University of Southern California Dissertations and Theses

Increased susceptibility of glutathione peroxidase-1 transgenic mice to kainic acid-related seizure activity and hippocampal neuronal cell death due to direct activation of the NMDA receptor by GSSG

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Increased susceptibility of glutathione peroxidase-1 transgenic mice to kainic acid-related seizure activity and hippocampal neuronal cell death due to direct activation of the NMDA receptor by GSSG

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Content INCREASED SUSCEPTIBILITY OF GLUTATHIONE PEROXIDASE-1 TRANSGENIC MICE TO KAINIC ACID-RELATED SEIZURE ACTIVITY AND HIPPOCAMPAL NEURONAL CELL DEATH DUE TO DIRECT ACTIVATION OF THE NMDA RECEPTOR BY GSSG by Rapee Boonplueang A Dissertation Presented to the FACULTY OF THE GRADUATE SCHOOL UNIVERSITY OF SOUTHERN CALIFORNIA In Partial Fulfillment of the Requirements for the Degree DOCTOR OF PHILOSOPHY (MOLECULAR BIOLOGY) December 2004 Copyright 2004 Rapee Boonplueang Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. UMI Number: 3155383 INFORMATION TO USERS The quality of this reproduction is dependent upon the quality of the copy submitted. Broken or indistinct print, colored or poor quality illustrations and photographs, print bleed-through, substandard margins, and improper alignment can adversely affect reproduction. In the unlikely event that the author did not send a complete manuscript and there are missing pages, these will be noted. Also, if unauthorized copyright material had to be removed, a note will indicate the deletion. ® UMI UMI Microform 3155383 Copyright 2005 by ProQuest Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code. ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, Ml 48106-1346 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. DEDICATION to My parents, brother and sister All teachers All experimental subjects and In memory of William E. Trusten Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ACKNOWLEDGEMENTS I have been told that I could be as much creatively expressive as I would like to when it comes down to writing the acknowledgements of my thesis. Since I have been very fortunate to have my life touched by so many wonderful people during my Ph.D. at USC, it is going to be a long one. First and foremost, I would very much like to thank my advisor, Dr. Julie K. Andersen, for her guidance, her total understanding of the nature of research work and never having given me a hard time when I did not have any new results to report as well as moral and financial support. I wish to thank Dr. John Petruska for his guidance and graciously accepting to be the co-chair of my dissertation committee. I also wish to thank my other committee members, Dr. Donald Arnold and Dr. Christian Pike for their valuable suggestions. I would very much like to thank Dr. John Walsh and Garnik Akopian for their wonderful electrophysiological work contributed to this project. I would also like to thank Dr. Shelly Lu and John Kuhlenkamp for their HPLC work. We could not have proven our hypothesis without their contributions. I wish to thank all my classmates, Mike Hsu, Deepali Shinde, Jaichandar Subramanian and Cathryn Harris for not leaving me behind when we were taking our coursework, written and oral qualifying exams. Thank you, guys. iii Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. I wish to thank all wonderful people in the Andersen lab, also including all summer students, for their helps and supports. Fang Stevenson, I would not be able to finish my animal works without your help. Dr. Yongqin Wu, thank you for techniques and tips you taught me. Dr. Deepinder Kaur, thank you for being such a nice bay-mate, your moral support and for never having complained when I played music too loud. Dr. Veena Viswanath, you made my first year in the lab easy for me. I wish to thank the faculty and the administrative staff of the Molecular and Computational Biology program, USC, especially the late Bill Trusten and Eleni Yokas, for their help and support. I wish to thank all wonderful people from all over the world I have had a privilege to know and work with at the Buck Institute for Age Research, especially, Andersen lab’s administrative assistants and vivarium staff. Also, I would like to thank Dr. Sylvia Chen for tissue culture tips and techniques and giving me chances to show off my Thai cooking skill. Dr. Corina Marx, thank you for Spanish 101 and your rhubarb pies, they are the best. Dr. Enrique Samper, thank you for the conversation we had that night. It literally saved my life. Christina Yau, thank you for all entertainment reports. This place is where I have come to realize that, regardless of their faith, race or nationality, we all are the same. I would also like to thank my friends. Ped, thank you for being there for me through ups and downs in my life and being such an understanding and wonderful friend. Sometimes, I wondered if you have ever had a negative thought. Puay, thanks for all oversea calls you made to me all these years but please remember that there is iv Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. a 15-hour difference. Kang, I could not imagine my first year at USC without your help. Obe, your stories always fascinated me. Tock, Pop and Patty, thanks for your emails and postcards. Lee, sorry, I missed your wedding day. My SC-12 friends, it is good that we are finally reuniting and thanks for your moral support. Bee, I’m going to miss your wedding as well, sorry. Thanks to Grant Welling and Daniel Comb for those unforgettable experiences. Also thanks to all new friends at the volleyball in Petaluma, if it was not for you, I would have gone insane. I would like to thank the Department of Biology, Faculty of Science, Mahidol University and the government of Thailand for giving me an opportunity to broaden my horizon. Last but not least, I would very much like to thank my parents, my brother and sister as well as my extended family. I would never have come this far without their unconditional love and support. All words in the dictionary are not enough to describe how fortunate I am to have them as a part of my life. Mom and dad, I have finally made it! Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. TABLE OF CONTENTS DEDICATION................................................................................................................... ii ACKNOWLEDGEMENTS ..........................................................................................iii LIST OF TABLES ....................................................................................................... viii LIST OF FIG U R ES......................................................................................................... ix A BSTRA CT..................................................................................................................... xi CHAPTER ONE Introduction to the Dissertation Research and Literature Review ............................ 1 Introduction to the Dissertation Research .........................................................1 Literature Review ............................................................................................... 2 1. Free radicals, reactive oxygen species (ROS), reactive nitrogen species (RNS) antioxidants and oxidative stress .................................................................................. 2 1.1. Terminology .......................................................................2 1.2. Sources of ROS and RNS .................................................3 1.1.1. Exogenous sources ............................................ 3 1.1.2. Endogenous sources .......................................... 3 1.3. R O S .......................................................................................4 1.3.1. Superoxide radical ............................................ 4 1.3.2. Hydrogen peroxide ........................................... 5 1.3.3. Hydroxyl radical ............................................... 6 1.4. ROS and DNA modification ............................................ 7 1.5. ROS and protein modification .........................................8 1.6. ROS and lipid modification ............................................. 8 1.7. ROS and apoptosis .............................................................9 1.8. Anti oxidants in the brain .............................................. 10 2. Glutathione peroxidase (GSHPx) ..................................................10 2.1. History .............................................................................. 10 2.2. GSHPx isoforms................................................................ 12 2.2.1. Cytosolic GSHPx .......................................... 12 2.2.2. Gastrointestinal GSHPx ..................................13 2.2.3. Glycosylated G S H P x....................................... 14 2.2.4. Phospholipid hydroperoxide GSHPx ...........14 2.2.5. Sperm nuclei GSHPx ......................................15 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2.3. Genetically altered GSHPx-1 models ....................... 15 2.3.1. Overexpression of GSHPx ............................. 15 2.3.2. GSHPx-1 knockout .........................................22 3. Kainic acid (KA) ............................................................................25 3.1. KA-induced seizures and neuronal cell death ............ 25 3.2. KA toxicity and oxidative stress ................................... 27 3.3. KA receptors units .......................................................... 29 CHAPTER TWO GSHPx-1 Transgenic mice are more susceptible to KA-induced toxicity ............. 31 Summary .............................................................................................................31 Introduction ........................................................................................................ 32 Materials and Methods .....................................................................................33 Results ................................................................................................................42 Discussion ........................................................................................................ 51 CHAPTER THREE GSHPx-1 Transgenic Mice Exhibit Increased Hippocampal GSSG Levels Resulting in Increased Susceptibility to KA Toxicity ................................................................................................................56 Summary .............................................................................................................56 Introduction ........................................................................................................ 57 Materials and Methods ......................................................................................58 Results .................................................................................................................68 Discussion .......................................................................................................... 82 CHAPTER FOUR Conclusions ..................................................................................................................... 88 REFERENCES ................................................................................................................91 v ii Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. LIST OF TABLES Table 1. ROS of interest in oxidative stress ..................................................................5 Table 2. Oligonucleotide primers for PCR analysis ...................................................35 viii Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. LIST OF FIGURES Figure 1. Pathways of ROS generation ....................................................................6 Figure 2. Schematic of the GSH metabolic pathway ........................................... 12 Figure 3. PCR analysis: templates, oligonucleotide primers and PCR products ......................................................................................36 Figure 4. PCR products ............................................................................................43 Figure 5. Hippocampal GSHPx activity in HM, HT and WT animals .............. 44 Figure 6. Assessment of KA-induced seizure activity in KA-treated animals ................................................................................... 45 Figure 7. Increased GSHPx activity results in increased KA-induced seizure activity ................................................ 47 Figure 8. Increased GSFIPx activity results in increased KA-induced neuronal cell death ............................................48 Figure 9. Assessment of hippocampal neuronal cell death ................................49 Figure 10. Increased GSHPx activity also results in increased susceptibility to KA in vivo ..................................................................... 50 Figure 11. Brain synaptosomal H 2O2 levels .......................................................... 68 Figure 12. Basal brain GSH/GSSG ratios in HM, HT and WT animals ...........69 Figure 13. Hippocampal GSSG and GSH levels .................................................. 71 Figure 14. Alterations in activity levels of enzymes involving in glutathione metabolism in GSHPx-1 transgenic and wild-type brains ................................................................................... 72 Figure 15. Extracellular GSSG levels ..................................................................... 74 ix Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Figure 16. Absolute and relative dithiothreitol (DTT)-induced changes in NMDA-evoked response in CA3 neurons from WT and HM hippocampal slices .................................................................................... 76 Figure 17. Western blot analysis of NMDA receptor subunits ...........................78 Figure 18. GSSG-evoked electrophysiological responses in WT CA3 neurons .......................................................................................79 Figure 19. GSSG induced response acts via direct NMDA receptor activation in CA3 neurons from WT mice ............................................ 81 x Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ABSTRACT Glutathione peroxidase (GSHPx) has been demonstrated in several in vivo studies to reduce both the risk and severity of oxidative stress mediated tissue damage. The neurotoxin kainic acid (KA) has been suggested to elicit its toxic effects through generation of oxidative stress. In this study we report that overexpression of murine GSHPx-1 in transgenic mice, resulting in a 4-6 fold increase in enzyme activity, surprisingly results in increased rather than decreased KA susceptibility including increased seizure activity and neuronal hippocampal damage. Isolated transgenic primary hippocampal culture neurons also display increased susceptibility to KA treatment compared with those isolated from wild- type animals. Our subsequent studies suggest that this is due to alterations in enzymes involved in the glutathione (GSH)-regeneration cycle including GSHPx, glutathione reductase (GR), glucose-6-phosphate dehydrogenase (G6PD) and 6- phosphogluconate dehydrogenase (6PGD), resulting in elevated glutathione disulfide (GSSG) levels leading to alterations in the glutathione redox state (GSH/GSSG ratio) and a shift in the cellular redox environment. We also observed increased basal N- methyl-D-aspartate (NMDA)-evoked response in transgenic hippocampal slices independent of the cellular redox status or the number of NMDA receptors present. Our data suggests that, in response to oxidative insults, accumulation of intracellular GSSG leads to increased GSSG efflux which in turn, can directly activate NMDA receptors involved in subsequent seizure activity. xi Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. CHAPTER ONE Introduction to Dissertation Research and Literature Review INTRODUCTION TO THE DISSERTATION RESEARCH Kainic acid (KA) has been shown to induce seizures and neuronal cell death. However, the mechanisms by which KA induces neuronal cell death are not completely understood. Reactive oxygen species (ROS) has been proposed to play an important role in KA-induced neuronal cell death. The original aim of this dissertation research was to investigate the role of oxidative stress in KA excitotoxicity. Since cytosolic glutathione peroxidase (GSHPx-1) can remove hydrogen peroxide (H2O 2), one of the ROS that appear to be involved in KA toxicity, theoretically overexpression of the enzyme should attenuate the toxicity. In this study, murine GSHPx-1 transgenic mice, exhibited 4-6 fold increases in the enzyme activity, primary neuronal cultures and acute brain slices were used to determine the role of oxidative stress, particularly H2O2 , in KA-induced neuronal cell death. 1 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. LITERATURE REVIEW 1. Free radicals, reactive oxygen species (ROS), reactive nitrogen species (RNS), antioxidants and oxidative stress. 1.1. Terminology. Free radicals, defined as “any species capable of independent existence that contains one or more unpaired electrons in an atomic or molecular orbital” (Halliwell, 1996), are highly reactive with other molecules. To become more stable, these free radicals remove electrons from nearby molecules. This can lead to irreversible damage of biological molecules including DNA, RNA, proteins and lipids. ROS is defined as any reactive molecules associated with oxygen atoms or oxygen equivalents, including not only oxygen-containing free radicals such as superoxide radical anion (02*"), hydroxyl radical (OH*), peroxyl radical (ROO*) but also other non-free radical reactive molecules such as H2O 2 and hypochlorous acid (H0C1) (Halliwell, 1996). RNS refers to reactive nitrogen-containing molecules such as nitric oxide (NO*), peroxynitrite anion (ONOO ) and nitroxyl anion (NO ) (Ohshima, 2003). Antioxidants are defense systems involved in prevention, interception and/or repair of damage caused by ROS and/or RNS. Antioxidants include non-enzymatic scavengers and quenchers such as tocopherols (vitamin E), ascorbate (vitamin C) and 2 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. carotenoids, and enzymatic systems such as superoxide dismutase (SOD), catalase and glutathione peroxidase (GSHPx) (Sies, 1991; Halliwell and Gutteridge, 1999). Oxidative stress is defined as an imbalance between oxidants and antioxidants potentially leading to damage (Sies, 1991). 