; Xu, J.X. Possible involvement of Ca2+ signaling in rotenone-induced apoptosis in human neuroblastoma SH-SY5Y cells. Neurosci. Lett. 2005, 376, 127–132. [Google Scholar] [CrossRef] [PubMed]
- Zhao, R.Z.; Jiang, S.; Zhang, L.; Yu, Z.B. Mitochondrial electron transport chain, ROS generation and uncoupling (Review). Int. J. Mol. Med. 2019, 44, 3–15. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Dringen, R.; Pfeiffer, B.; Hamprecht, B. Synthesis of the antioxidant glutathione in neurons: Supply by astrocytes of CysGly as precursor for neuronal glutathione. J. Neurosci. Off. J. Soc. Neurosci. 1999, 19, 562–569. [Google Scholar] [CrossRef]
- McBean, G.J. Cysteine, Glutathione, and Thiol Redox Balance in Astrocytes. Antioxidants 2017, 6, 62. [Google Scholar] [CrossRef] [Green Version]
- Grunewald, A.; Kumar, K.R.; Sue, C.M. New insights into the complex role of mitochondria in Parkinson’s disease. Prog. Neurobiol. 2019, 177, 73–93. [Google Scholar] [CrossRef] [PubMed]
- Surmeier, D.J.; Schumacker, P.T. Calcium, bioenergetics, and neuronal vulnerability in Parkinson’s disease. J. Biol. Chem. 2013, 288, 10736–10741. [Google Scholar] [CrossRef] [Green Version]
- Cali, T.; Ottolini, D.; Brini, M. Calcium signaling in Parkinson’s disease. Cell Tissue Res. 2014, 357, 439–454. [Google Scholar] [CrossRef]
- He, Y.; Hof, P.R.; Janssen, W.G.; Rothstein, J.D.; Morrison, J.H. Differential synaptic localization of GluR2 and EAAC1 in the macaque monkey entorhinal cortex: A postembedding immunogold study. Neurosci. Lett. 2001, 311, 161–164. [Google Scholar] [CrossRef]
- Holmseth, S.; Dehnes, Y.; Huang, Y.H.; Follin-Arbelet, V.V.; Grutle, N.J.; Mylonakou, M.N.; Plachez, C.; Zhou, Y.; Furness, D.N.; Bergles, D.E.; et al. The density of EAAC1 (EAAT3) glutamate transporters expressed by neurons in the mammalian CNS. J. Neurosci. Off. J. Soc. Neurosci. 2012, 32, 6000–6013. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Underhill, S.M.; Wheeler, D.S.; Li, M.; Watts, S.D.; Ingram, S.L.; Amara, S.G. Amphetamine modulates excitatory neurotransmission through endocytosis of the glutamate transporter EAAT3 in dopamine neurons. Neuron 2014, 83, 404–416. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Bauer, D.E.; Jackson, J.G.; Genda, E.N.; Montoya, M.M.; Yudkoff, M.; Robinson, M.B. The glutamate transporter, GLAST, participates in a macromolecular complex that supports glutamate metabolism. Neurochem. Int. 2012, 61, 566–574. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Gegelashvili, G.; Bjerrum, O.J. Glutamate transport system as a key constituent of glutamosome: Molecular pathology and pharmacological modulation in chronic pain. Neuropharmacology 2019, 161, 107623. [Google Scholar] [CrossRef] [PubMed]
- Genda, E.N.; Jackson, J.G.; Sheldon, A.L.; Locke, S.F.; Greco, T.M.; O’Donnell, J.C.; Spruce, L.A.; Xiao, R.; Guo, W.; Putt, M.; et al. Co-compartmentalization of the astroglial glutamate transporter, GLT-1, with glycolytic enzymes and mitochondria. J. Neurosci. Off. J. Soc. Neurosci. 2011, 31, 18275–18288. [Google Scholar]
- Robinson, M.B.; Lee, M.L.; DaSilva, S. Glutamate Transporters and Mitochondria: Signaling, Co-compartmentalization, Functional Coupling, and Future Directions. Neurochem. Res. 2020, 45, 526–540. [Google Scholar] [CrossRef] [PubMed]
