New Strategies in Stroke Intervention: Ionic Transporters, Pumps, and New Channels (Contemporary Neuroscience)

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Contemporary Neuroscience

For further volumes: http://www.springer.com/series/7626

Lucio Annunziato Editor

New Strategies in Stroke Intervention Ionic Transporters, Pumps, and New Channels

Editor Lucio Annunziato Department of Neuroscience School of Medicine ‘‘Federico II’’ University of Naples Via Sergio Pansini, 5 80131 Napoli Italy [email protected]

ISBN 978-1-60761-279-7 e-ISBN 978-1-60761-280-3 DOI 10.1007/978-1-60761-280-3 Library of Congress Control Number: 2009932562 # Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer ScienceþBusiness Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper springer.com

Preface

Stroke is a leading cause of serious, long-term disability in the world. Every 45 seconds someone is affected by stroke and on average every 3–4 min someone dies of focal ischemia. In fact, 15 million people suffer from stroke worldwide each year. Of these, five million die and another five million are permanently disabled. Stroke ranks among the third most common cause of death after heart disease and cancer. In developed countries, the incidence of stroke is declining – largely because of the efforts to lower blood pressure and reduce smoking. However, given that age is one of the most substantiated risk factors for stroke, the aging of the world population implies a growing number of people at risk. The mean lifetime cost of a single ischemic stroke in the US is estimated at $140,048, including impatient care, rehabilitation, and follow-up care for longlasting deficits. The estimated direct and indirect costs of stroke for 2009 are $ 68.9 billion in the United States and $ 32.3 billion in the European Union countries. Stroke is caused by a blood clot that lodges in the brain and reduces oxygen supply to critical tissue. Two general therapeutic approaches are available: the first strategy is aimed at reducing the failure of arterial oxygen and glucose delivery to the local brain tissue by performing a thrombolysis of the arterial thrombus within few hours from the onset of symptoms and to reduce the tissue back-pressure occurring when failure of the blood–brain-barrier causes vasogenic edema. The second, not yet validated, therapeutical strategy is to exert a neuroprotective action on the surviving brain ischemic tissue. Despite the great effort generated in the attempt to identify new pharmacological treatments, the only current drug therapy used in the treatment of stroke belongs to the first thrombolytic strategy. However, this pharmacological approach has a narrow window of time for therapeutic application and dose-limiting adverse effects. In fact, the clot busting drug recombinant tPA can save lives and diminish disability, but its use is limited to about 3–4% of all stroke patients. In part, this is because the drug must be administered within 3 hours after the beginning of stroke symptomatology. Furthermore, tPA has also risky side effects such as haemorrhage, edema, and