1.2. Sources of ROS and RNS. 1.2.1. Exogenous sources. Exposure to radiation is known to cause free radical production. For example, gamma ray exposure induces OH* and 02*" production (Sarma et al., 1996). Protein oxidation is also known to occur during exposure of protein to ultraviolet (UV) or visible light via 02*’ production (Davies, 2003). UV light also can cleave the covalent bond in water, generating OH* (Halliwell, 1996). 1.2.2. Endogenous sources. There are four prominent sources of oxidants produced in the cells (Ames et ah, 1993). (i) The electron transport chain: Most organisms need oxygen for energy generation as the terminal electron acceptor in the electron transport chain in the mitochondria. However an error in electron transfer at any point in the electron transport chain can result in an electron being accepted by oxygen leading to 02*’ formation and eventually OH* production in the presence of metal ions such as Fe2 + and Cu+. Under physiological conditions, approximately 2-3% of the oxygen consumed by aerobic cells is converted into 02*" and other ROS (Sohal and Weindruch, 1996). In each rat cell, 2 x 101 0 02*" and H 2O2 3 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. are generated daily (Ames et al., 1993). Roughly 2 kg of 02*', is generated in a human body every year (Halliwell, 1996). Each day, in a rat cell, DNA suffers about 100,000 oxidative attacks, 10,000 attacks in a human cell (Ames et al., 1993). (ii) Phagocytic activity: to rid the cell of bacteria and virus, phagocytic cells use an oxidative burst of NO*, 02*' and H 2O2 . Chronic infection of bacteria, virus or parasites causes a continual phagocytic activity leading to chronic inflammation, increasing the risk of cancer, (iii) Peroxisomes: H 2O2 is generated as a by-product of fatty acids and other molecular degradation which take place in peroxisomes. Under some conditions, H 2O 2 escapes from being degraded by catalase within the peroxisome, resulting in its leakage into other compartments of the cell and subsequent oxidative damage, (iv) Cytochrome P450 enzyme: This enzyme is one of the primary defense systems against toxins. During the process of preventing the acute effects of toxins, oxidant by-products are generated, leading to oxidative damage (Ames et al., 1993). 1.3. ROS There are several ROS generated and involved in oxidative damage as shown in Table 1 (Sies, 1991). Three major ROS: 02*", OH* and H 2O 2 are described in this section. 1.3.1. Superoxide radical. 02*', a weakly reactive radical, is generated by several pathways (Figure 1); (i) oxidation by nicotinamide adenine dinucleotide phosphate (NADPH) oxidase; (ii) oxidation of xanthine or 4 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. hypoxanthine by xanthine oxidase; (iii) reducing equivalents (e.g. reduced nicotinamide adenine dinucleotide (NADH), NADPH and reduced flavin adenine dinucleotide (FADH 2)); (iv) autoxidation of monoamines (e.g. dopamine, epinephrine and norepinephrine), flavins and hemoglobin in the presence of metals; (v) one-electron reduction of O2 by cytochrome P-450; and (vi) one-electron reduction of O2 by nitric oxide synthase (NOS) (Fang et al., 2002). Compound Remarks O 2V superoxide anion One-electron reduction state, generated in several autoxidation reactions. H 0 2 *, perhydroxyl radical Protonated form of 02*", more lipid soluble H 2O 2 , hydrogen peroxide Two-electron reduction state, generated by dismutation from 0 2 *’ or directly from O 2 OH*, hydroxyl radical Three-electron reduction state, generated by Fenton reaction, Haber-Weiss reaction, highly reactive RO*, alkoxy radical Oxygen-centered organic radical ROOH, hydroperoxide Organic hydroperoxide ROO*, peroxy radical Generated from ROOH by hydrogen abstraction JA g 0 2 or '0 2 Singlet molecule oxygen, first excited state, 22 kcal/mol above ground state (triplet) 3 0 2 Table 1. ROS of interest in oxidative stress (Sies, 1991). 1.3.2 Hydrogen peroxide. H2O 2 is produced through dismutation of 02*" by SOD. Two-electron reduction of O2 by cytochrome P-450, D-amino acid 5 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. oxidase, acetyl coenzyme A oxidase, or uric acid oxidase also generates H 2O 2 . In the pathway of glycine metabolism, the oxidation of sarcosine also leads to H 2O2 generation (Fang et al., 2002) (Figure 1). H 2O2 is poorly reactive but at high concentration can cause some cellular damage (Hyslop et al., 1988). NADH + H+ (or FADH2 ) NAD+ (or FAD) NADPH Oxidase T V " NADPH+ H+ NADP+ HOC1 - MP0 (X H+ + Cl' Xanthine Xanthine oxidase Uric acid P-450 ETSJ 1 h 2 o 2 Oxidized Monoamines and Hemoglobin o 2 + h 2 o Vit C Vit C o • J t Autoxi dation Fe2+ or Cu+ Figure 1. Pathways of ROS generation (Fang et al., 2002). ETS, electron transport system; FAD, oxidized flavin adenine dinucleotide; FADH 2 , reduced flavin adenine dinucleotide; H2O2 , hydrogen peroxide; IR, ionizing radiation; MPO, myeloperoxidase; NAD+, oxidized nicotinamide adenine dinucleotide; NADH, reduced nicotinamide adenine dinucleotide; NADP+, oxidized nicotinamide adenine dinucleotide phosphate; NADPH, reduced nicotinamide adenine dinucleotide phosphate; NOS, nitric oxide synthase; O2V superoxide anion radical; OH», hydroxyl radical; P-450, cytochrome P-450; SOD, superoxide dismutase; Vit C, vitamin C. 1.3.3. Hydroxyl radical. OH* is extremely reactive. It reacts with a broad range of both organic and inorganic molecules including DNA, lipid and 6 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. proteins (Cheng et al., 2002). Interactions between 02*' and H 2O2 can generate OH* through the metal-dependent Fenton reaction (reaction 2) and the iron-catalyzed Harber-Weiss reaction (reaction 3) (Kehrer, 2000). Fe3 + + 02*’ Fe2 + + 0 2 reaction (1) Fe2 + + H2 0 2 Fe3 + + OH+OH« ............. reaction (2) The net reaction: 0 2 » ~ + H2 0 2 -> 0 2 + OH' + OH» ................reaction (3) 1.4. ROS and DNA modification. ROS can induce several forms of DNA modification such as oxidation of bases, abasic sites and DNA strand breaks (Klungland et al., 1999). The guanine base in DNA is highly sensitive to oxidative attack. Thus its modifications are used as markers for oxidative-mediated DNA damage (Kehrer, 2000). More than 20 oxidative-mediated modifications have been identified (Henle et al., 1996). Levels as low as 25 x 10'1 5 mol of 8-hydroxy-2’-deoxyguanosine (8-OHdG), one of the oxidative-mediated guanine modifications, can be detected (Kehrer, 2000). It has been estimated that, under steady state conditions, there are 100 8-OHdG residues per genome (Klungland et al., 1999). Even though, a number of repairing systems existing in the cell that are responsible of repairing such damage (Lindahl and Wood, 1999), if the DNA lesions occur at important sites or are left unrepaired, these oxidized DNA can cause several adverse effects such as an increase in genetic mutations and transcriptional errors (Kehrer, 2000). 7 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 1.5. ROS and protein modification. ROS can generate several forms of oxidized amino acids and proteins such as 2-oxohistidine, glutamic acid hydroperoxide and protein carbonyls (Dean et al., 1997). The oxidation of amino acids and proteins can lead to protein malfunctions. However, no protein repair systems are present because it is more efficient either to prevent the oxidation of proteins via antioxidant systems to begin with or to remove the damaged protein through proteolysis. Accumulations of unremoved oxidized proteins appears to play a role in a variety of diseases such as diabetes and several neurodegenerative diseases (Kehrer, 2000). 1.6. ROS and lipid modification. Alterations in lipids can lead to cell death due to the failure to maintain membrane structures and functions. The double bonds in polyunsaturated fatty acids are sensitive to oxidative attack. Once ROS removes a hydrogen atom from a lipid double bond, this generates a new radical species that can react with oxygen leading to formation of a lipid peroxyl radical. The lipid peroxyl radical can initiate a chain reaction by removing a hydrogen atom from another fatty acid, generating another lipid peroxyl radical and a lipid hydroperoxide which is unstable and eventually converted to a variety of molecules such as malodialdehyde. These processes can cause alterations of ion channels, inactivation of membrane transport proteins and enzymes, and permeabilization of the lipid bilayer leading to disruption of ion 8 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. homeostasis. Some oxidized fatty acids are also able to affect apoptotic signaling pathways (Kehrer, 2000). 1.7. ROS and apoptosis. Apoptosis is a form of cell death. Literally, apoptosis, derived from the Greek, means “the falling off of leaves from a tree, or the falling off of petals from a flower” (Takahashi et al., 2004). Characteristically, an apoptotic cell shrinks remarkably. Its plasma membrane becomes ruffled and blebbed. The cytoplasm becomes condensed due to loss of water and ions within intact organelles. The cell breaks up into small sealed compartments, called apoptotic bodies. There is no intracellular leakage. Its chromatin becomes very densely associated and fragmented (Cohen, 1993). Apoptosis can be triggered via several genetic pathways including those which are either caspase-dependent or caspase-independent. Several caspase- dependent pathways have been identified; the Fas-associated death domain protein (FADD)/caspase-8 pathway, the mitochondrial/caspase-9 pathway and the endoplasmic reticulum/caspase-12 pathway. Caspase-independent pathways such as BAX, a Bel-family protein, can activate other proteases (reviewed in Assuncao Guimaraes and Linden, 2004) (BAX activation is also part of caspase-dependent pathways). Generation of ROS, such as H 2O 2 , is also known to induce apoptotic cell death which can be inhibited by antioxidants such as N-acetylcysteine (NAC). Since the major endogenous source of ROS production is the mitochondria, an inhibition of 9 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. its ROS production by a mitochondrial complex I inhibitor, rotenone, can suppress ceramide-induced apoptosis. Contrarily, apoptosis can be potentiated by antimycin A, a complex III inhibitor which blocks electron flow downstream of ubiquinone (Coenzyme Q10) and induces mitochondria ROS production (Quillet-Mary et al., 1997). 1.8. Antioxidants in the brain. There are several antioxidant systems present in the brain, for example; (i) antioxidant enzymes such as SOD and GSHPx; (ii) GSH and cysteine; (iii) vitamin C and E (Halliwell and Gutteddge, 1999). In the next section, GSHPx will be described extensively. 2. Glutathione peroxidase (GSHPx). Glutathione peroxidase (GSHPx, glutathione:hydrogen-peroxide oxidoreductase, also known as selenium (Se)-glutathione peroxidase and reduced glutathione peroxidase) is found ubiquitously in mammals and it is best known for its role in the elimination of ROS such as H2O2 and ONOO\ 2.1. History. GSHPx was first described in 1957 by Mills (Mills, 1957) as an enzyme isolated from erythrocytes, in the presence of GSH, which prevented the H 2O 2- induced oxidation of hemoglobin. Since the enzyme had a peroxidatic activity with 10 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. GSH serving as a hydrogen donor, the enzyme was called glutathione peroxidase, suggesting its physiological role in protecting erythrocytes from oxidative stress by catalyzing the oxidation of GSH by H 2O2 , resulting in water and oxidized glutathione (GSSG, also known as glutathione disulfide), as represented by reaction (4); H 2O2 + 2GSH----------------► 2H2 0 + GSSG ......... reaction (4) GSHPx GSSG + NADPH + H+ ----------------► 2GSH + NADP+ ........reaction (5) GR However, Mills and Randall (Mills, 1957; Mills and Randall, 1958) suggested that, to maintain the physiological role, a continual supply of GSH is required. GSH reductase (GR, EC 1.8.1.7, also known as GSSG reductase) was shown to be the enzyme that catalyzes the reduction of GSSG in the presence of reduced nicotinamide adenine dinucleotide phosphate (NADPH, previously known as reduced triphosphopyridine nucleotide or TPNH) which serves as a coenzyme resulting in GSH and nicotinamide adenine nucleotide phosphate (NADP+, previously known as triphosphopyridine nucleotide or TPN+ ) as represented by reaction (5) (Rail and Lehninger, 1952). Glucose-6-phosphate dehydrogenase (G6PD) and 6-phosphogluconate dehydrogenase (6PGD), rate-limiting enzymes in the pentose phosphate pathway (PPP) which coverts glucose into pentose sugars, are also required to generate NADPH in order to maintain a sufficient amount of GSH through the reduction of GSSG by GR (Cohen and Hochstein, 1963; Salvemini et al., 11 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 1999; Mehta et al., 2000; Grabowska and Chelstowska, 2003; Leopold et al., 2003; Spolarics et al., 2004) (Figure 2). G_6_P\ G6PD ) NADP+ ) GR 2 GSH ) GSHPx ( Ru-5-P / 6PGD + co2 NADPH GSSG 2H 2 0 Figure 2. Schematic of the GSH metabolic pathway. Glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase generate NADPH which is required to maintain an adequate amount of GSH, a substrate of H 2C > 2-scavenging GSHPx (Mehta et al., 2000). G-6-P, glucose-6-phosphate; Ru-5-P, ribulose-5- phosphate; G6PD, glucose-6-phosphate dehydrogenase; 6PGD, 6-phosphogluconate dehydrogenase; GR, glutathione reductase. 2.2. GSHPx isoforms. Several selenocysteine-containing enzymes in the glutathione peroxidase family have been identified including cytosolic GSHPx (GSHPx-1), gastrointestinal GSHPx (GSHPx-GI or GSHPx-2), glycosylated plasma GHSPx (GSHPx-P or GSHPx-3), phospholipid hydroperoxide GSHPx (PHGSHPx or GSHPx-4) and sperm nuclei GSHPx (snGSHPx) (Armstrong et al., 1998). cGSHPx, EC 1.11.1.9). GSHPx-1 was first identified as a Se-dependent enzyme by Rotruck and coworkers (Rotruck et al., 1973) since hemolyzates prepared from erythrocytes of Se-deficient rats did not protect hemoglobin from H202-induced oxidative stress in the presence of GSH. To prove that GSHPx-1 contains Se, rats 2.2.1. Cytosolic GSHPx (GSHPx-1 or classical GSHPx or 12 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 'JC were injected with Se as sodium selenite. 2-4 weeks later, GSHPx-1 was purified from hemoglobin obtained from the Se injected rats by using DEAE Sephadex chromatography. 60% of 75Se was detected in the fractions that contained GSHPx activity. The fractions were then concentrated and purified using Sephadex G-150. 70% of the applied 75Se was detected in the peaks of molecular weight 90 kDa which contained GSHPx activity, and 25 kDa which contained a significant level of GSHPx activity. By using neutron activation analysis technique on GSHPx-1 prepared from bovine blood, Flohe and coworkers described GSHPx-1 as a selenoenzyme since it contains Se but no heme, nor flavin, assumingly, Se is the active part of the enzyme (Flohe et al., 1973). The finding of GSHPx-1 as a selenoenzyme explains many Se deficiency related pathological conditions that involve oxidative stress, for example Phrabhu and colleagues reported a 17-fold decrease in GSHPx-1 level and increased oxidative stress in Se-deficient RAW 264.7 cells, a murine macrophage cell line. Lipopolysaccharide (LPS) treated Se-deficient RAW 264.7 cells show increased levels of expression of the inducible isoform of NOS (iNOS) and NO* production (Prabhu et al., 2002). Forstorm and colleagues were the first group to demonstrate that the Se in GSHPx-1 was in the form of selenocysteine, the catalytic site of the enzyme (Forstrom et al., 1978). 2.2.2. Gastrointestinal GSHPx (GSHPx-GI or GSHPx-2). GSHPx - 2 was characterized by Chu and coworkers as a Se-dependent tetrameric protein 13 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. present in cytoplasm (Chu et al., 1993). In human, GSHPx-GI expression was detectable in liver, colon, stomach, small intestine and, occasionally, breast but not in heart or kidney. In rats, GSHPx-3, expressed in intestinal epithelium, is the major cellular GSHPx in the gastrointestinal tract (Chu and Esworthy, 1995). 2.2.3. Glycosylated plasma GSHPx (GSHPx-P or GSHPx-3). GSHPx-P was purified and described by Takahashi and colleagues as a 100-kD tetrameric glycoprotein consisting of 23 kD subunits (Takahashi et al., 1987). It contains 4 atoms of Se per mole. Avissar and coworkers demonstrated that, even though endothelial cells and myeloid cells are known to secrete and release plasma proteins, only hepatic cells synthesize and secrete this enzyme (Avissar et al., 1989). GSHPx-P is found in plasma (Takahashi et al., 1987), milk (Avissar et al., 1991), kidney and lung (Yoshimura et al., 1991). 2.2.4. Phospholipid hydroperoxide GSHPx (PHGSHPx or GSHPx-4). PHGSHPx (EC 1.11.1.12) was discovered by Ursini and colleagues (Ursini et al., 1982). This monomeric GSHPx is found in spermatids and testis and play an important role in spermatogenesis (Roveri et al., 1992). It has been shown to be able to reduce phospholipids, hydroperoxides and cholesterol hydroperoxides (Thomas et al., 1990). It was also been reported to play a protective role as an anti- apoptotic in hydroperoxide-mediated mitochondrial death pathways (Nomura et al., 1999). 14 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2.2.5. Sperm nuclei GSHPx (snGSHPx). The latest Se-dependent GSHPx, snGSHPx found in testis, was recently identified by Pfeifer and coworkers (Pfeifer et al., 2001a; Pfeifer et al., 2001b) as a specific sperm nucleic specific enzyme. Though it differs from PHGSHPx at its N-terminal, its properties are similar to those of PHGSHPx. In rats, snGSHPx, the only selenoenzyme present in the nuclei of late spermatids, plays important roles in chromatin condensation and as an anti-oxidant agent which are necessary for spermatogenesis and male fertility (Pfeifer et al., 2001a; Pfeifer et al., 2001b). 2.3. Genetically altered GSHPx-1 models. Several genetically engineered GSHPx-1 models both overexpressing and null mutations, have been generated. These models have been used by several laboratories to study G SH Px-l’s protective capability against a range of oxidative insults. 