- Kirischuk, S.; Kettenmann, H.; Verkhratsky, A. Membrane currents and cytoplasmic sodium transients generated by glutamate transport in Bergmann glial cells. Pflug. Arch. Eur. J. Physiol. 2007, 454, 245–252. [Google Scholar] [CrossRef]
- Verkhratsky, A. Physiology of neuronal-glial networking. Neurochem. Int. 2010, 57, 332–343. [Google Scholar] [CrossRef]
- Plaitakis, A.; Shashidharan, P. Glutamate transport and metabolism in dopaminergic neurons of substantia nigra: Implications for the pathogenesis of Parkinson’s disease. J. Neurol. 2000, 247 (Suppl. 2), II25–II35. [Google Scholar] [CrossRef]
- Castaldo, P.; Macri, M.L.; Lariccia, V.; Matteucci, A.; Maiolino, M.; Gratteri, S.; Amoroso, S.; Magi, S. Na+/Ca2+ exchanger 1 inhibition abolishes ischemic tolerance induced by ischemic preconditioning in different cardiac models. Eur. J. Pharmacol. 2017, 794, 246–256. [Google Scholar] [CrossRef]
- Kritis, A.A.; Stamoula, E.G.; Paniskaki, K.A.; Vavilis, T.D. Researching glutamate—Induced cytotoxicity in different cell lines: A comparative/collective analysis/study. Front. Cell. Neurosci. 2015, 9, 91. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Nampoothiri, M.; Reddy, N.D.; John, J.; Kumar, N.; Nampurath, G.K.; Chamallamudi, M.R. Insulin blocks glutamate-induced neurotoxicity in differentiated SH-SY5Y neuronal cells. Behav. Neurol. 2014, 2014, 674164. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Sun, Z.W.; Zhang, L.; Zhu, S.J.; Chen, W.C.; Mei, B. Excitotoxicity effects of glutamate on human neuroblastoma SH-SY5Y cells via oxidative damage. Neurosci. Bull. 2010, 26, 8–16. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Cunha, M.P.; Lieberknecht, V.; Ramos-Hryb, A.B.; Olescowicz, G.; Ludka, F.K.; Tasca, C.I.; Gabilan, N.H.; Rodrigues, A.L. Creatine affords protection against glutamate-induced nitrosative and oxidative stress. Neurochem. Int. 2016, 95, 4–14. [Google Scholar] [CrossRef] [PubMed]
- Gao, M.; Zhang, W.C.; Liu, Q.S.; Hu, J.J.; Liu, G.T.; Du, G.H. Pinocembrin prevents glutamate-induced apoptosis in SH-SY5Y neuronal cells via decrease of bax/bcl-2 ratio. Eur. J. Pharmacol. 2008, 591, 73–79. [Google Scholar] [CrossRef]
- Magi, S.; Piccirillo, S.; Maiolino, M.; Lariccia, V. Amoroso, NCX1 and EAAC1 transporters are involved in the protective action of glutamate in an in vitro Alzheimer’s disease-like model. Cell Calcium 2020, 91, 102268. [Google Scholar] [CrossRef] [PubMed]
- McKenna, M.C. The glutamate-glutamine cycle is not stoichiometric: Fates of glutamate in brain. J. Neurosci. Res. 2007, 85, 3347–3358. [Google Scholar] [CrossRef] [PubMed]
- Dienel, G.A.; Cruz, N.F. Astrocyte activation in working brain: Energy supplied by minor substrates. Neurochem. Int. 2006, 48, 586–595. [Google Scholar] [CrossRef]
- Parpura, V.; Verkhratsky, A. Astrocytes revisited: Concise historic outlook on glutamate homeostasis and signaling. Croat. Med. J. 2012, 53, 518–528. [Google Scholar] [CrossRef] [Green Version]
- Moussawi, K.; Riegel, A.; Nair, S.; Kalivas, P.W. Extracellular glutamate: Functional compartments operate in different concentration ranges. Front. Syst. Neurosci. 2011, 5, 94. [Google Scholar] [CrossRef] [Green Version]