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potential neurotoxic effects have been hypothesized by some experimental studies. Rupture or blockade of a blood vessel in the brain causes rapid cell death in the core of the injured region and triggers mechanisms in the surrounding area – the penumbra – that lead among several mediators to changes in the concentrations of several ions such as intracellular Ca2+, Na+, H+, K+, and radicals such as reactive oxygen species (ROS) and reactive nitrogen species (RNS). All these transductional factors might initiate cell death. In particular, it is widely accepted that a critical factor in determining neuronal death during cerebral ischemia is the progressive accumulation of intracellular Na+ ions, which can precipitate necrosis and apoptosis of vulnerable neurons. Whereas the detrimental action of [Na+]i increase is attributable to both cell swelling and microtubular disorganization – phenomena that lead to cell necrosis – a change in Ca2+,Na+, K+, H+ ions has been shown to be a key factor in ischemic brain damage, for it modulates several death pathways, including oxidative and nitrosative stress, mitochondrial dysfunction, protease activation, and apoptosis. Since Olney’s seminal work firstly suggested that excitatory amino acids could elicit neurotoxicity, a large amount of work has been accumulated showing that glutamate extracellular concentrations briskly rise during acute brain injury, thus triggering an influx of Ca2+ and Na+ ions into neurons through ionotropic glutamate receptor subtypes. This evidence has led to the elaboration of the paradigm of glutamate excitotoxicity that explains ischemic neuronal cell death as a mere consequence of Na+ and Ca2+ influx through glutamate receptors. Although this theory has been guiding basic research in the field of neurodegeneration for almost three decades, more recently it has become the object of serious criticism and reassessment. What has aroused such skepticism among researchers has been the fact that although first, second, and third generation glutamate receptor antagonists have long yielded promising results in animal models of brain ischemia, they have failed to elicit a neuroprotective action in stroke and traumatic brain injury in humans. Therefore, the theory of excitotoxicity, though a fascinating paradigm, can only explain some of the events occurring in the acute phase of anoxic insult but cannot be seen as a major target for developing new therapeutic avenues for brain ischemia. In the last decade, several seminal experimental works are markedly changing the scenario in this field. In fact, it has been shown that some integral plasma-membrane proteins, involved in the control of Ca2+, Na+, K+, H+ ions influx or efflux and, therefore, responsible for maintaining the homeostasis of these four cations, might function as crucial players in the brain ischemic process. Indeed, these proteins, by regulating Ca2+, Na+, K+, H+ homeostasis, may provide the molecular basis underlying glutamateindependent Ca2+ overload mechanisms in neuronal ischemic cell death and, most importantly, may represent more suitable molecular targets for therapeutic intervention. Targeting these mechanisms is a promising route for the development of innovative therapies for stroke treatments.

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The main goal of this book is to provide readers involved in basic and clinical research with an overview of glutamate receptor-independent channels, pumps, and ionic exchangers regulating Na+ and Ca2+ homeostasis, involved in the pathophysiology of the ischemic damage and potentially targetable in the attempt to develop a new therapy in stroke intervention. Twelve chapters are included in this book. Chapter ‘‘Basis of ionic dysregulation in cerebral ischemia’’ describes the basis of ionic dysregulation in cerebral ischemia. Chapter ‘‘Why have ionotropic and metabotropic glutamate antagonists failed in stroke therapy?’’ summarizes the reasons of the failure of the glutamate theory. Chapters ‘‘Mitochondrial channels as potential target for a pharmacological strategies in brain ischemia’’ and ‘‘Endoplasmic reticulum calcium homeostasis and neuronal pathophysiology of stroke’’ deal with the role of the two intracellular organelles, mitochondria and endoplasmic reticulum, in the pathophysiology of the ischemic event. The subsequent four chapters of the book, chapters ‘‘The Naþ/Ca2þ exchanger: a target for therapeutic intervention in cerebral ischemia’’, ‘‘The ‘loop’ diuretic drug bumetanide-sensitive Naþ-Kþ-Cl– cotransporter in cerebral ischemia’’, ‘‘The Naþ/Hþ exchanger: A target for therapeutic intervention in cerebral ischemia’’, and ‘‘The Naþ/Kþ-ATPase as a drug target for ischemic stroke’’, present data regarding the potential involvement of ionic transporters responsible for the control of neuronal ionic homeostasis in the development of stroke lesion. In particular, the following transporters are described: Naþ/ Ca2þ exchangers, NCXs, Na+/K+/Ca2+/Cl– cotransporters, NKCCs, Na+/ H+ Exchangers, NHEs, and Naþ/Kþ ATPase. In the second part of the book, chapters ‘‘Acid-sensing ion channels (ASICs): New targets in stroke treatment’’, ‘‘Role of TRPM7 in ischemic CNS injury’’, ‘‘Subtypes of voltage-gated Ca2+ channels and ischemic brain injury’’, and ‘‘The diverse roles of K+ channels in brain ischemia’’, it is described the role of important membrane ionic channels such as Acid-Sensing Ionic Channels, ASIC, the Transient Receptor Potential Channels, TRPC; Voltage Operated Ca2+ Channels, VOCC and K+ channels. Each chapter dedicated to most of the molecular targets involved in the maintenance of ionic homeostasis describes in a coordinated way the relevant information related to: (1) gene structure, (2) structural features, (3) molecular biology, (4) cellular and tissue distribution, (5) biophysical and electrophysiological properties, (6) receptorial, transcriptional and transductional regulatory mechanisms, (7) physiological properties, (8) pathophysiological relevance in stroke, (9) pharmacological modulation, (10) preliminary clinical trials, and (11) therapeutic perspectives. This book written by the most authoritative scientists in the field should provide to the readers the most relevant information to understand the role played by the most recently discovered mechanisms involved in the pathophysiology of stroke thus providing the characterization of new pharmacological avenues for the cure of this relevant neurological disease. Naples, Italy