2.3.1. Overexpression of GSHPx. Kelner and colleagues investigated whether overexpression of human GSHPx-1 in human MCF7 and murine NIH3T3 cancer cells, transfected with human GSHPx-containing vectors, would protect the cells from oxidative insults (Kelner et al., 1995). With 2-4 fold increases in GSHPx activity levels, the cancer cells were resistant to /-butyl hydroperoxide, H 2O2 and lipid hydroperoxide-mediated paraquat toxicities. This provides evidence of the enzyme’s ability to remove other peroxides besides H 2O 2. 15 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Mirochnitchenko and his group created transgenic mice expressing human GSHPx-1 and GSHPx-P driven by the mouse hydroxy-methylglutaryl-coenzyme A reductase gene promoter (Mirochnitchenko et al., 1995). In GSHPx-1 transgenic mice, a 400% increase of GSHPx activity was found in the brain, 160% in muscle, 127% in liver and 150% in kidney compared to those of control animals. In GSHPx - P transgenic mice, a 55% increase of GSHPx activity was found in blood compared to control animals. Hyperthermia was induced in both of the transgenic lines and non-transgenic controls. Both transgenic lines showed increased sensitivity to the hyperthermia having shorter survival times during exposure to elevated temperatures than control animals. Overexpression of GSHPx-1 or GSHPx-P, resulting in lower levels of H 2O2 and lipid peroxides which are effective inducers of heat shock protein 70 (HSP70), leading to lowered level of HSP70 activation and increased sensitivity to heat-induced hyperthermia. Though, no significant differences in GSH or GSSG levels were detected in transgenic vs. wild-type animals after heat-induced stress, the effect of local imbalance in the GSH/GSSG ratio can not be eliminated. Spector and colleagues created a GSHPx-1 transgenic mouse line using the 5.3 kb mouse GSHPx-1 genomic DNA consisting of 1.4 kb of the gene, 2.2 kb of 5’ flanking region and 1.7’ kb of 3’ flanking region (Spector et al., 1996). The activities of GSHPx-1 in the lens, brain, heart and liver of transgenic animals were 4.5, 3.4, 2.4 and 1.2 times higher than those of wild-type animals, respectively. To study the protective activity of GSHPx-1 against H202-induced oxidative stress, lens epithelial cells prepared from GSHPx-1 transgenic animals were treated with H2O2 . 16 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Surprisingly, the degradation of H 2O 2 rate in cells prepared from GSHPx-1 transgenic animals was not significantly different from those prepared from wild- type animals or GSHPx-1 null mutation animals. A possible explanation for increased GSHPx-1 activity not being efficient in H 2O2 degradation is that the system was unable to provide or maintain a high enough level of GSH, the substrate of the enzyme. This is probably due to the imbalance of GSHPx-1 and GR activity ratio; in the whole lens, 11.7 in wild-type versus 51 in GSHPx-1 transgenic animals; 1.5 in lens epithelial cells prepared from wild-type animals versus 12 in those prepared from GSHPx-1 transgenic animals. Yoshida and colleagues (Yoshida et al., 1996) used the same line of GSHPx - 1 transgenic mice as Spector and co-workers (Spector et al., 1996) to test the hypothesis that the cellular antioxidant enzyme could protect the animals against oxidative stress mediated myocardial ischemia/reperfusion injury. A 30-munite ischemia followed by a 20-minute reperfusion was introduced to the hearts of GSHPx-1 transgenic animals. Transgenic mouse hearts exhibited less ischemia/reperfusion mediated injury as assessed by creatine kinase release and myocardial infarct size, and faster post-ischemic ventricular recovery rate compared to wild-type hearts. Wiesbort-Lefkowitz and co-workers used another line of GSHPx-1 transgenic mice to demonstrate the antioxidant enzyme’s protective capacity against focal cerebral ischemia/reperfusion (Weisbrot-Lefkowitz et al., 1998). Ischemia/reperfusion was achieved in the brains of human GSHPx-1 transgenic mice 17 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. by a one-hour middle cerebral artery occlusion followed by 24-hour reperfusion. Before the brains were removed for assessments of infarct size, hemispheric enlargement and histological damages, neurological deficits were evaluated. Four fold overexpression of human GSHPx-1 was shown to attenuate oxidative stress- mediated ischemia/reperfusion damage, as assessed by reduced behavioral deficits, smaller infarct sizes, less brain edema and less histological damage compared to wild-type mice. This suggested that GSHPx might be a potential treatment for stroke. Mirochnitchenko and his group investigated how transgenic mice overexpressing human GSHPx-1 and GSHPx-P responded to acute acetaminophen overdose which known to cause fatality (Mirochnitchenko et al., 1999). Overexpression of GSHPx-P appeared to be protective against the toxicity of acetaminophen intraperitoneal (IP) injection; the mortality rate following exposure was 25% compared to 75% in acetaminophen treated non-transgenic animals. Surprisingly, overexpression of GSHPx-1 showed an opposite effect in acetaminophen IP injected GSHPx-1 transgenic mice; all animals died within 5.5 hours of treatment compared to 75% mortality rate in non-transgenic animals over a period of 36 hours after treatment. Injections of GSHPx into tail veins of non- transgenic mice appeared to be protective against the acetaminophen toxicity. This indicated that increased GSHPx activity in the blood, either genetically or injectionally, protected the animals against the toxicity of acetaminophen. Interestingly, acetaminophen-treated GSHPx-1 transgenics had lower GSH levels at 8 hours after treatment compared to GSHPx-P and non-transgenic mice in which 18 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. GSH levels had returned to baseline. The researchers hypothesized that GSHPx-1 transgenic mice were unable to regenerate GSH efficiently, resulting in an increased susceptibility of the animals to acetaminophen toxicity. Lu and colleagues reported that overexpression of GSHPx-1 or GSHPx-1 and SOD increased carcinogenic response to a series of 7,12-dimethylbenz[a]anthracene (DMBA) and 12-0-tetradecanoylphorbol-13-acetate (TPA) treatments (Lu et al., 1997). After the treatments, transgenic animals, both GSHPx-1 alone and GSHPx-1 and SOD, developed 10.9 and 11 tumors per animal, respectively, compared to 3.9 tumors per animal in non-transgenic animals. At 10 weeks after the treatments had stopped, 9-11% of the tumor-bearing transgenic mice showed complete degeneration of tumors compared to 26% in tumor-bearing non-transgenic mice. Mirochnitchenko and colleagues investigated the role of GSHPx in response to lipopolysaccharide(LPS)-induced endotoxemia by using transgenic mice overexpressing human GSHPx-1 and GSHPx-P (Mirochnitchenko et al., 2000). Both strains of the transgenic animals, injected with a lethal dosage of LPS, showed higher survival rates and blood pressure compared to those of LPS-injected non-transgenic animals. At 8 and 18 hours following LPS injection, GSHPx activity levels in several organs, including heart, kidney, liver and platelets, were not altered by LPS treatments in both transgenic and non-transgenic mice. However, at 18 hours, GSHPx activity levels in lungs of GSHPx-1 and GSHPx-P transgenic animals were slightly lower than those before LPS treatment. LPS-induced NO* production was detected in all animals. Nitrite/nitrate levels in transgenic animals were 30-40% less 19 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. than those in non-transgenic animals. iNOS protein levels and its activity levels in macrophages from non-transgenic mice were higher compared to those from transgenic animals. Lipid peroxidation levels in liver and kidney from LPS-treated transgenic mice were lower than those from LPS-treated non-transgenic mice. In 2002, Gouaze and colleagues demonstrated that overexpression of human GSHPx-1 in the human breast cell line T47D, inhibited CD95-induced apoptosis via inhibition of effector caspase activation and ROS production (Gouaze et al., 2002). Moreover, in an experiment in human GSHPx-1 transgenic mice injected with anti- CD95 antibody, the animals were more resistant to anti-CD95 triggered cell death. Wang and colleagues reported decreased susceptibility to oxygen/glucose deprivation (OGD) in the hippocampal via herpes simplex virus-1 mediated overexpression of human GSHPx-1. An 18% increase of GSHPx activity in the infected hippocampal cultures was found to be neuroprotective, resulting in a 33% decrease in susceptibility to OGD. Infected cortical cultures, with more than a 30% increase of GSHPx activity, were found to be more resistant (80%) to the toxicity of 50 |iM KA compared to the controls, empty vector infected cortical cultures. The increase of GSHPx activity in cortical cultures was also found to reduce by approximately 60% of the glutamate-induced hydrogen peroxide accumulation and completely block the toxicity of 50 ftM glutamate and glutamate-induced lipid peroxidation (Wang et al., 2003). Shiomi and coworkers used overexpressing human GSHPx-1 transgenic mice to demonstrate that overexpression of the enzyme protects the heart against oxidative 20 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. stress from left ventricle structural remodeling and functional failure after myocardial infarction (MI) (Shiomi et al., 2004). MI was introduced surgically to the animals. 4 weeks later, infarct size and myocardial histopathology were performed. Antioxidant enzyme activities, GSHPx, catalase and SOD, and lipid peroxidation in myocardial tissues with MI were also determined. Only increased GSHPx activity levels, but not catalase nor SOD, in noninfarcted left ventricle obtained from transgenic mice with MI were detected. Lipid peroxidation levels in noninfarcted left ventricle obtained from transgenic mice with MI were significantly lower compared to those of wild-type with MI. An increased survival rate but unaltered infarct size was detected in transgenic mice with MI. Cell death in the noninfarcted left ventricle of transgenic mice with MI, assessed by Terminal Deoxynucleotide Transferase- mediated dUTP Nick-End Labeling (TUNEL) staining, was significantly lower compared to those of wild-type with MI. These suggest that overexpression of GSHPx can protect the heart from ROS-mediated left ventricular remodeling and failure after MI. Liu and colleagues reported that adenovirus-mediated overexpression of human GSHPx-1 (AdGSHPx) in human primary pancreatic undifferentiated adenocarcinoma cells of ductal epithelial cell origin (MIA PaCa-2 cells) showed slower in vitro growth when compared to parent cells (Liu et al., 2004). AdGSHPx also lowered plating efficiency and growth in soft agar of the tumor. In addition, in in vivo studies, AdGSHPx slowed growth of MIA PaCa-2 induced tumors in nude mice. 21 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2.3.2. GSHPx-1 Knockout. Taylor and co-workers (Taylor et al., 1993) demonstrated in DG44 cells, a Chinese hamster ovary cell line, transfected with an antisense bovine GSHPx expression vector resulting in an 80% decrease of GSHPx activity, that the reduced GSHPx activity caused the infected cells to become more sensitive to oxidative-mediated paraquat and adriamycin toxicities, assessed by a viability assay. This was also confirmed by a treatment of the cell line with buthionine sulfoximine (BSO), which reduces GSH substrate. Ho and colleagues created a null GSHPx-1 deficient mouse line (knockout), containing an interruption due to presence of the neomycin resistance gene (Ho et al., 1997). The expression of GSHPx-1 (GSHPx-1 mRNA) and GSHPx activities in the brain, heart, kidney, liver and lung of GSHPx-1 heterozygous knockout animals were found to be about 40-60% of those of wild-type animals where, in GSHPx-1 homozygous knockout animals, GSHPx-1 mRNA was not found in those tissues. Although very low GSHPx activity can be detected in the GSHPx-1 homozygous knockout, this may be due to the expression of other GSHPx isoforms. Both GSHPx- 1 heterozygous and homozygous knockout animals appeared healthy compared to wild-type animals. The numbers of erythrocytes, reticulocytes and differential leukocytes of the knockouts were comparable to those of wild-type animals. No significant difference in other antioxidant enzymes activities, protein oxidation and lipid peroxidation were detected in the knockout animals. When lung tissues prepared from GSHPx-1 homozygous knockout animals were exposed to hyperoxia, 22 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. there was no significant difference in median survival times, protein carbonyl content and lipid peroxidation compared to those of wild-type animals. This suggested a very limited role of the antioxidant enzyme in animals under normal physiological conditions and in pulmonary defense against hyperoxia. De Haan and coworkers reported that GSHPx-1 null mutation (knockout) mice showed an increased sensitivity to oxidative-mediated paraquat toxicity (de Haan et al., 1998). Non-detectable to very low levels of GSHPx activity, less than 4% of wild-type animals, was measured in liver, lung and brain of homozygous knockout mice with an exception of a 32% residual activity in the heart, presumably, due to other GSHPx isoforms. Knockout animals were healthy compared to wild- type animals. A 100% mortality was observed in homozygous knockout animals within 5 hours of paraquat treatment, at a dosage of 30 mg/kg body weight, approximately a half of the LD5 0 of wild-type animals. Up to 10 days after paraquat treatment at the same dosage, no lethality was observed in wild-type animals. At a smaller dosage of paraquat, 20 mg/kg body weight, a 76% mortality was observed between 6 and 14 hours after the treatment in knockout animals. Only 1 out of 11 knockout animals died at 15 hours after a 10-mg/kg body weight treatment of paraquat. This suggested a dose-dependent sensitivity to oxidative-mediated paraquat toxicity in the knockout animals. In addition, paraquat has been shown to up-regulate GSHPx-1 expression in the human hepatocytes cell line, HepG2, and Chinese hamster ovary cell line, CHO, transfected with human GSHPx-1 promoter/1 uciferase vector. This provided evidence supporting a role for GSHPx-1 in protecting against 23 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. oxidative-mediated paraquat toxicity. Neuronal primary cultures prepared from GSHPx-1 knockout embryos were used to study the effect of another oxidant, H 2O2 . Cortical neurons cultures were treated with various concentrations of H 2O 2 . 70% of knockout cultures survived a 65-pM treatment of H 2O 2 versus a 100% viability in wild-type cultures. At higher concentrations of H2 O2 , 80 and 100 pM, only approximately one third and none of knockout cultures survived versus 66% and 25% viabilities observed in wild-type cultures, respectively. This suggested an antioxidant role for GSHPx in H 2O 2 toxicity. Fu and colleagues investigate roles of GSHPx-1 in apoptosis induced by diquat (DQ), a ROS (i.e. 02*" generator, and peroxynitrite (PN), an RNS generator (Fu et al., 2001). Primary hepatocytes isolated from GSHPx-1 knockout and wild- type mice were treated with either 0.5 mM DQ or 0.1-0.8 mM PN for 3, 6, 9 or 12 hours. GSHPx-1 knockout hepatocytes were more susceptible to DQ toxicity but yet interestingly, more resistant to PN toxicity compared to wild-type hepatocytes. DNA fragmentation, TUNEL-positive apoptotic cells, cytochrome-c release and caspase-3 activation were detected in DQ-treated GSHPx-1 knockout and PN-treated wild-type hepatocytes but not in untreated, DQ-treated wild-type and PN-treated GSHPx-1 knockout hepatocytes. GSH levels in DQ-treated and PN-treated wild-type hepatocytes were lower than in treated GSHPx-1 knockout cells. DQ induced increases of GSSG levels in wild-type hepatocytes but not in GSHPx-1 knockout cells. PN did not have effects on GSSG levels in GSHPx-1 knockout hepatocytes but altered GSSG levels of wild-type cells during the first 30 minutes after PN 24 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. treatments. GSH/GSSG ratios were higher in treated GSHPx-1 knockout hepatocytes than in treated wild-type cells. Protein nitration was detected in PN-treated GSHPx-1 knockout and PN-treated wild-type cells but not in DQ-treated cells. Jiang and colleagues demonstrated that GSHPx-1 knockout mice were more resistant to KA-induced epileptic seizures and neurodegeneration compared to wild- type animals (Jiang et al., 2000). However, KA-treated knockouts had higher levels of oxidative stress indices in the brain including ROS, lipid peroxidation and carbonyl contents. Following KA treatment, increased GSSG levels were observed in the brains from wild-type animals, however, no significant increase of GSSG levels were observed in the brains of the GSHPx-1 knockouts. The resistance to KA- induced seizure was found to be due to oxidation of the NR1 subunit of N-methyl-D- aspartate (NMDA) receptor leading a reduction in its function. 3. Kainic acid (KA). 3.1. KA-induced seizures and neuronal cell death. KA, a potent excitotoxin, is an agonist of non-NMDA receptors (a-amino-3- (3-hydroxy-5-methylisoxazol-4-yl) propionic acid (AMPA)/KA receptors). KA, originally isolated from the sea weed Digenea simplex, once was used to treat intestinal worms, ascarides, in Japan (Sperk, 1994) Since it was first described as a neurotoxin in 1974 by Olney and co workers (Olney et al., 1974) it has been used as an effective tool for exploring excitatory amino acid transmission in both invertebrates and mammals (Sperk, 1994) With its excitotoxic properties, KA 25 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. induces both acute and sub-acute epileptical seizure activity both via intracerebral injection of KA, which also generates both local and remote seizure-related lesions in several brain areas (Ben-Ari et al., 1979; Ben-Ari et al., 1980a; Ben-Ari et al., 1980b; French et al., 1982; Pollard et al., 1994), and systemic administration of KA, which also generates morphological changes similar to the former (Ben-Ari et al., 1981). Since KA cannot cross the blood-brain-barrier very well, only a few percent of the systematically administered KA can reach its receptors in the brain. This might contribute to the differences in susceptibility to KA of various animal strains (Sperk, 1994). Although KA receptors are present in other portions of the CNS, most KA receptors can be found in the CA3 region of the hippocampus (Foster et al., 1981; Ozawa et al., 1998). This causes the CA3 region to be particularly vulnerable to KA toxicity (Pollard et al., 1994; Ben-Ari and Cossart, 2000). KA not only generates seizure and neuronal degeneration in the CA3, but it also induces lesser degeneration in other areas including the CA1, CA4 and amygdala (French et al., 1982; Pollard et al., 1994; Venero et al., 1999; Ben-Ari and Cossart, 2000) and to even a lesser extent in the CA2 and dentate gyrus (Sperk et al., 1983). The clinical signs generated by KA administration can be divided into several phases; staring, head nodding, wet-dog shaking, individual recurrent limbic motor seizures, forepaws tremor, rearing and loss of postural control. Then the seizures become more complex and prolonged with shorter pauses. Finally, the animals develop a full status epilipticus; continuous convulsions (Ben-Ari, 1985). 