- Ha, J.S.; Lee, C.S.; Maeng, J.S.; Kwon, K.S.; Park, S.S. Chronic glutamate toxicity in mouse cortical neuron culture. Brain Res. 2009, 1273, 138–143. [Google Scholar] [CrossRef] [PubMed]
- McKenna, M.C.; Sonnewald, U.; Huang, X.; Stevenson, J.; Zielke, H.R. Exogenous glutamate concentration regulates the metabolic fate of glutamate in astrocytes. J. Neurochem. 1996, 66, 386–393. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Hawkins, R.A. The blood-brain barrier and glutamate. Am. J. Clin. Nutr. 2009, 90, 867S–874S. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Onaolapo, A.Y.; Onaolapo, O.J. Dietary glutamate and the brain: In the footprints of a Jekyll and Hyde molecule. Neurotoxicology 2020, 80, 93–104. [Google Scholar] [CrossRef] [PubMed]
- Kim, A.Y.; Jeong, K.H.; Lee, J.H.; Kang, Y.; Lee, S.H.; Baik, E.J. Glutamate dehydrogenase as a neuroprotective target against brain ischemia and reperfusion. Neuroscience 2017, 340, 487–500. [Google Scholar] [CrossRef]
- Rink, C.; Gnyawali, S.; Stewart, R.; Teplitsky, S.; Harris, H.; Roy, S.; Sen, C.K.; Khanna, S. Glutamate oxaloacetate transaminase enables anaplerotic refilling of TCA cycle intermediates in stroke-affected brain. FASEB J. Off. Publ. Fed. Am. Soc. Exp. Biol. 2017, 31, 1709–1718. [Google Scholar] [CrossRef] [Green Version]
- Jantas, D.; Greda, A.; Golda, S.; Korostynski, M.; Grygier, B.; Roman, A.; Pilc, A.; Lason, W. Neuroprotective effects of metabotropic glutamate receptor group II and III activators against MPP+-induced cell death in human neuroblastoma SH-SY5Y cells: The impact of cell differentiation state. Neuropharmacology 2014, 83, 36–53. [Google Scholar] [CrossRef]
- Verkhratsky, A.; Nedergaard, M. Physiology of Astroglia. Physiol. Rev. 2018, 98, 239–389. [Google Scholar] [CrossRef]
- Iovino, L.; Tremblay, M.E.; Civiero, L. Glutamate-induced excitotoxicity in Parkinson’s disease: The role of glial cells. J. Pharmacol. Sci. 2020, in press. [Google Scholar] [CrossRef]
- Verkhratsky, A.; Parpura, V.; Pekna, M.; Pekny, M.; Sofroniew, M. Glia in the pathogenesis of neurodegenerative diseases. Biochem. Soc. Trans. 2014, 42, 1291–1301. [Google Scholar] [CrossRef]
- Rodriguez, J.J.; Verkhratsky, A. Neuroglial roots of neurodegenerative diseases? Mol. Neurobiol. 2011, 43, 87–96. [Google Scholar] [CrossRef] [PubMed]
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Piccirillo, S.; Magi, S.; Preziuso, A.; Castaldo, P.; Amoroso, S.; Lariccia, V. Gateways for Glutamate Neuroprotection in Parkinson’s Disease (PD): Essential Role of EAAT3 and NCX1 Revealed in an In Vitro Model of PD. Cells 2020, 9, 2037. https://doi.org/10.3390/cells9092037
Piccirillo S, Magi S, Preziuso A, Castaldo P, Amoroso S, Lariccia V. Gateways for Glutamate Neuroprotection in Parkinson’s Disease (PD): Essential Role of EAAT3 and NCX1 Revealed in an In Vitro Model of PD. Cells. 2020; 9(9):2037. https://doi.org/10.3390/cells9092037
Chicago/Turabian StylePiccirillo, Silvia, Simona Magi, Alessandra Preziuso, Pasqualina Castaldo, Salvatore Amoroso, and Vincenzo Lariccia. 2020. "Gateways for Glutamate Neuroprotection in Parkinson’s Disease (PD): Essential Role of EAAT3 and NCX1 Revealed in an In Vitro Model of PD" Cells 9, no. 9: 2037. https://doi.org/10.3390/cells9092037