Lucio Annunziato

Acknowledgments

We thank Dr. Paola Merolla for editorial revision, Ms Giuliana Pellegrini of the Art Studio Design ‘‘Ciotola’’ for revising all the figures of the book, and Dr. Fabrizio Esposito, Associate Researcher at the Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, for providing us with the fMRI image of human brain used in the cover of the present book.

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Contents

Basis of Ionic Dysregulation in Cerebral Ischemia. . . . . . . . . . . . . . . . . . . Thiruma V. Arumugam, Eitan Okun, and Mark P. Mattson

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Why have Ionotropic and Metabotropic Glutamate Antagonists Failed in Stroke Therapy? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gianfranco Di Renzo, Giuseppe Pignataro, and Lucio Annunziato

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Mitochondrial Channels as Potential Targets for Pharmacological Strategies in Brain Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rosemary H. Milton and Michael R. Duchen

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Endoplasmic Reticulum Calcium Homeostasis and Neuronal Pathophysiology of Stroke. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alexei Verkhratsky

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The Naþ/Ca2þ Exchanger: A Target for Therapeutic Intervention in Cerebral Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lucio Annunziato, Pasquale Molinaro, Agnese Secondo, Anna Pannaccione, Antonella Scorziello, Giuseppe Pignataro, Ornella Cuomo, Rossana Sirabella, Francesca Boscia, Alessandra Spinali, and Gianfranco Di Renzo The ‘‘Loop’’ Diuretic Drug Bumetanide-Sensitive Naþ-Kþ-Cl– Cotransporter in Cerebral Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dandan Sun

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The Naþ/Hþ Exchanger: A Target for Therapeutic Intervention in Cerebral Ischemia. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Jin Xue and Gabriel G. Haddad The Naþ/Kþ-ATPase as a Drug Target for Ischemic Stroke. . . . . . . . . . . Melissa A. Gottron and Donald C. Lo

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Acid-Sensing Ion Channels (ASICs): New Targets in Stroke Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Giuseppe Pignataro Role of TRPM7 in Ischemic CNS Injury . . . . . . . . . . . . . . . . . . . . . . . . . . Michael F. Jackson, Hong-Shuo Sun, Michael Tymianski and John F. MacDonald

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Subtypes of Voltage-Gated Ca2þ Channels and Ischemic Brain Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Soon-Tae Lee, Daejong Jeon, and Kon Chu

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The Diverse Roles of Kþ Channels in Brain Ischemia . . . . . . . . . . . . . . . . Hiroaki Misonou and James S. Trimmer

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Clinical Trials with Drugs Targeting Ionic Channels, Antiporters, and Pumps in Ischemic Stroke . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alessandra Spinali, Giuseppe Pignataro, Gianfranco Di Renzo, and Lucio Annunziato Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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List of Contributors