26 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3.2. KA toxicity and oxidative stress. There are several pieces of evidence suggesting that oxidative stress plays a role in KA excitotoxicity (Coyle and Puttfarcken, 1993; Bruce and Baudry, 1995; MacGregor et al., 1996; Chen and Chuang, 1999; Carrasco et al., 2000; Erakovic et al., 2000; Gluck et al., 2000; Liang et al., 2000; Patel et al., 2001; Patel and Li, 2003). KA administration has been found to produce increased oxidative stress marker levels such as lipid peroxidation (Bruce and Baudry, 1995; Carrasco et al., 2000; Gluck et al., 2000; Patel et al., 2001), protein oxidation (Bruce and Baudry, 1995; Carrasco et al., 2000; Gluck et al., 2000), DNA oxidation (Liang et al., 2000; Patel and Li, 2003); increased antioxidant enzymes activity levels such as SOD (Bruce and Baudry, 1995; Liang et al., 2000) and catalase (Bruce and Baudry, 1995); and an alteration in cellular redox system i.e. decreased GSH levels which lead to an decreased GSH/GSSG ratio (Chen and Chuang, 1999; Gluck et al., 2000). Treatment with anti oxidants such as melatonin (Chen and Chuang, 1999) or ascorbic acid (MacGregor et al., 1996), attenuates KA-induced toxicity. A mechanism of KA-mediated neurodegeneration via ROS production has been proposed (Coyle and Puttfarcken, 1993; Olanow, 1993). Several lines of evidence suggest that KA treatment induces excess Ca2 + influx (Choi, 1992; Dutrait et al., 1995; Bar-Peled et al., 1996) and release of Ca2 + from intracellular Ca2 + stores (Frandsen and Schousboe, 1993) leading to increased intracellular Ca2 + concentrations. It is believed that activation of KA receptors results in a sodium (Na+ ) influx resulting in membrane depolarization (Choi, 1988) and, eventually, a 27 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. chloride (Cl ) influx (Van Damme et al., 2003). Depolarization activates the release of the magnesium (Mg2+ ) ion block on NMDA receptors and activation of NMDA receptors by glutamate released from the presynaptic terminal (Garthwaite, 1991) resulting in a Ca2 + influx through NMDA receptor-gated ion channels and voltage- gated Ca2 + channels (Coyle and Puttfarcken, 1993; Frandsen and Schousboe, 1993). Two possible pathways explaining how KA induces increased intracellular Ca2 + concentrations leading to cell death have been proposed (Olanow, 1993). In the first pathway, it is believed that the KA-induced Ca2 + influx may activate Ca2+ - activated neutral cysteine protease calpain I (Siman et al., 1989; Olanow, 1993) which eventually activates xanthine oxidase (Cheng and Sun, 1994), causing H 2O2 and 02*" formation (Dykens et al., 1987). The inhibition of xanthine oxidase is found to protect cerebellar neurons from KA-induced cell death (Dykens et al., 1987). H2O2 may trigger apoptosis through a caspase-mediated cascade (DiPietrantonio et al., 1999; Faherty et al., 1999). Moreover, in the presence of iron (Fe2 + ), H 2O 2 is converted into OH* through the Fenton reaction (reaction 2) (Aust et al., 1985; Coyle and Puttfarcken, 1993; Olanow, 1993; Shoham and Youdim, 2004). In the second pathway, it is believed that KA-induced Ca2 + influx activates NOS causing NO* formation (Olanow, 1993; Milatovic et al., 2002). Reaction between NO* and 02*’ results in ONOO' formation (Coyle and Puttfarcken, 1993) and eventually OH* formation (Leist and Nicotera, 1998). OH* is believed to cause functional alterations to lipids, proteins and DNA, leading cell death (Simonian and 28 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Coyle, 1996). It is also believed that ONOO can induce apoptosis by activating the caspase-3 cascade (Lin et al., 1998). It has been shown that following KA treatment, rate of ROS generation in rat cerebral cortical tissue culture cells increases (Bondy and Lee, 1993). Levels of both ROS and lipid peroxidation were also found to increase in rat cerebellar granular cell cultures and rat glial cell cultures following KA administration (Puttfarcken et al., 1993; Matsuoka et al., 1999). There is evidence suggesting that following KA treatment, activated caspase-3 expression is detectable in mouse brains (Becker et al., 1999; Faherty et al., 1999). The inhibition of caspase-3 activation is found to protect cortico-hippocampal cultures from KA-induced cell death (Osaka et al., 1999). The presence of the baculoviral caspase inhibitor p35 in mice brain both in vivo and in situ is also found to lower caspase-3 activities and decrease KA-induced cell death (Viswanath et al., 2000). 3.3. KA receptor subunits. There are five kainic acid receptor subunits: GluR5, GluR6, GluR7, KA1, and KA2. GluR5 forms receptors for glutamate, KA, domate and AMPA. GluR6 forms homomeric channels that are not sensitive to AMPA. GluR7 forms homomeric channel for glutamate but not AMPA or domate. GluR5-7 also forms heteromerically among each other and KA1 or KA2 to form functional kainic acid receptors. This results in a diversity of KA receptors in the brain (reviewed in Lerma, 2003). 29 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Porter et al., 1997 has reported the distribution of KA receptor subunit expression in the human hippocampus, neocortex and cerebellum (Porter et al., 1997). GluR5 mRNA is expressed only in Purkinje cells and some hippocampal neurons of the CA1. GluR6 expression was found ubiquitously, especially in the dentate gyrus, in pyramidal neurons of CA3, in cerebellar granule cells and in superficial and deep laminae of the neocortex. GluR7 expression was seen in the dentate gyrus, neocortex and cerebellum. KA1 mRNA was detectable in the dentate gyrus and CA3 but neither in the neocortex nor cerebellum. KA2 was abundantly found in the dentate gyrus, CA2, CA3 and the neocortex and cerebellum. Lower expression of KA2 was also detectable in CA1 and CA4. Recently, it has been reported that, at hippocampal mossy fiber synapses, GluR6 forms KA receptors with KA1 and KA2, presynaptically and postsynaptically, respectively (Darstein et al., 2003). 30 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. CHAPTER TWO GSHPx-1 Transgenic Mice Are More Susceptible To KA-induced Toxicity SUMMARY Local and systemic kainic acid (KA), a potent excitotoxin, have been shown to induce seizures and neuronal cell damage, primarily in the hippocampal CA3 area. Oxidative stress has been proposed to play an important role in KA-induced neuronal cell death. To investigate the role of oxidative stress in KA excitotoxicity, murine GSHPx-1 transgenic mice, with 4-6 fold increases in the enzyme activity, were used in this study. The animals were systemically administered with KA at a dosage of 35 mg/Kg body weight. After KA treatments, seizure severity and neuronal cell death were observed and compared to KA-treated wild-type controls. Interestingly, transgenic mice displayed increases in both seizure severity and neuronal cell damage, assessed histologically by terminal deoxynucleotide transferase-mediated dUTP nick-end labeling (TUNEL), haematoxylin and eosin, and cresyl violet. The neuronal damage was observed not only in the CA3 area but also in CA1, CA2 and CA4 of the hippocampus as well. An in vitro study also supported these findings; hippocampal cultures prepared from GSHPx-1 transgenic embryos were more sensitive to KA treatment than those from wild-type controls. 31 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. INTRODUCTION Kainic acid, a very powerful excitotoxin, is an agonist of non-NMDA receptors. Intracerebral and systemic administration of KA generates seizures and seizure-related neuronal cell death in several brain areas (Ben-Ari et al., 1979; Ben- Ari et al., 1980a; Ben-Ari et al., 1980b; Ben-Ari et al., 1981; French et al., 1982; Pollard et al., 1994). KA not only causes neuronal cell death in the CA3 region of the hippocampus, which is the most vulnerable region to KA toxicity (Pollard et al., 1994; Ben-Ari and Cossart, 2000), but the damage can also propagate to other regions such as CA1 and CA4 as well as the amygdala (French et al., 1982; Pollard et al., 1994; Venero et al., 1999). Although it is not fully understood how KA mediates neurodegeneration, oxidative stress has been proposed to play an important role in its toxicity (Coyle and Puttfarcken, 1993; Olanow, 1993; Cheng and Sun, 1994; MacGregor et al., 1996; Milatovic et al., 2002; Cheng et al., 2004; Parihar and Hemnani, 2004). Several studies have shown that treatments with antioxidants such as melatonin (Chen and Chuang, 1999; Dabbeni-Sala et al., 2001; Chung and Han, 2003) vitamin C (MacGregor et al., 1996) and vitamin E (Milatovic et al., 2002), iron-depleted diet (Shoham and Youdim, 2004) and genetically increased antioxidant enzymes such as catalase (Wang et al., 2003) and Cu/ZnSOD (Hirata and Cadet, 1997; Kondo et al., 1997; Schwartz and Coyle, 1998) can attenuate KA toxicity. However, some evidence suggests contradictory results, for example, treatment of GSH did not protect murine cortical cell cultures from KA toxicity (Regan and Guo, 32 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 1999), and overexpression of Cu/ZnSOD in cortical cultures derived from Cu/ZnSOD transgenic mice either did not protect (Ying et al., 2000) or exacerbated KA toxicity (Bar-Peled et al., 1996). Whether and how oxidative stress plays a role in KA toxicity still remains incompletely understood. GSHPx-1 is an important enzyme in protecting cells from oxidative stress by, with GSH, converting H 2O2 to water. Hypothetically, an increase of GSHPx activity, resulting form overexpression of GSHPx-1, should result in a decrease in H2O 2 levels leading to less OH* production. OH*, the most reactive of the reactive oxygen species, contributes to oxidative damage caused by its reactions with DNA, protein and lipid. In this study, GSHPx-1 transgenic animals were used to investigate the relationship between overexpression of GSHPx-1 and KA neurotoxicity in order to investigate whether GSHPx-1 has neuroprotective properties against KA-induced seizure activity and neuronal cell death. MATERIALS AND METHODS Materials. All chemicals were purchased from Sigma-Aldrich, Saint Louis, MO, unless otherwise stated. 33 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Transgenic animals. GSHPx-1 transgenic mice were generated by Y.S. Ho, Wayne State University, Detroit, Michigan (Spector et al., 1996; Yoshida et al., 1996). A mouse cellular GSHPx (GSHPx-1) clone, matched with a rat GSHPx-1 cDNA clone (Ho et al., 1988) was isolated from a bacteriophage FIX II genomic library. A 5.3-Kb Sac I fragment, containing 1.05-Kb of GSHPx-1 gene was isolated and subcloned into pBluscript SK (pSK), named pSK-mGP21. To prepare DNA for transgenic generation, the 5.3-Kb Sac I fragment containing 2.2 Kb of 5’ and 2.1 Kb of 3’ flanking sequences was purified and then microinjected into B6C3F1 fertilized eggs to create GSHPx-1 transgenic animals (Cheng et al., 1997). Animals used in the study were bred in-house and were housed according to standard animal care protocols, fed ad libitum, kept on a 12 hr light/dark cycle, and maintained in a pathogen-free environment in the Buck Institute Vivarium, accredited by AALAC. All animal experiments were approved by local IACUC review and conducted according to the policies on the use of animals of The National Institutes of Health. DNA extraction. 0.5 cm of mouse tail was cut from 3-week-old mice was digested in 600 (x l of extraction buffer (50 mM Tris-HCl, pH 8.0, 100 mM EDTA, 100 mM NaCl, 1% sodium dodecyl sulfate (SDS) and 35 |xl of 10 mg/ml Proteinase K in TE buffer (10 mM Tris-Cl, pH 8.0, 1 mM EDTA) at 55°C, overnight. DNA was isolated twice with phenol:chloroform and once with chloroform, then precipitated with cold 100% 34 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ethanol then washed with cold 70% ethanol. The purified DNA was then dissolved in TE buffer. The concentration of the DNA was then estimated from the absorbance at 260/280 nm using a spectrophotometer, DU 530 Life Science UV/Vis Spectrophotometer (Beckman Coulter, Fullerton, CA). The DNA concentration was diluted to 20 pg/ml in TE buffer for genotyping. Polymerase Chain Reaction. Polymerase chain reaction (PCR) was used for genotyping the animals using oligonucleotide primers (Figure 2) complementary to endogenous mouse GSHPx-1 gene and flanking regions of adjacent transgenes (Cudkowicz et al., 2002). Primer Sequence(5’- 3 ’) Wild-type : forward (WTF) CCT GTG ACT CAT CGG GAA ACC TC Wild-type : reverse (WTR) ATG GTA CGA AAG CGG CGG CTG TA 5X GSHPx-1 : forward (5XF) TGA AAC TGG AAT CCT ATT ATC C 5X GSHPx-1 : reverse (5XR) CTA ACA TGA GAC CAC AGA ACT G Table 2. Oligonucleotide primers for PCR analysis. WTF and WTR primers, complementary to intron 1 and exon 2 of endogenous mouse GSHPx-1 gene, respectively, generated a 400 bp product used as a positive control. 5XF and 5XR, complementary to the splice region of the GSHPx- 1 transgenes concatamers, generated a 163 bp product (Figure 3). 35 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. a WTF 5XF GSHPx-1 5XR WTR 1 Kb endogenous GSHPx-1 WTF PCR WTR _ 400 bp PCR product transgenic GSHPx-1 concatamer 400 bp PCR products 163 bp PCR products WTF 5XF WTF 5XF m - m — WTF 5XF m - m — WTF 5XF m - m — WTR 5XR WTR 5XR WTR 5XR WTF ^ _ m - m - WTR 5XR WTR PCR I Figure 3. PCR analysis: templates, oligonucleotide primers and PCR products, (a) WTF and WTR are complimentary to intron 1 and exon 2 of GSHPx-1 gene. 5XF and 5XR are complementary to the splice region of the GSHPx-1 transgenes, (b) A 400-bp PCR product generated from WTF and WTR was used as positive control. A 163-bp PCR product generated by 5XF and 5XR was used as an indication of transgenic GSHPx-1 (Cudkowicz et al., 2002). Each PCR reaction contained: 2 pi of 20 pg/ml of extracted DNA template, 1 pi of PCR buffer (160 mM (N H ^S O ^ 670 mM Tris, pH 8.8, 1% Tween 20, and 15 mM MgCl2), 2 pi of 1 mM dNTPs, 0.45 pi of WTF primer, 0.45 pi of WTR primer, 0.3 pi of 5XF primer, 0.3 pi of 5XR primer, 3.43 pi of double distilled water (ddH20) and 0.07 pi of TAQ polymerase (Eppendorf, Westbury, NY). A PCR 36 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. program of 30 cycles of a 58-second denaturation at 94°C followed by a 65-second hybridization at 58°C and a 60-second extension at 72 extension temperature, was performed in an Eppendorf Mastercycler PCR machine (Eppendorf, Westbury, NY). The PCR products were separated on a 1.8% agarose gel in IX TAE buffer (40 mM Tris, pH 8.0, 20 mM acetic acid, 1 mM EDTA) containing 0.25 mM ethidium bromide at a constant voltage of 100 volts (Model 250 Power Supply, Life Technologies, Carlsbad, CA), visualized and photographed using a Chemilmager (Alpha Innotech Corporation, San Leandro, CA). A 100-bp DNA ladder maker (Invitrogen, Life Technologies, Carlsbad, CA) was used to estimate the sizes of the PCR products. GSHPx activity assay. A GSHPx assay kit (BIOXYTECH GPx-340, Oxis International, Portland, Oregon) was used to determine hippocampal GSHPx activity levels. The assay is based on a change in NADPH absorbance at 340 nm. Briefly, animals were anesthetized with ether then perfused with 0.9 % NaCl containing 0.16 mg/ml heparin. Hippocampi were removed and homogenized in cold 50 mM Tris-HCl, pH 7.5, 5 mM EDTA, 1 mM 2-mercaptoethanol and supernatant collected after centrifugation at 10,000 x g for 10 minutes at 4°C. Protein concentration was first estimated by using the Bradford reagent (Bio-Rad, Hercules, California). 15 pi of hippocampal lysate was then added to 150 pi of 0.05 M Tris-HCL, pH 7.6, 5 mM EDTA containing 0.24 pmol glutathione, 0.12 U glutathione reductase (GR) and 37 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 0.048 jimol (3-NADPH. 75 pL of 0.007% tert-butyl hydroperoxide was added to the sample mixture. Absorbance was recorded at 340 nm for 3 minutes. GSHPx activity levels were determined against a standard curve. One GSSG activity unit is defined as the amount of enzyme catalyzing the oxidation of one 1 pM of NADPH per minute. Though this assay measures all GSHPx activities and is not specific to GSHPx-1, since the GSHPx-1 is the most abundant GSHPx, the GSHPx activity is expressed as GSHPx-1 activity (Spector et al., 1996). In vivo treatment with KA and seizure activity assessment. A dosage of 35 mg/kg body weight KA (Ocean Produce International, Nova Scotia, Canada) in 1 x PBS was injected intraperitoneally into 8-12 week old homozygous transgenics, heterozygous transgenics and wild-type mice. For a period of 2 hours after KA administration, seizure activities of each animal were observed and rated according to an arbitrary scale (Yiswanath et al., 2000): 0, no seizures; 1, head nodding, staring, very light seizures of forepaws; 2, repeated mild seizures, rearing and loss of postural control; 3, whole body seizures for a short time; 4, seizures become severe and prolonged. Histological staining for CA3 hippocampal cell death. 24 hours after KA administration, brains were collected, fresh frozen and kept at -80°C. 16-pm coronal sections through the hippocampi were obtained from the fresh frozen brains using an Microm 505 cryostat (RMC Cryosystems, Tucson, AZ). Sections were stained by Terminal Deoxynucleotide Transferase-mediated 38 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. dUTP Nick-End Labeling (TUNEL), cresyl violet (CV) and haematoxylin and eosin (H&E) stain. For TUNEL staining, a TdT-Fragel DNA fragmentation detection kit (Oncogene Research Products, Cambridge, Massachusetts) was used as per the manufacturer’s manual. Briefly, brain coronal sections were fixed in 4% formaldehyde in 1 x phosphate buffered salt (PBS), pH 7.4 (Invitrogen, Life Technologies, Carlsbad, CA) at room temperature, for 15 minutes then washed in lx Tris buffered saline (TBS), pH 8 (20 mM Tris, pH 8.0, 140 mM NaCl) for 15 minutes at room temperature. The specimens then were permeabilized with 20 pg/ml proteinase K in 10 mM Tris, pH 8.0 for 10 minutes at room temperature and briefly washed with lx TBS. Endogenous peroxidase in the sections was inactivated by a treatment of 3% H 2O2 in methanol for 5 minutes at room temperature. The sections were, again, briefly rinsed with lx TBS, covered with lx TdT Equilibration Buffer (0.2 M sodium cacodylate, 30 mM Tris, 0.3 mg/ml BSA, 0.75 mM C 0 CI2, pH 6.6) and incubated at room temperature for 30 minutes. After the 30-minute incubation, the TdT Equilibration Buffer were blotted from the sections, TdT Labeling Reaction Mixture (a mixture of biotin-labeled and unlabeled deoxynucleotides, and TdT enzyme) was applied to the sections, followed by a 2-hour incubation at 37°C. The sections were then rinsed with lx TBS. To terminate the labeling reaction, Stop Solution (0.5 M EDTA, pH 8.0) was applied to the specimens, followed by a 5- minute incubation at room temperature. Blocking Solution (4% BSA in IX PBS) was then applied to block additional labeling. After a 10-minute incubation, the Blocking 39 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Solution was blotted from the sections. The biotinylated deoxynucleotides bound the exposed 3’-OH end of DNA fragments catalyzed by TdT enzyme were detected using lx Conjugate (a streptavidin-horseradish peroxidase conjugate). The sections were covered with the lx Conjugate, incubated in a humidified chamber for 30 minutes, at room temperature and rinsed with lx TBS. Then the streptavidin- horseradish-biotinylated deoxynucleotides were colorized using DAB solution (0.7 mg/ml 3,3’ diaminobenzidine and 0.6 mg/ml ITCVurea in tap water). After 15 minutes, the sections were rinsed in dHaO. In the final step, the sections were counterstained with methyl green and mounted using a xylene-based mounting medium, Cytoseal 60 (Stephen Scientific, Kalamazoo, MI). TUNEL-positive cells, which appear dark brown in color, were viewed and quantitated under a light microscope. For CV staining, brain coronal sections were immersed in 0.5 % cresyl violet for 15-30 minutes. Then they were differentiated in running tap water for 3-5 minutes. The sections were then dehydrated through a series of 70%, 80%, 90%, 95% and 100% ethanol, 1 minute each, and xylene for 5 minutes, and mounted using Cytoseal 60. Stained sections, with stained chromatin and Nissl substances appear violet in color, were viewed and compared with adjacent TUNEL-stained sections under a light microscope. For H&E staining, brain coronal sections were immersed in 80% ethanol for 5 minutes and rinsed with dH^O. The sections were stained with Haematoxylin for 5- 10 minutes, rinsed with running water for 5 minutes to allow the stain to develop, 40 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. immersed in acid ethanol (1% HC1 in 75% ethanol) for 1 minute and stained with Eosin Y for 30 seconds. The sections were immersed in 95% ethanol for 30 seconds, 100% ethanol for 5 minutes then xylene for 5 minutes and mounted using Cytoseal 60. The sections were viewed and compared with adjacent TUNEL-stained sections under a light microscope. Haematoxylin-stained nuclei and Eosin Y-stained cytoplasm appeared blue to purple and red to pink, respectively. Quantitation of TUNEL staining in the hippocampal area. The severity of cell death in hippocampal (CA) areas was evaluated in each area, CA1, CA2, CA3 and CA4 on the following arbitrary score: + for when less than 20% cells were TUNEL positive, ++ for when 20-50% of cells were TUNEL positive, and +++ when more than 50% of cells were TUNEL positive (Jiang et al., 2000). In vitro treatment with KA and cell viability assay. Primary hippocampal neuronal cultures were isolated from E l8 animals. Briefly, hippocampal tissues were dissected in Ca2+ -Mg2 + free Hank’s salt solution (Life Technologies, Carlsbad, California) and digested with 0.25% EDTA-trypsin. Cells were dissociated in 10% fetal bovine serum (FBS)-DMEM medium (Life Technologies, Carlsbad, California) and plated onto poly-L-lysine coated 48-well plates. One hour after plating, the medium was changed to Neurobasal medium (Life Technologies, Carlsbad, California) with B-27 supplement Minus AO (Life 41 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Technologies, Carlsbad, California) and 2 mM glutamine (Life Technologies, Carlsbad, California) to remove non-adherent cells. Cells were maintained in a humidified atmosphere of 95% air and 5% CO 2 at 37°C. Medium was half-changed every 4 days. Cultures were treated with 10 pM cytosine P-D-arabinofuranoside on days 3 and 6 to remove non-neuronal cells. At days 11-14 in vitro (DIV), the cultures were treated with various concentrations of KA in neurobasal medium with B-27 supplement minus AO and glutamine: 0, 25, 50, 100, 250 and 500 pM for 24 hours. Cell viability assay was performed on KA-treated cultures by incubating cultures with 5 mg/ml of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) at 37°C for 3 hours. At the end of the incubation, the medium was removed and the converted dye was solubilized with DMSO. Absorbance of converted dye was measured at a wavelength of 570 nm with background subtraction at 660 nm using a SpectraMAX 340PC spectrophotometer (Molecular Devices, Sunnyvale, California). RESULTS Genotyping using PCR. GSHPx-1 transgenic mice used in the current experiments exhibited no physical abnormalities and grew and reproduced normally as reported elsewhere (Cheng et al., 1997; Cheng et al., 1998). For genotyping the animals, two sets of oligonucleotide primers complementary to the endogenous mouse GSHPx-1 gene 42 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. and flanking regions of concatamer transgenes were used in the PCR, generating a 400 bp PCR product, a positive control, and a 163 bp product, an indication of the presence of at least one copy of the transgene (Figure 4). M l 2 3 4 5 6 7 8 9 400 bp PCR product 163 bp PCR product Figure 4. PCR products. PCR products were separated on a 1.8% agarose gel in 1 x TAE buffer containing 0.25 mM ethidium bromide; M, 100-bp DNA ladder marker; lane 1-3, PCR product generated from wild-type animals DNA templates; lane 4-6, PCR products generated from GSHPx-1 transgenic animals DNA templates; lane 7, PCR product generated from a known wild-type animal DNA template, positive control; lane 8, PCR product generated form a known GSHPx-1 transgenic DNA template, positive control; lane 9, PCR product generated in a reaction with out DNA template, negative control. GSHPx-1 transgenic animals have increased levels of hippocampal GSHPx-1 activity. To investigate hippocampal GSHPx activity levels, an assay based on the rate of NADPH consumption was used. GSHPx-1 in concert with GSH is able to reduce H2 0 2 to water. GSH is oxidized in the process and becomes a disulfide dimer, GSSG, which can be reduced to GSH by GR, catalyzed by NADPH. So the change in NADPH absorbance indirectly represents the activity of GSHPx-1. 43 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. C A 2 0 0 -i HM HT W T Figure 5. Hippocampal GSHPx activity in HM, HT and WT animals. GSHPx, glutathione peroxidase; HM, homozygous; HT, heterozygous; WT, wild-type, n = 4- 5, * p = 0.01 HM vs. HT, ** p = 0.01 HM and HT vs. WT. Hippocampal GSHPx activity in HT (heterozygous, 1 copy of the mouse GSHPx transgene) demonstrated a four-fold increased in GSHPx activity (102.49 ± 11.44 mU/mg protein) and HM (homozygous, 2 copies of the transgene) a six-fold increase (160.10 ± 6.05 mU/mg protein) compared to wild-type (WT) hippocampi (26.38 ± 4.51 mU/mg protein) (Figure 5). In the same line of GSHPx-1 transgenic mice, increased mRNA GSHPx-1 levels have been reported in heart, eye, brain, lung, muscle, spleen, tongue and kidney (Yoshida et al., 1996; Cheng et al., 1997). Also, increased GSHPx activity levels in several organs have been reported including lens epithelial cells, brain, heart, lung, muscle, kidney, intestines and stomach (Spector et al., 1996; Yoshida et al., 1996; Cheng et al., 1997; Cudkowicz et al., 2002). It has been reported that increase of GSHPx-1 level does not have effect on basal levels of other anti oxidant 44 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. enzymes such as GSHPx-P, PHGSHPx, glutathione-S-trasferase (GST), MnSOD, Cu/ZnSOD, GR, catalase and G6PD (Yoshida et al., 1996; Cheng et al., 1997). GSHPx-1 transgenic animals displayed increased KA-induced seizure activities. u > < u C/3 < U >- 3 N '5 3 1/3 « 4 - l o < u u t- b O < D o HM HT WT 1 1 1 1 0 30 60 90 120 ir 30 60 90 120 Time (min) 30 60 90 120 Figure 6. Assessment of KA-induced seizure activity in KA-treated animals. Representative seizure severity and frequency in HM (homozygous, 2 copies of the mouse GSHPx-1 transgene) and HT (heterozygous, 1 copy of the transgene) vs. WT (wild-type) animals (n = 3 each) over a 2-hour period following systemic KA administration. 0, no seizures; 1, head nodding, staring, very light seizures of forepaws; 2, repeated mild seizures, rearing and loss of postural control; 3, whole body seizures for a short time; 4, seizures become severe and prolonged. Higher levels of hippocampal GSHPx activity in transgenic animals (6-fold in HM and 4-fold in HT) were hypothesized to be protective against the effects of 45 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. systemic KA treatment. 8-12 weeks old animals were intraperitoneally injected with KA (35 mg/kg body weight) and for a period of 2 hours immediately after KA administration, seizure activities were observed, scored and graphed. Within 20-30 minutes following KA administration, mice began to display motionlessness, head nodding, staring and very light seizure of forepaws (arbitrary scale 1). The next 30 minutes, mice began to display symptoms of mild seizure (scale 2, rearing and loss of postural control), and occasionally, the whole body seizures for a short period (scale 3) as well as heavy salivation and foaming at the mouth occasionally mixed with blood. The next 1-2 hours, mice developed full status epilepsy (scale 4) alternating with repeated mild seizure (scale 2) and short period whole body seizure (scale 3). The animals that experienced scale 3 and scale 4 were defined as having severe seizure in this study. Surprisingly, HM and HT mice showed a clear increase in the frequency and severity of seizures induced with a dosage of 35 mg/Kg body weight compared to WT animals. Figure 6 represents an experiment where animals, 3 of each genotype, were intraperitoneally injected with KA. All 3 HMs and 3 HTs animals displayed severe KA-induced seizure (scale 3 and 4) while WT animals displayed only mild seizure during a 2-hour period observation immediately following KA administration. Overall, 63.41% of KA-treated HMs (26 out of 41), 48.84% of KA- treated HTs (21 out of 43) and 35.71% of KA-treated WTs (15 out of 42) displayed whole body seizure (scale 3) and/or severe, prolonged seizures (scale 4) (Figure 7). 46 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 80 70 - C/3 13 I 60 H § x> 50 - < D jg 40 - 3 3 0 “ «g 2 0 - £ 10 - HM HT WT Figure 7. Increased GSHPx activity results in increased KA-induced seizure activity. Percentages of KA-administered animals displaying whole body seizure (scale 3) and/or severe, prolonged seizure (scale 4); HM, homozygous, 2 copies of the mouse GSHPx-1 transgene; HT, heterozygous, 1 copy of the transgene; WT, wild-type animals; n = 41, 43 and 42, respectively. Systemic KA administration induced increased neuronal cell death in GSHPx-1 transgenic animals. To investigate if systemic KA administration would also result in increased neuronal cell death in GSHPx-1 transgenic animals, the degree of neuronal damage in hippocampus was assessed using TUNEL, CV and H&E stainings. Although TUNEL staining principally is based on in situ DNA fragmentation detection, a specific characteristic of apoptotic cell death, in this experiment TUNEL staining was used as an indicator of neuronal damage without distinguishing whether it was apoptotic or necrotic (Charriaut-Marlangue and Ben-Ari, 1995; Jiang et al., 2000). 24 hours following KA injection, brains of surviving animals were removed, fresh-frozen, sectioned and stained with TUNEL, CV and H&E. Coronal brain 47 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. sections showed increased damage in area CA3 in both HM and HT compared to WT mice. Almost half of all KA-treated HMs (47.62%, 10 out of 21), 28% of KA- treated HTs (7 out of 25) and 19.05% of KA-treated WTs (4 out of 21) also demonstrated damage to the CA3 (Figure 8). Not all surviving animals that displayed severe KA-induced seizure showed neuronal damage. 80 3 70 " ^ 60 - e a < D t* 50 - 40 - a 3° - 20 - o 10 - HM HT WT Figure 8. Increased GSHPx activity results in increased KA-induced neuronal cell death. Percentages of systematic KA-administered animals displaying neuronal cell death in CA1, CA2 and CA3 regions of hippocampus; HM, homozygous, 2 copies of the mouse GSHPx-1 transgene; HT, heterozygous, 1 copy of the transgene; WT, wild-type animals; n = 21, 25 and 21, respectively. Although, it has been shown that the CA3 area of the hippocampus is the most vulnerable brain area to seizure-induced cell death (Ben-Ari, 1985) some cell damage was observed in CA1, CA2 and CA4 (Figure 9). 48 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Reproduced w ith permission o f th e copyright owner. Further reproduction prohibited without permission. HM HT WT -i^ c v • • ■ - ~ - £ v * H&E 1 m 'm m ’ Mm “sm .. . .■ •■ • TUNEL ' '^iSSs- * 1 ' . f * - i^ s - • » j □ Figure 9. Assessment of hippocampal neuronal cell death. Assessment of hippocampal neuronal cell death, using cresyl violet (CV), haematoxylin & eosin (H&E) and terminal deoxynucleotide trasferase-mediated dUTP nick-end labeling (TUNEL) stainings, in representative coronal sections collected from animals 24 hours after systemic KA administration. Top insets, CA1 region; side insets, CA3 region. HM, homozygous, 2 copies of mouse GSHPx-1 transgene; HT, heterozygous, 1 copy of the transgene; WT, wild-type animals; bar = 1 mm. Hippocampal primary cultures isolated from GSHPx-1 transgenic animals displayed increased susceptibility to KA toxicity. Figure 10. Increased GSHPx activity also results in increased susceptibility to KA in vivo. Cell death in primary hippocampal neuronal cultures from HM vs. WT mice 24 hours following KA treatment at dosages of 0, 25, 50, 100, 250 and 500 pM. HM, homozygous, 2 copies of the mouse GSHPx-1 transgene; WT, wild-type animals; n = 4. Open bars, WT; filled bars, HM; * p = 0.01. To study whether or not GSHPx-dependent susceptibility to KA toxicity also occurs in vitro, the effects of KA on dispersed hippocampal primary neuronal cultures from HMs and WTs were studied. Cultures were treated with various concentrations of KA (0-500 pM) and cell viability assayed 24 hours following KA addition. At lower concentrations of KA (25 pM), no significant differences between HM and WT were observed (Figure 10). However at higher concentrations of KA (50, 100, 250 and 500 pM), a significantly reduced level of cell survival was 125 0 Z * * * p — ■ — * — i 1 I I I I I 0 25 50 100 250 500 K A (pM ) 50 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. observed in the HM cultures (10-12% of untreated cultures). In contrast, higher concentrations of KA did not increase the amount of cell death in WT cultures. DISCUSSION Neurotoxicity of kainic acid was first demonstrated by Olney and colleagues (Olney et al., 1974). Both local and systemic administration of kainic acid can cause acute and sub-acute epileptical seizure activity, and seizure-related neurodegeneration, most prominently in the CA3 area of the hippocampus (Ben-Ari et al., 1979; Ben-Ari et al., 1980a; Ben-Ari et al., 1980b; Ben-Ari et al., 1981; French et al., 1982; Pollard et al., 1994). Since KA administration has be shown to increase production of oxidative stress markers such as lipid peroxidation, protein oxidation and DNA oxidation, oxidative stress has been proposed to play an important role in KA-induced neurodegeneration (Coyle and Puttfarcken, 1993; Bruce and Baudry, 1995; Carrasco et al., 2000; Gluck et al., 2000; Liang et al., 2000; Patel et al., 2001; Patel and Li, 2003). Despite several studies having shown that antioxidants prevent the toxicity (MacGregor et al., 1996; Hirata and Cadet, 1997; Kondo et al., 1997; Schwartz and Coyle, 1998; Chen and Chuang, 1999; Dabbeni-Sala et al., 2001; Milatovic et al., 2002; Chung and Han, 2003; Wang et al., 2003), some studies have reported the opposite (Bar-Peled et al., 1996; Regan and Guo, 1999; Jiang et al., 2000; Ying et al., 2000). The relationship between antioxidants and KA-induced neurodegeneration therefore requires further investigation. 51 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Activation of AMPA/KA receptors by KA causes a Na+ influx (Choi, 1988) resulting in depolarization of the cell membranes which, in turn, triggers Cl' influx (Van Damme et al., 2003). More sustained depolarization of the membrane triggers release of the Mg2 + blockade which activates NMDA receptors causing prolonged neuronal bursting and repetitive discharges, in turn, generating limbic seizure (Stanton et al., 1987; Chen et al., 1999), and allowing a Ca2 + influx leading to neuronal cell death (Garthwaite, 1991). Treatment with a non-competitive NMDA receptor antagonist such as MK-801 (Clifford et al., 1990; Baran et al., 1994), antioxidants such as melatonin (Skaper et al., 1999; Chung and Han, 2003), vitamin C (MacGregor et al., 1996) and vitamin E (Milatovic et al., 2002), and iron-depleted food (Shoham and Youdim, 2004) has been shown to attenuate KA-induced toxicity. Since GSHPx-1 is the major cellular form of GSHPx, a key enzyme in counteracting ROS, initially we hypothesized that the increased activity levels of the enzyme in transgenic mice would protect the mice from KA toxicity. Interestingly, mice with increased activity levels of GSHPx-1 were more susceptibility to KA toxicity compared to wild-type animals as assessed by both seizure activity and neuronal cell death. Despite increased GSHPx activity levels, GSHPx-1 transgenic mice were more susceptible to KA-induced neurotoxicity compared to wild-type animals. GSHPx-1 transgenic displayed 4- and 6 -fold increased levels in hippocampal GSHPx activity in HT and HM animals, respectively, compared to wild-type 52 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. animals. Increases in GSHPx-1 expression levels (mRNA) and GSHPx activity levels have been reported in several organs of the same line of transgenic mice including the heart, eye, brain, lung, muscle, and kidney (Spector et al., 1996; Yoshida et al., 1996; Cheng et al., 1997; Cudkowicz et al., 2002). Whole brain GSHPx activity levels in GSHPx-1 transgenic mice has been reported to be increased 3-fold (Yoshida et al., 1996) and 4-fold increased (Cudkowicz et al., 2002) compared to wild-type which is comparable to 4- to 6 - fold increases of hippocampal GSHPx activity observed in this study, depending on the genotype of the animals, i.e. HT, single copy of the transgene or HM, 2 copies of GSHPx-1 transgene. In spite of the 4-6 fold increased GSHPx activity levels, systematic KA administration, at a dosage of 35 mg/Kg body weight, surprisingly, caused increased in vitro and in vivo KA-mediated neuronal cell death and KA-induced seizure activities in GSHPx-1 transgenic mice. This contradicts several reports suggesting either increases in GSHPx activity levels could attenuate oxidative stress mediated damage or decreases in GSHPx activity levels could exacerbate such damage. For example, Kelner and co-workers demonstrated that human breast cancer cells overexpressing human GSHPx-1 were more resistant to paraquat toxicity (Kelner et al., 1995). An ovary cell line, with an 80% decrease of GSHPx activity, became more sensitive to oxidative-mediated paraquat and adriamycin toxicities (Taylor et al., 1993). Cortical neuron cultures isolated from GSHPx-1 knock out mice showed increased susceptibility to H 2O2 toxicity in a dose-dependent manner (de Haan et al., 53 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 1998). Hepatocytes isolated from GSHPx-1 knock out mice were more susceptible to diaquat, an ROS generator, compared to untreated cells (Fu et al., 2001). In terms of in vivo studies, Yoshida and co-workers (Yoshida et al., 1996) demonstrated that a 5-fold overexpression of murine GSHPx-1 in a separate line of transgenic mice protected the animals from ischemia reperfusion-mediated myocardial injury through the enzyme’s ^CV scavenging property. Overexpression of human GSHPx-1, resulting in a 4-fold increase in GSHPx activity in transgenic mice, also protected the animals against focal cerebral ischemia/reperfusion (Weisbrot-Lefkowitz et al., 1998). Human GSHPx-1 overexpressing mice, with 38% and 62% increases in enzyme activities in macrophage and plasma, respectively, have been reported to be resistant to lipopolysaccharide (LPS)-induced endotoxemia which is mediated through oxidative stress (Mirochnitchenko et al., 2000). Overexpression of human GSHPx-1 in mice was also reported to protect the hearts against left ventricular failure after myocardial infarction (Shiomi et al., 2004). Almost completely depleted GSHPx activity in homozygous GSHPx-1 knock-out mice has been shown to increase the susceptibility of the animals to paraquat toxicity (de Haan et al., 1998). However, a number of studies reported adverse effects of overexpression of GSHPx-1. For example, an increase in GSHPx activity in human GSHPx-1- containing transgenic mice resulted in thermosensitivity of the animals which was triggered by lowered levels of HSP70 activation due to lower levels of H 2O 2 and lipid peroxides (Mirochnitchenko et al., 1995). A 1-fold increase in human GSHPx-1 54 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. in transgenic mice also resulted in increased tumorigenesis of the animals (Lu et al., 1997). Overexpression of human GSHPx-1 in transgenics also failed to prevent acetaminophen-induced hepatic cell death resulting in an increased mortality of the animals (Mirochnitchenko et al., 1999). In addition, a study in GSHPx-1 knock out mice has shown that the animals were more resistant, rather than susceptible, to KA- toxicity compared to wild-type animals (Jiang et al., 2000). Moreover, ebselen, 2- phenyl-l,2-benzisoselenazol-3(2H)-one, a seleno-organic compound possessing GSHPx-like activity and anti-inflammatory properties, has been shown to induce apoptosis in a hepatoma cell line (Yang et al., 2000a; Yang et al., 2000b). Several studies have suggested that the possible reason for the inability of elevated GSHPx-1 activity level to efficiently remove H 2O2 may be that the system was unable to provide and/or maintain a sufficient level of endogenous intracellular GSH leading to an imbalance in the GSH/GSSG ratio (Mirochnitchenko et al., 1995; Spector et al., 1996; Fisher et al., 1999; Yang et al., 2000a; Yang et al., 2000b). A further study on the imbalance of GSH/GSSG was conducted to elucidate whether an alteration in the GSH/GSSG ratio may play a role in the adversarial, instead of protective, effect of elevated GSHPx-1 in terms of KA neurotoxicity. 55 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. CHAPTER THREE GSHPx-1 Transgenic Mice Exhibit Increased Hippocampal GSSG Levels Resulting in Increased Susceptibility to KA Toxicity SUMMARY Possible mechanisms underlying the unanticipated increased susceptibility of the GSHPx-1 transgenic mice to KA-induced toxicity were investigated in this current study. In the previous study, described in Chapter Two, we demonstrated that even though the transgenic mice had 4-6 fold elevations in GSHPx activity levels in the hippocampus compared to wild-type animals, they were more susceptible to KA- toxicity i.e. displayed increased seizure activity and neuronal damage. Our study suggests that, despite a small increase in GR, the increase in GSHPx and decreases in G6 PD and 6 PGD enzyme activity levels, all important enzymes in the GSH- regenerating cycle, may cause an accumulation of GSSG in the hippocampus of the GSHPx transgenics resulting in a shift of cellular redox environment. This in turn leads to cellular GSSG efflux in an attempt to maintain the reducing intracellular redox state and to prevent subsequent GSSG toxicity. This in turn results in an increased basal hippocampal NMDA-evoked response in the transgenics compared to wild-type animals which is independent of the NMDA receptor redox state or 56 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. NMDA receptor number, likely due to a direct activation of the receptor by the excreted extracellular GSSG. INTRODUCTION Oxidative stress has been proposed to play an important role in KA-induced toxicity (Coyle and Puttfarcken, 1993; Olanow, 1993; Milatovic et al., 2002; Cheng et al., 2004; Parihar and Hemnani, 2004). Free radical scavengers and antioxidant enzymes have been reported to attenuate such toxicity (MacGregor et al., 1996; Schwartz and Coyle, 1998; Chen and Chuang, 1999; Milatovic et al., 2002; Chung and Han, 2003; Wang et al., 2003). Therefore, we initially hypothesized that an increase in H 2 0 2 -scavenging GSHPx enzyme activity levels resulting from overexpression of GSHPx-1 in transgenic mice would protect the animals from KA toxicity. Interestingly, in our previous study (Chapter Two), we observed that despite 4-6 fold increases in GSHPx activity levels in the GSHPx-1 transgenic mice, these animals were more susceptible to KA toxicity, displaying increased seizure activity and increased neuronal damage. A recent report has demonstrated that an alteration in the GSH regenerating cycle i.e. GSHPx/GR ratio can effect the ability of the system to cope with oxidative insults (Spector et al., 1996). The GSH system is important in maintaining a reducing environment in cells and tissues (Filomeni et al., 2002a). An increase in GSSG and/or a decrease in GSH levels, in response to oxidative insults, could result in a shift of the intracellular redox environment 57 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. (Cereser et al., 2001; Schafer and Buettner, 2001; Armstrong and Jones, 2002; Biroccio et al., 2002; Filomeni et al., 2002b; Monteiro et al., 2004). There are several molecular mechanisms to restore the GSH redox status, for example increase of GR activity (Filomeni et al., 2002a) and/or GSSG transport into the extracellular space (Meister and Anderson, 1983; Kondo et al., 1987; Hirrlinger et al., 2001). However in neuronal cells, extracellular GSSG has been reported to activate NMDA receptors resulting in increased intracellular Ca2 + (Leslie et al., 1992). Since the GSHPx-1 transgenics display 4-6 fold increased levels of GSHPx activity levels compared to wild-type animals, hypothetically it would be possible that this could result in disruption of the GSH system in the transgenic animals in response to oxidative stimuli, leading to increased intracellular GSSG accumulation and GSSG efflux which in turn in hippocampal neurons could activate the NMDA receptor. In this study, we explore whether KA induces a shift in the GSH redox status in the transgenics leading to GSSG efflux and activation of the hippocampal NMDA receptors. MATERIALS AND METHODS Materials. All chemicals were purchased from Sigma-Aldrich, Saint Louis, MO, unless otherwise stated. 58 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Transgenic animals and primary hippocampal neuronal cultures. The methods are the same as those previously described in the Material and Methods section of Chapter Two. GSH/GSSG ratios. A GSH/GSSG assay kit (BIOXYTECH GSH/GSSG-412, Oxis International, Portland, Oregon) was used to determine the GSH/GSSG ratio. The assay is based on the coloric development of 5,5’-dithiobis-2-nitrobenzoic acid (DTNB) reacting with GSH at 412 nm. Briefly, brains were homogenized in 1 ml of ice-cold 0.05 M Tris- HCL, pH 7.6 containing 5 mM EDTA and 1 mM mercaptoethanol with or without 33 mM l-methyl-2-vinylpyridine (M2VP), a GSH scavenger. Supernatant was collected after a centrifugation at 10,000 x g for 10 minutes at 4°C. Protein concentrations of the lysates were estimated using the Bradford reagent (Bio-Rad, Hercules, California). Samples were then extracted with 3 volumes of cold 5% metaphosphoric acid (MPA). MPA extracts were collected after a centrifugation at 1,000 x g for 10 minutes at 4°C. 200 pi of 1.262 mM DTNB in 100 mM NaP0 4 containing 5 mM EDTA, pH 7.5 with 5% ethanol and 3 U of GR in 100 mM NaP0 4 containing 5 mM EDTA, pH 7.5 were added to 200 pi of the extract. After a 5- minute incubation at room temperature and addition of 0.1 mmol NADPH, the absorbance was recorded at 412 nm for 3 min. GSH and GSSG levels were determined against a standard curve. GSH/GSSG ratios were calculated from the formula (GSH-2GSSG)/GSSG. 59 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Brain synaptosomal preparations. Brain synaptosomes were prepared from the whole brain obtained 24 hours following KA or saline treatment using a modified protocol of a previously described procedure (Chalmers and Nicholls, 2003). Briefly, brains were removed and homogenized with a Dounce homogenizer in 5 ml of ice-cold isolation buffer containing 320 mM sucrose, 5 mM Tes, 1 mM EGTA, pH 7.2). The homogenate was centrifuged at 1,000 x g for 10 minutes at 4°C. After collecting the supernatant, the pellet was re-homogenized in 5 ml of ice-cold isolation buffer and centrifuged at 1,000 x g for 10 minutes at 4°C. The supernatant from the secondary homogenization was pooled with the supernatant collected previously. The pooled supernatant was centrifuged at 8,500 x g for 10 minutes at 4°C. The pellet was then resuspended in 2 ml of ice-cold isolation buffer, layered onto a discontinuous gradient consisting of 3 ml 6 % Ficoll, 1 ml 9% Ficoll and 3 ml 12% Ficoll (all prepared in isolation buffer) and centrifuged at 75,000 x g for 30 minutes. A doublet of synaptosomal fractions formed in 9% Ficoll layer was collected and pooled together. The pooled synaptosomal fraction was then resuspended with 2 ml of ice-cold 250 mM sucrose, 10 mM Tes, 0.16 pM bovine serum albumin, pH 7.2 and centrifuged at 15,000 x g for 10 minutes at 4°C. The synaptosome-containing pellet was then resuspended in ice-cold 100 pi of the 250 mM sucrose buffer for protein concentration estimation (Bio-Rad, Hercules, California). 60 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. H 2O2 assay. An H 2O2 assay kit (Amplex Red Hydrogen Peroxide Assay Kit, Molecular Probes, Eugene, OR) was used to determine whole brain synaptosomal hydrogen peroxide levels. The assay was performed per manufacturer’s instructions. In principle, the assay uses a H 2O2 sensitive and stable probe, 10-acetyl-3,7- dihydroxyphenoxazine (Amplex Red reagent). In the presence of horseradish peroxidase (HRP), the Amplex Red reagent reacts with H 2O2 with a 1:1 stoichiometry resulting in stable and highly fluorescent resorufin. The fluorescence is measured in a fluorescence microplate spectrofluorometer (Gemini, Molecular Devices, Sunnyvale, CA) using an excitation wavelength of 345 nm and an emission wavelength of 590 nm. Sample H 2O2 levels were calculated from a standard curve constructed from fluorescence measurements of a set of known H 2O 2 concentration standards. GR, Glucose-6-phosphate dehydrogenase (G-6-PD) and 6-phosphogluconate dehydrogenase (6-GPD) assays. Brains were homogenized in cold lx PBS containing 1 mM EDTA. After centrifugation at 8,500 x g for 10 minutes at 4°C, supernatants were collected for protein concentration estimation (Bio-Rad, Hercules, California) and enzyme activity assays. A GR assay kit (BIOXYTECH GR-340, Oxis International, Portland, Oregon) was used to determined brain GR activity levels according to the manufacturer’s instructions. The assay is based on the oxidation of NADPH to 61 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. NADP+ catalyzed by GR. GR activity levels were determined by a change in NADPH absorbance at 340 nm. One GR activity unit is defined as the amount of GR catalyzing the reduction of 1 pM of GSSG per minute. A G-6 -PD/6 -GPD assay kit (BIOXYTECH G6PD/6PGD-340, Oxis International, Portland, Oregon) was used to determined brain G-6 PD and 6 -GPD levels, according to the manufacturer’s instructions. The assay is based on the increase of NADPH absorbance at 340 nm catalyzed by G-6 -PD and 6 -GPD. One unit of either enzyme activity is defined as the amount of enzyme producing 1 pM of NADPH per minute. Western blot analysis of NMDA receptor subunits. Hippocampi (n=3 each group, HM, HT, WT) were dissected 24h after 0.9% NaCl administration and homogenized in RIPA buffer (1 x PBS, 1% Igepal CA-630, 0.5% sodium deoxycholate, 0.1% SDS containing 10 mg/ml PMSF and 30 pg/ml aprotinin). 1 0 0 pg protein per lane of samples were electrophoresed on 1 0 % SDS/PAGE gels. Samples were then transferred to poly-vinylidene difluoride (PVDF) membranes. After incubation in 5% non-fat milk in TBS for 1 hour at room temperature, membranes were incubated overnight at 4°C with rabbit polyclonal antibodies against either glutamate receptor NMDAR1 (NR1) (1:1000, Chemicon, Temecula, CA), glutamate receptor NMDAR2A (NR2A) (1:200, Chemicon, Temecula, CA) or glutamate receptor NMDAR2B (NR2B) (1:200, Chemicon, Temecula, CA). A horseradish peroxidase-conjugated anti-rabbit antibody (1:2000- 1:3000, Amersham Pharmacia Biotech, Piscataway, New Jersey) was used as 62 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. secondary antibody. A chemiluminescence substrate system (ECL, Amersham Pharmacia Biotech, Piscataway, New Jersey) was used to detect antibody binding. |3- actin antibody (1:5000, Amersham Pharmacia Biotech, Piscataway, New Jersey) binding was used to normalize optical densities of protein bands. The relative optical densities of protein bands were quantitated using a Chemilmager 5500 gel documentation system (Alpha Innotech Corporation, San Leandro, California). High-Performance Liquid Chromatography (HPLC) analysis of GSSG. For hippocampal GSSG tissue measurements, animals were perfused with 0.9 % NaCl containing 0.16 mg/ml heparin. Hippocampi were removed, weighed, frozen in liquid nitrogen and homogenized in 1 ml of 10% perchloric acid (PCA) with 1 mM bathophenanthrelinedisulfonic acid (BPDS) with or without an internal control, 1 pM GSSG. The homogenate was then sonicated at power 3 for 20 seconds (Sonic Dismembrator model 550, Fisher Scientific, Pittsburgh, PA) frozen and thawed. 0.5 ml of supernatant obtained after a 3-minute centrifugation at 15,000 x g was added to a tube containing 50 pi of 100 mM iodoacetic acid in 0.2 mM m-cresol purple solution. 0.5 ml of 2 M KOH - 2.4 M KHCO 3 was then added to the tube to raise the pH of the solution up to pH 8-9. After a 10-minute incubation in the dark at room temperature, 1 ml of 1% fluorodinitrobenzene was added to the solution. The solution was then mixed and stored at 4°C overnight to allow derivatization to completely occur. For extracellular GSSG measurements, at 11-13 DIV primary hippocampal culture neurons isolated form E l 8 animals were treated with 0 or 50 63 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. pM KA for 24 hours, the media collected then precipitated and derivatized using the same method as for hippocampi preparations. Hippocampal and extracellular GSSG levels were determined by the HPLC method of Fariss and Reed (Fariss and Reed, 1987) (LC-10ATVP pump, SCL-10AVP system control, Shimadzu) with a SPD- 10AVP UV detector and a SIL-10ADVP autosampler (Shimadzu) using a 3-amino propyl 5pm column (4.6 x 200 mm, Cel Associates, Inc., Pearland, TX). GSSG was identified by measuring absorbance at 365 nm at a sensitivity scale of 0.01. The amount of GSSG in each sample was calculated from a standard curve of GSSG prepared at the same time as the samples. The identity of GSSG peak was also confirmed by spiking the sample with GSSG standard. Brain slice preparation. GSHPx-1 transgenic and WT animals (1-2 months old) were anesthetized with halothane vapors and decapitated. Brains were quickly removed and placed in cold low-sodium sucrose substituted saline artificial cerebral spinal fluid (sucrose ACSF: 90 mM NaCl, 105 mM sucrose, 1.3 mM MgSC>4 , 3 mM KC1, 1.25 mM NaH2P 0 4 , 25 mM NaHC 0 3 , 2.4 mM CaCl2 , and 10 mM glucose, modified from Aghajanian and Rasmussen’s methods (Aghajanian and Rasmussen, 1989)). 200-400 pM coronal sections of the brains through hippocampi were cut with a vibroslicer (Camden Instruments, Loughborough, UK). The slices were then bathed in aerated (95% O, 5% C 0 2 normal ACSF (124 mM NaCl, 1.3 mM M gS04, 3 mM KC1, 1.25 mM NaH2PC > 4 , 25 mM NaHC0 3 , 2.4 mM CaCl2, and 10 mM glucose) for at least 1 64 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. hour prior to electrophysiological recording. In some experiments, MgSO.* was omitted to create Mg-free conditions. Electrophysiological recordings. Whole-cell recordings were obtained from neurons in brain slices bathed in normal ACSF using a fixed stage microscope (Axioscop, Zeiss, Germany) and water immersion lenses. Patch electrodes used for whole-cell recording were pulled on a Flaming/Brown p-87 micropipette puller (Sutter Instrument, Novato, California) and filled with internal solutions either i) 120 mM Cs-gluconate, 2 mM MgCl2, 0.5 mM EGTA, 10 mM HEPES, 10 mM TEA, 3 mM QX-314, 3 mM Na-ATP, pH 7.2, 270- 280 mOsm for voltage clamp readings or ii) 120 mM K-gluconate, 10 mM HEPES, 30 mM KC1, 0.2 mM EGTA, 3 mM Na ATP, 2 mM MgCl2, pH 7.2, 270-280 mOsm for current-clamp readings. The electrodes were positioned using a 3-axis MP-285 motorized micromanipulator (Sutter Instrument, Novato, California) for fine positioning and a mechanical manipulator, Newport MX 110 (Newport, Irvine, California) for coarse positioning. pCLAMP data acquisition software (Axon Instruments, Union City, California) and an Axopatch-ID patch clamp amplifier (Axon Instruments, Union City, California) were used to monitor electrode resistance in voltage clamp mode. Series of resistance were monitored throughout the experiment by measuring the instantaneous current response to a -10 mV voltage step from -70 mV. A gravity-fed array of inflow tubes of 0.58 mm inner diameter and an outflow tube attached to a vacuum reservoir provide solution flow which 65 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. allowed the slice to be continuously perfused with oxygenated solution during the search for neurons, rapid solution changes, and a small volume when the flow was stopped during recording for exposure to calcium channel antagonists. The ground electrode was consisted of a salt bridge constructed from glass electrode filled with agar. Drug Application. 100 pM NMD A were applied for 1 second to brain slices by using SF-77B Perfusion Fast Step (Warner Instrument, Hamden, Connecticut) Control responses were evoked for 6 minutes to ensure the stability of the response to NMDA and then 5 mM dithiothreitol (DTT) was perfused through the recording chamber. Responses were recorded for 6 minutes after the addition of DTT. DTT-mediated changes in the response to NMDA application were plotted over the time of DTT exposure as a percentage of the control NMDA response amplitude. For GSSG pressure application, 20 mM GSSG were applied by Pneumatic PicoPump (PV-820, World Precision Instruments, Saratosa, Florida), at holding membrane potentials of -60, or -75 with constant current or a holding membrane potential of -80 mV with small hyperpolarizing pulses of current (200 ms). For GSSG bath application, 20 mM GSSG was applied at a holding membrane potential of -70 mV. For GSSG or sucrose perfusion application, 20 mM GSSG or 20 mM sucrose was applied by Perfuse Fast Step System (Warner Instruments, Hamden, Connecticut) at a holding membrane potential of -55 mV. Voltage-clamp ramps were used to examine the voltage- 66 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. dependency of NMDA and GSSG responses. Cells were held at a holding potential of -70 mV and the membrane potential was slowly ramped over a one second period to a final peak potential of +40 mV. The membrane potential was then repolarized to -70 mV using a 200 ms ramp. Net NMDA and GSSG induced currents were obtained by subtracting currents obtained from ramps delivered before agonist application from currents obtained after agonist application. Acute isolation of CA3 neurons. Neurons were isolated using a procedure described earlier (Schumacher et al., 1998). The isolation procedure is as follows: Hippocampal slices were prepared in ice-cold solution containing 90 mM NaCl, 3 mM KC1, 2 mM CaCl2 , 2 mM MgSC>4 , 1 mM Na-pyruvate, 10 mM HEPES, 10 mM glucose, 105 mM sucrose, pH 7.4, 100 % O2 and transferred to the ASCF. Isolation of viable neurons has been possible up to 10 h after the preparation of slices. After an equilibration period of 30-45 min the individual slice were transferred to a polystyrene tube with 5 ml of saline containing 126 mM NaCl, 2.5 mM KC1, 2 mM CaCl2, 2 mM MgCl2, 1.25 mM NaH 2P 0 4, 26 mM PIPES, 10 mM glucose, pH 7.3, 100% O 2 . Pronase, 1.5 mg/ml (protease type XIV, Sigma) was added to the oxygenated medium. After an incubation period of 25-30 min at 32 C the slice was washed in the same ice-cold, enzyme-free saline. The CA3 region was dissected and triturated in 2 ml of ice-cold saline through fire polished Pasteur pipettes of decreasing apertures. The cells were then transferred to one compartment of two-compartment recording chamber and allowed to settle for 67 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 5-10 min. Neurons with pyramidal shaped soma were patched, lifted and transferred to the second small-volume (300-400 pi) compartment of recording chamber where they were perfused with the desired experimental solution. RESULTS GSHPx-1 transgenic mice displayed decreased H2O2 levels after KA treatment. < D > < D o ^ <N . £ 2 c d s o C/3 O o l-l O h S Q a & & ■ C/3 O S W T HM Figure 11. Brain synaptosomal H 2O 2 levels. H 2O 2 levels in synaptosomes isolated from the brains of KA-treated or saline-treated HM and WT animals; open bars, saline; filled bars, KA; HM, homozygous; WT, wild-type; * p = 0.01, n = 2-4. Despite having increased GSHPx activity, there was no difference in H 2O2 levels as assessed by the Amplex Red H2O2 assay kit in synaptosomes prepared from the brains of saline-treated HM vs. WT, 0.071283 ± 0.002145 mM/mg protein vs. 0.073237 ± 0.003565 mM/mg protein, respectively (Figure 11). However, upon KA treatment at a dosage of 35 mg/kg body weight, overexpression of GSHPx-1 was 68 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. capable of lowering H 2O2 levels in the brains of transgenic mice, 0.063689 ± 0.00232 mM/mg protein compared to 0.078837 ± 0.003403 in KA-treated WT brains. KA has been suggested to raise the levels of H 2O2 (Dykens et al., 1987), KA- treated WT brains displayed slightly higher H 2O2 levels than saline-treated WT brains but these values did not achieve significance. GSHPx-1 transgenic animals had decreased basal brain GSH/GSSG ratios. 80-, 0 6 0 - t-i O 0 0 oo O o 2 0 ■ 4 0 - HM HT W T Figure 12. Basal brain GSH/GSSG ratios in HM, HT and WT animals. GSH, reduced glutathione; GSSG, oxidized glutathione; HM, homozygous; HT, heterozygous; WT, wild-type; n = 3-5, * p = 0.05 HM vs. HT, ** p = 0.05 WT vs. HM and HT. Since GSHPx plays an important role in removing H 2O 2 via oxidation of GSH, having 4-6 fold increased levels of GSHPx-1 activity in GSHPx-1 transgenic mice might affect the basal redox status of the system leading to increased 69 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. vulnerability to oxidative insults. To test this hypothesis, basal brain redox status were measured in the transgenic mice and wild-type controls using a GSH/GSSG ratio assay kit. Interestingly, basal brain GSH/GSSG ratios were found to be significantly reduced in HM (37.81 ± 2.95) and HT (43.01 ± 2.40) GSHPx-1 transgenics compared to WT (62.49 ± 8.40) prior to KA administration (Figure 12). This is likely due to increased conversion of GSH to GSSG via elevated GSHPx levels (Mirochnitchenko et al., 1999). GSSG levels were elevated in GSHPx-1 transgenic mice. To investigate if the shift in redox status i.e. GSH/GSSG ratios, was caused by an increase in GSSG levels, hippocampal GSSG levels of untreated HM and WT were measured via HPLC measurements. Levels of cellular GSSG were found to be significantly elevated in the transgenics, 38.83 ± 5.10 pM/mg tissue in hippocampi isolated from HM, 31.44 ± 1.20 pM/mg tissue in those of HT compared to 23.24 ± 1.45 pM/mg tissue in those of wild-type animals (Figure 13). Though, the levels of GSSG were affected in transgenics, there was no detectable difference in GSH levels in transgenics vs. wild-type, 356.51 ± 54.40 pM/mg tissue in hippocampi isolated from HM, 322.62 ± 45.40 pM/mg tissue in those of HT compared to 313.595 ± 17.92 pM/mg tissue in those of WT likely due to large levels of GSH vs. GSSG. This leads to a decreased GSH/GSSG ratios in transgenic mice compared to WT animals. 70 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Figure 13. Hippocampal GSSG and GSH levels. HPLC measurements of hippocampal GSSG and GSH of GSHPx-1 transgenic mice and wild-type controls; GSSG, oxidized glutathione; GSH, glutathione; HM, homozygous; HT, heterozygous; WT, wild-type, (a) Hippocampal GSSG levels, n = 3, * p = 0.05. (b) Hippocampal GSH levels, n = 3. Decreases in G6PD and 6PGD activities, but not GR, may cause intracellular GSSG accumulation in GSHPx-1 transgenic mice. In the GSH metabolic pathway (Figure 2), GR plays an important role in reducing GSSG to GSH in the presence of NADPH. Two rate-limiting enzymes in the pentose phosphate pathway, G6 PD and 6 PGD, are required for NADP+ reduction to NADPH. To investigate if GSSG accumulation was caused by alterations in activities of any the enzymes involving in GSSG reduction, spectrophotometric assay kits were used to detect the activity levels of GR, G6 PD and 6 PGD in hippocampi of untreated GSHPx-1 transgenic vs. wild-type controls. 71 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ■ 3 120 - HM HT WT c '5 - 4 — > o o, o o £ to £ O o - V O a 10 8 6 4 2 0 HM HT WT C 10 - i c * 5 3 o W a O , 00 E £ ' - w ' O O c- v o 6 - 4- 2 - HM HT WT Figure 14. Alterations in activity levels of enzymes involving in glutathione metabolism in GSHPx-1 transgenic and wild-type brains; HM, homozygous; HT, heterozygous; WT, wild-type, (a) hippocampal glutathione reductase (GR) activity levels, n =3-6, * p = 0.05 HM vs. HT, ** p = 0.05 HT vs. WT. (b) hippocampal glucose-6 -phosphate dehydrogenase (G6 PD) activity levels, n =3-6, * p = 0.01 HM vs. WT. (c) hippocampal 6 -phosphogluconate dehydrogenase (6 PGD) activity levels, n = 3-6, * p = 0.01 HM vs. WT, ** p = 0.05 HT vs. WT. GR activity levels in HM (132.75 ± 2.97 mU/mg protein) and HT (119.27 ± 6.82 mU/mg protein) were slightly but significantly higher than in WT (97.86 ± 7.53 72 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. mU/mg protein) (35.66% and 21.88% increase respectively) (Figure 14a). However this is unlikely to compensate for the 400-600% increase in GSHPx activity in HT and HM transgenics, respectively (Figure 5). An approximately 10% decrease in G 6 PD activity levels was observed in HM (6.99 ± 0.14 mU/mg protein) compared to WT mice (7.83 ± 0.33 mU/mg protein). There was no significant difference between HT (7.80 ± 0.99 mU/mg protein) and WT animals (Figure 14b). Unlike G6 PD, activity levels of 6 PGD were decreased both in HM (3.34 ± 0.10 mU/mg protein) and HT (3.92 ± 0.88 mU/mg protein) compared to WT (4.94 ± 0.25 mU/mg protein) (32.37% and 20.80% respectively, Figure 14c). Decreased G6 PD and 6 PGD could conceivably act to limit NADPH generation required for conversion of GSSG to GSH despite the small compensatory GR elevation. GSSG levels in the media of KA-treated primary hippocampal neurons were elevated. It has been suggested that excessive intracellular GSSG is unidirectionally transported into extracellular space to prevent inhibition of intracellular enzymes caused by high levels of GSSG (Srivastava and Beutler, 1969). Activation of glutamate receptors, by either KA or NMDA, and malonate-induced oxidative stress have been reported to result in increased extrusion of GSSG from cultured cells and acute brain slices (Zeevalk et al., 1998; Wallin et al., 1999). In this study, overexpression of GSHPx-1 enzyme in the transgenic mice was shown to increase intracellular GSSG levels (Figure 13a). To investigate if KA-mediated oxidative 73 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. stress results in increased GSSG efflux in GSHPx-1 transgenic mice compared to wild-type animals, we measured GSSG levels via HPLC in the extracellular media from primary cultures derived from GSHPx-1 vs. WT hippocampi 24 hours following addition of 0 or 50 pM KA. We found GSSG levels to be significantly increased in the media from 50-pM KA-treated GSHPx-1 cells, 15.4940 ± 1.1536 mmol/mg protein vs. 11.3847 ± 1.2135 mmol/mg protein in the media from 50-pM KA-treated WT (Figure 15). There was no significant difference of GSSG levels in the media from 0-pM treated HM cultures, 10.6439 ± 1.0963 mmol/mg protein, and those from 0-pM treated WT cultures, 10.9939 ± 2.1397 mmol/mg protein. Figure 15. Extracellular GSSG levels. HPLC measurement of extracellular GSSG levels in the media of primary cultures derived from embryonic homozygous and wild-type hippocampi at 24 hours after 0 or 50 pM KA treatment. GSSG, oxidized glutathione; Open bars, wild-type; filled bars, homozygous, n = 3-5, * p = 0.01. 74 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. GSHPx-l-mediated adverse effect is independent of NMDA receptor redox state and NMDA receptor numbers. Based on our data, increased GSHPx activity levels in the transgenic animals appears to be responsible for an elevation in extracellular GSSG levels. Since, activation of NMDA receptors triggers seizures (Demarque et al., 2004) and GSSG has been reported to activate NMDA receptors (Leslie et al., 1992), we assessed the possible direct interaction of extracellular GSSG with the NMDA receptor. Previously, it has been reported that exogenous application of GSSG and to a lesser extent GSH can stimulate Ca2 + entry into dissociated neurons which is prevented by the NMDA receptor agonist MK801 (Leslie et al., 1992). GSSG has also been demonstrated to be capable of directly binding to NMDA receptors although the electrophysiological effects of this binding on receptor activity have not been directly tested (Janaky et al., 2000; Hermann et al., 2002). GSHPx-mediated elevation of intracellular GSSG levels coupled with KA-induced extrusion of GSSG may result in an increase in GSSG available to bind and directly activate surface NMDA receptors. This would explain the increase in seizures and CA3 hippocampal cell death in the GSHPx-1 transgenic animals following KA administration. An electrophysiological study, performed on acute hippocampal slices from HM vs. WT animals, demonstrated that the NMDA-evoked response in HM slices was approximately 3 times higher than in WT slices (min 2, 4, 6 following NMDA application, HM = 2451.00 ± 457.07 pA, 2496.64 ± 458.82 pA, 2582.59 ± 449.22 75 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. pA; WT = 845.10 ± 278.70 pA, 844.15 ± 239.18 pA, 799.22 ± 250.53 pA, respectively, Figure 16a). a < Q Z, 0 < D "O B 1 < 6.000 - I 5.000 - JS, 4,000 - | 3,000- & 2,000 - < D ’ 1,000 0 b < 250 - I Q o £ J - i 200- Z c o ID T3 o C m 150 - a O N® o« 0s 100- S < L > 73 c O o a 50- % - > s3 T 3 I n 0- Pi C - I - 2 i 2 — r- 4 6 -i— 8 T o" ~i 12 Time (min) 4 6 8 Time (min) 10 12 Figure 16. Absolute and relative dithiothreitol (DTT)-induced changes in NMDA- evoked response in CA3 neurons from WT and HM hippocampal slices. 100 pM NMDA was applied using the Perfusion Fast Step system. At minute 6 , 5 mM DTT was added via Fast Step. Responses were recorded for 6 minutes after the addition of DTT; NMDAR, NMDA receptor, (a) Absolute amplitude NMDA receptor response values, (b) Relative amplitude NMDA receptor response. 76 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. The NMDA receptor has been reported to contain a redox modulatory site which can effects its activity (Aizenman et al., 1990; Gilbert et al., 1991). Oxidation of the NR1 subunit of the NMDA receptor has been suggested to cause decreased function of the receptor leading to increased resistance to KA-induced seizure and neuronal cell death in GSHPx-1 knockout mice (Jiang et al., 2000). To investigate if a change in redox status of NMDA receptor was responsible for the increase in NMDA-evoked activity observed in the GSHPx-1 transgenic hippocampus, 5 mM DTT was applied to the slice cultures after minute 6 . No difference in relative DTT- induced NMDA-evoked activity was observed between WT and HM cultures suggesting that the GSHPx-mediated effects is independent of NMDA receptor redox state (minute 8 , 10, 12; HM =1.38 ± 0.04, 1.63 ± 0.04, 1.77 ± 0.07 pA; WT=1.27 ± 0.03,1.47 ± 0.08,1.64 ± 0.17, respectively) (Figure 16b). In addition, results from immunoblotting assays demonstrated no increase in numbers of NMDA receptor subunits expressed in the hippocampus including NMDAR1 (NR1), NMDAR2A (NR2A) nor NMDAR2B (NR2B) (Figure 17) suggesting that the NMDA-evoked response is also separate from alterations in NMDA receptor numbers. Indeed, there appears to be a decrease in NR1 receptor subunit protein levels in the GSHPx-1 transgenics; this would be expected to potentially result in decreased rather than increased NMDA receptor activity. 77 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. a NRl P-actin HM HT WT NR2A P-actin O f : ♦ . HM HT WT HM HT WT NR2B p-actin b 4 - 0' 3.0' | 2. 0 . 1 .0 - ()■ d 2 - 0' 1.5 ■ < S i - o ■ A 0.5 • O' f 2.0- 1.5- 0 3 at 1.0- A 0.5- 0 ■ * HM HT WT HM HT WT HM HT WT Figure 17. Western blot analysis of NMDA receptor subunits, (a) Western blot of hippocampal NMDA receptor subunit NRl and (b) its corresponding arbitrary units relative to (3-actin band densities, (c) and (d), NR2A. (e) and (f), NR2B; HM, homozygous; HT, heterozygous; WT, wild-type, * p = 0.05, HM vs. WT. Extracellular GSSG induces depolarization in CA3 neurons. To investigate the possibility that high levels of extracellular GSSG may directly activate CA3 neurons, GSSG was directly applied to individual neurons in acute hippocampal brain slices from WT animals. Application of 10-20 mM GSSG via pressure pipette and recording of electrophysiological responses by whole-cell current clamp revealed large GSSG-evoked depolarizations generating a barrage of action potentials reminiscent of the “paroxysmal depolarizing shift” seen in models of epilepsy (Johnston and Brown, 1984; Bradford, 1995) (Figure 18a). 78 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. a G S SG 20 m M G SSG 20 m M 20 mV -75 2 sec -70 1 min GSSG 20 mM d -55 20 mV 2 sec Sucrose 20 mM 20 mv|___ -55 2 sec Figure 18. GSSG-evoked electrophysiological responses in WT CA3 neurons, (a) Response of CA3 neuron to pressure application of 20 mM GSSG (Vm = -75 mV). (b) Response of CA3 neuron to slow bath application of 20 mM GSSG. GSSG (20 mM) produced a depolarization and an associated barrage of action potentials (top trace, Vm = -60 mV). (c) Pressure application of 20 mM GSSG generates a depolarization and action potentials while application of equimolar sucrose (d) did not produce a response (Vm = -55mV). To eliminate the possibility of depolarization being generated by pressure pipette application, GSSG was also applied via bath perfusion. Similar results were obtained, i.e. a slow depolarization associated with a series of action potentials (Figure 18b). The effects of GSSG application could not be explained via osmotic effects as equimolar sucrose application showed no response (Figure 18c-d). 79 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. GSSG directly activates NMDA receptors. The bursting behavior induced by GSSG application suggested GSSG could be acting by directly activating NMDA receptors. To test this hypothesis we compared response of direct application of GSSG to CA3 neurons. Individual CA3 neurons were voltage-clamped at a holding potential of -70 mV and exposed to a 1 second depolarizing ramp which peaked at a membrane potential of +40 mV and then returned to -70 mV during a 200 millisecond (ms) repolarizing ramp. The neurons were then exposed to GSSG and the ramp was re-delivered. Subtraction of the control ramp from the ramp obtained after GSSG exposure revealed a voltage- dependent GSSG-induced current which looked similar to that induced by NMDA (Figure 19a-b). Application of 100 pM 5-aminophosphonovaleric acid (AP-5) was able to block subsequent GSSG-evoked responses (Figure 19b). Similarly, AP-5 blocked responses to NMDA application in the same cell (Figure 19a). It is possible that GSSG could cause a release of glutamate and thus not be directly activating NMDA receptors. To eliminate this possibility, we examined the response of CA3 neurons to GSSG application in neurons acutely isolated from surrounding hippocampal tissue. This method allows recording from single CA3 neurons (cell body and small primary dendrite) in total isolation from tissue (other cells axons, synapses, dendrites, cell bodies). We found GSSG also evoked an inward current in these isolated neurons (Figure 19c). 80 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. I (pA) a NM DA 1 sec -70 200 -80 -40 AP-5 -400- b 100- GSSG AP-5 '100- c GSSG 20 mM 50 pA 1 sec Figure 19. GSSG induced response acts via direct NMDA receptor activation in CA3 neurons from WT mice. Response to application of (a) 100 pM NMDA alone (NMDA) or 100 pM NMDA after application of 100 pM AP-5 (AP-5) (b) Response to 20 mM GSSG alone (GSSG) or 20 mM GSSG after an application of 100 pM AP- 5 (AP-5) (Perfusion Fast Step) (holding potential = -70 mV, 1 second depolarizing ramp, peaked at +40 mV and returned to -70 mV during a 200 ms repolarizing ramp), (c) Response of an acutely isolated CA3 neuron to application of 20 mM GSSG (holding potential = -40 mV). 81 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. DISCUSSION In this study, we have obtained evidence suggesting that a shift in redox status (GSH/GSSG) mediated by overexpression of GSHPx-1 can lead to GSSG efflux which in turn may lead to direct activation of extracellular NMDA receptors. We demonstrated that mice with increased GSHPx activity levels displayed increased intracellular GSSG levels. This is possibly due to decreased activities of G6PD and 6PGD, rate-limiting enzymes required for generation of NADPH which is necessary for reduction of GSSG into GSH by GR. The increase in GSSG levels leads to a shift in the GSH/GSSG ratio and subsequent GSSG efflux. Exogenous application of GSSG has been shown to be able to directly activate neuronal NMDA receptors isolated from the hippocampal CA3 area of WT animals. Moreover, the NMDA-evoke response of CA3 neurons in hippocampal brain slices isolated from transgenic animals was increased independently of either the redox status or the numbers of NMDA receptors. Taken together, these data suggest that the increased susceptibility of GSHPx-1 transgenic animals to KA toxicity observed in the previous study is due to increased NMDA receptor function caused by direct activation by extracellular GSSG. 82 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Redox status alteration may cause the increased vulnerability of GSHPx-1 transgenic animals to KA toxicity. In the current study, increased GSHPx activity was found to induce increased intracellular GSSG levels leading to a shift in redox status and decreased GSH/GSSG. The glutathione (GSH/GSSG) redox status is important for maintaining a reducing environment in cells and tissues. Any alterations in the ratio of GSSG to GSH could cause a shift in the cellular redox environment (Schafer and Buettner, 2001; Filomeni et al., 2002a). Several pieces of evidence suggest that alterations in the redox state can cause adverse effects. This has been suggested to be specifically due to either GSH depletion or increases in GSSG levels. For example, depletion of GSH via either the GSH synthesizing inhibitor buthionine sulfoxamine (BSO) or diethyl maleate (DEM) in Bcl-2 overexpressing HL60 cells leads to increased ROS production and selective cell death (Armstrong and Jones, 2002). Similar reductions in GSH via 2-cyclohexen-l-one (CHX) has also been reported to enhance activator protein-1 (AP-1) DNA binding in the murine hippocampus and to result in more severe KA-induced seizures (Ogita et al., 2001). Elevated GSSG has also been reported to induce detrimental effects. Exogenous GSSG application, for example, has been shown to result in apoptosis in a promonocytic cell U937 by altering the intracellular redox environment resulting in activation of the p38 mitogen protein pathway (Filomeni et al., 2002b). Biroccio and colleagues demonstrated that down- regulation of c-myc also induces program cell death in melanoma cells via an accumulation of intracellular GSSG (Biroccio et al., 2002). Cereser and coworkers 83 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. demonstrated that thiram, a dithiocarbamate fungicide, induces increased GSSG levels and lowers cell viability in human fibroblast cultures (Cereser et al., 2001). Decreased G6PD and 6PGD activity levels and increased GSHPx/GR ratio may cause an increase in intracellular GSSG in GSHPx-1 transgenic animals. In the GSH metabolic pathway (Figure 2), to prevent the toxicity of GSSG due to its accumulation (Meister and Anderson, 1983) and to maintain the GSH pool, GR plays an important role in reducing GSSG to GSH in the presence of NADPH which is generated by G6PD and 6PGD, rate-limiting enzymes in the pentose phosphate pathway (Mehta et al., 2000; Grabowska and Chelstowska, 2003; Leopold et al., 2003; Spolarics et al., 2004). In the same line of GSHPx-1 transgenic mice, it has been reported that the levels of GR activity in whole lens and lens epithelium were not significantly different from those of wild-type animals (Spector et al., 1996). However, in the current study, GSHPx-1 transgenic mice displayed increased hippocampal GR activity levels compared to wild-type animals. Despite the compensatory increase in GR activity, the ratio of GSHPx/GR in the transgenic animals was still 4.5 fold higher than that of wild-type animals. It has been suggested that excess GSHPx over GR may cause an inability of GR to reduce GSSG and thereby maintain GSH levels (Spector et al., 1996). Moreover, decreased G6PD and 6PGD activities levels observed in the transgenics may also affect the reduction of GSSG to GSH 84 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. (Salvemini et al., 1999; Mehta et al., 2000; Leopold et al., 2003) leading to an accumulation of intracellular GSSG. Extracellular GSSG may directly activate the NMDA receptor In the current study, increased GSHPx activity was found to induce elevated intracellular GSSG levels and eventually increased extracellular GSSG levels in response to KA administration in vitro. Active transport of GSSG has been proposed to be an emergency mechanism to protect the system from GSSG toxicity (Meister and Anderson, 1983). Though the active transport of GSSG via its efflux was discovered a few decades ago (Srivastava and Beutler, 1969), its molecular mechanism was not completely understood. Recently, it has been proposed that the transporters for GSSG efflux are a GSSG-Mg2 + -ATPase (Kondo et al., 1987) and/or the multidrug resistance-associated protein (MRP) (Sharma et al., 2000; Hirrlinger et al., 2001). GSSG efflux has been suggested to be correlated with intracellular GSSG levels; the higher the intracellular GSSG levels, the higher the GSSG efflux and subsequent extracellular GSSG levels (Akerboom et al., 1982a; Akerboom et al., 1982b). In addition, in tumor cells extracellular GSSG has been suggested to be capable of altering intracellular redox status, through depletion of GSH and induction of cell death via oxidization of cysteine residues of transmembrane pro- apoptotic proteins leading to an activation of the mitochondrial apoptotic pathway (Filomeni et al., 2002a; Filomeni et al., 2002b). 85 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. GSSG has been demonstrated to have direct electrophysiological effects via activation of NMDA receptors. Several groups have reported that GSSG either inhibited (Janaky et al., 1993) or decreased NMDA receptor responses (Gilbert et al., 1991; Sucher and Lipton, 1991; Varga et al., 1997) apparently via oxidation of the NMDA receptor. In this study, hippocampal slices prepared from HM animals displayed approximately 3 times higher levels of NMDA-evoked electrophysiological responses than WT hippocampal slices, independent of NMDA receptor numbers. To investigate if redox status alteration was responsible for the increased responses of HM slices, DTT, a thiol reducing agent, was applied to the hippocampal slices isolated from both HM and WT animals. If the NMDA receptors of WT were in a more oxidized state, DTT treatment should alter the NMDA-evoke response of WT slices even more than the response of DTT-treated HM slices. However, alterations in basal NMDA-evoked electrophysiological responses in GSHPx transgenic hippocampal neurons in our study did not appear to be explainable by redox effects since there was no significant difference in relative NMDA-evoked responses of hippocampal slices prepared from HM and WT animals following DTT application (Jiang et al., 2000). It has been previously demonstrated that GSSG binds to NMDA receptors and increases intracellular Ca2 + via NMDA receptor activation which can be blocked by MK801, an NMDA inhibitor (Leslie et al., 1992). In this study, we explored the possible relationship between effluxed GSSG and the NMDA receptor. We found that exogenous application of GSSG elicited bursts of neuronal activity similar to 86 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. those seen in electrophysiological models of epilepsy (Wyler et al., 1975). GSSG responses were blocked by the selective NMDA receptor antagonist AP-5. These data support a role for elevated GSSG in the initiation of seizures through direct activation of NMDA receptors. Our study suggests that alterations in the GSH/GSSG ratio via increased GSHPx expression results in elevated extracellular GSSG levels which can directly interact with and activate NMDA receptors. This in turn appears to result in increased seizure activity and CA3 hippocampal neuronal cell death associated with KA administration. This not only explains increased seizure and cell death in the GSHPx-1 transgenics following KA administration, but also could have larger global implications in terms the effects of the glutathione redox state in seizure-mediated cell death. 87 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. CHAPTER FOUR CONCLUSIONS Keeping the intracellular redox environment in a reduced state is known to be essential for cell survival. Changes in GSH and/or GSSG levels, the major redox couple within cells, would cause a shift in the intracellular redox status leading to vulnerability to oxidative stimuli. There are several factors that can cause an imbalance in the GSH/GSSG ratio. For example, oxidative stress-mediated GSH depletion and alterations in GSH-regenerating cycle enzymes activity levels i.e. GR and GSHPx, could have a significant impact on cellular redox status. If an imbalance in the redox system persists due to prolonged oxidative stress, the cells would be no longer able to cope with subsequent oxidative insults resulting in irreversible cell damage and death. Reducing agents such as vitamin C and vitamin E and antioxidant enzymes such as the 02*'-removing enzyme, Cu/ZnSOD, and the H202-scavenging enzyme, catalase, have been shown to be protective against oxidative stress by removing ROS from the cellular system. In this study, by using a genetically engineered mouse model that overexpresses the murine gene for the H 2O 2- scavenging enzyme GSHPx-1, one of the major cellular antioxidant enzymes, we investigated the role of this enzyme in oxidative stress-mediated KA toxicity. 88 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. In the current study, murine GSHPx-1 transgenic mice which displayed 4-6 fold increases in levels of hippocampal GSHPx activity levels, were systemically administered with KA at a dosage of 35 mg/kg body weight. Following KA administration, rather than being resistant to KA toxicity as we had originally predicted, transgenic mice displayed increased seizure activity compared to KA- treated wild-type animals. Twenty four hours following KA treatment, transgenic mice also displayed increased neuronal cell damage especially in the hippocampal CA3 and CA1 regions with less damage in the CA2 and CA4 regions of the hippocampus compared to those of KA-treated wild-type animals. In subsequent in vitro studies, we also observed an increased sensitivity to KA toxicity in primary hippocampal cultures isolated from transgenic mice compared to KA-treated cultures isolated from wild-type animals. Given these unexpected results, we proposed to investigate whether the increase in GSHPx activity may play an important part in the increased sensitivity of the transgenic mice to KA toxicity by introducing an imbalance in the cellular GSH/GSSG ratio. Our observations suggest that despite a small compensatory increase in GR activity, an overwhelming increase in GSHPx-1 enzyme activity and decreased activity in two other important GSH-regenerating enzymes, G6PD and 6PGD, may result in an accumulation of intracellular GSSG leading to an imbalanced GSH/GSSG ratio. In our in vitro study, increased GSSG levels were detected in the medium of KA-treated primary hippocampal cultures derived from transgenic animals. This finding is in line with other studies that suggest to maintain 89 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. the reducing cellular redox environment in response to oxidative stress and to prevent the toxicity of intracellular GSSG accumulation, GSSG must actively be transported outwards into the extracellular space. Since seizure and neurodegeneration have been previously shown to be associated with activation of the NMD A receptor, we investigated the receptor’s activity in transgenic vs. WT mice. In electrophysiological studies, we found that hippocampal slices isolated from transgenic mice displayed increased basal NMDA- evoked response compared to those isolated from wild-type animals, independent of the NMDA receptor redox state or NMDA receptor numbers. Studies from other laboratories have suggested that extracellular GSSG is capable of binding and modulating NMDA receptor activity. In our study, we demonstrated that exogenous GSSG can directly activate the NMDA receptors of wild-type animals. Taken together, our observations strongly suggest that alterations in the redox state of the glutathione system in GSHPx-1 transgenic mice resulting in a shift of the cellular redox status results in elevated extracellular GSSG levels which, in turn, directly activate NMDA receptors resulting in subsequent seizure activity and neurodegeneration. Our findings also suggest that therapeutic strategies in counteracting oxidative stress may need to be re-examined since an overwhelming dosage of an antioxidant, instead of being protective, could cause undesirable effects. 90 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. REFERENCES Aghajanian, G. K., and Rasmussen, K. (1989). Intracellular studies in the facial nucleus illustrating a simple new method for obtaining viable motoneurons in adult rat brain slices. Synapse 3(4), 331-8. Aizenman, E., Hartnett, K. A., and Reynolds, I. J. (1990). 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Asset Metadata

Creator Boonplueang, Rapee (author)

Core Title Increased susceptibility of glutathione peroxidase-1 transgenic mice to kainic acid-related seizure activity and hippocampal neuronal cell death due to direct activation of the NMDA receptor by GSSG

Contributor Digitized by ProQuest (provenance)

School Graduate School

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag biology, molecular,biology, neuroscience,OAI-PMH Harvest

Language English

Advisor Andersen, Julie K. (committee chair), Petruska, John (committee chair), Arnold, Donald (committee member), Pike, Christian (committee member)

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c16-565105

Unique identifier UC11340230

Identifier 3155383.pdf (filename),usctheses-c16-565105 (legacy record id)

Legacy Identifier 3155383.pdf

Dmrecord 565105

Document Type Dissertation

Rights Boonplueang, Rapee

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the au...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus, Los Angeles, California 90089, USA