Lucio Annunziato Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Thiruma V. Arumugam Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX 79106, USA, [email protected] Francesca Boscia Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Kon Chu Stroke & Stem Cell Laboratory, Department of Neurology, Clinical Research Institute, Comprehensive Epilepsy Center, Seoul National University Hospital, Program in Neuroscience, Seoul National University, Seoul, 110-744, South Korea, [email protected] Ornella Cuomo Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Gianfranco Di Renzo Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Michael R. Duchen Department of Physiology, University College London, London WC1E 6BT, UK, m.duchen @ucl.ac.uk Melissa A. Gottron Center for Drug Discovery and Department of Neurobiology, Duke University Medical Center, Durham, NC 27710, USA, [email protected] Gabriel G. Haddad Departments of Pediatrics and Neuroscience, University of California San Diego, La Jolla, CA 92093, USA, and Rady Children’s Hospital, San Diego, CA 92123, USA, [email protected] Michael F. Jackson Robarts Research Institute, Molecular Brain Research Group, University of Western Ontario, London, ON N6A 5K8, Canada, [email protected] xiii

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List of Contributors

Daejong Jeon Stroke & Stem Cell Laboratory, Department of Neurology, Clinical Research Institute, Comprehensive Epilepsy Center, Seoul National University Hospital, Program in Neuroscience, Seoul National University, Seoul, 110-744, South Korea, [email protected] Soon-Tae Lee Stroke & Stem Cell Laboratory, Department of Neurology, Clinical Research Institute, Comprehensive Epilepsy Center, Seoul National University Hospital, Program in Neuroscience, Seoul National University, Seoul, 110-744, South Korea, [email protected] Donald C. Lo Center for Drug Discovery and Department of Neurobiology, Duke University Medical Center, Durham, NC 27710, USA, [email protected] John F. MacDonald Robarts Research Institute, Molecular Brain Research Group, University of Western Ontario, London, ON N6A 5K8, Canada, and Department of Physiology, University of Toronto, King’s College Circle, Toronto, ON M5S 1A8, Canada, [email protected] Mark P. Mattson Laboratory of Neurosciences, National Institute on Aging Intramural Research Program, Baltimore, MD 21224, USA, [email protected] Rosemary H. Milton Department of Physiology, University College London, Gower Street, London WC1E 6BT, UK, [email protected] Hiroaki Misonou Department of Neural and Pain Sciences, Program in Neuroscience, Dental School, University of Maryland, Baltimore, MD 21201, USA, [email protected] Pasquale Molinaro Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Eitan Okun Laboratory of Neurosciences, National Institute on Aging Intramural Research Program, Baltimore, MD 21224, USA, [email protected] Anna Pannaccione Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Giuseppe Pignataro Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Antonella Scorziello Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected]

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Agnese Secondo Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Alessandra Spinali Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Rossana Sirabella Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Italy, [email protected] Dandan Sun Department of Neurological Surgery, University of Wisconsin School of Medicine and Public Health, Madison, WI 53705, USA, [email protected] Hong-Shuo Sun Department of Physiology, University of Toronto, King’s College Circle, Toronto, ON M5S 1A8, Canada, and Toronto Western Hospital Research Institute, Toronto, ON M5T 2S8, Canada, [email protected] James S. Trimmer Department of Neurobiology, Physiology, and Behavior, College of Biological Sciences; and Department of Physiology and Membrane Biology, School of Medicine, University of California, Davis, CA 95616, USA [email protected] Michael Tymianski Department of Physiology, University of Toronto, King’s College Circle, Toronto, ON M5S 1A8, Canada, and Toronto Western Hospital Research Institute, Toronto, ON M5T 2S8, Canada, [email protected] Alexei Verkhratsky Faculty of Life Sciences, The University of Manchester, Manchester M13 9PT, UK and Institute of Experimental Medicine, ASCR, 142 20 Prague 4, Czech Republic, [email protected] Jin Xue Departments of Pediatrics, University of California San Diego, La Jolla, CA 92093, US, [email protected]

Basis of Ionic Dysregulation in Cerebral Ischemia Thiruma V. Arumugam, Eitan Okun, and Mark P. Mattson

1 Introduction Stroke causes 9% of all deaths around the world and is the second most common cause of death after ischemic heart disease. Enhancing recovery from stroke and limiting ischemic damage are major goals to decrease stroke morbidity and mortality [15]. Brain tissue is extremely sensitive to oxygen and glucose deprivation, and even brief ischemia can initiate a complex sequence of events that ultimately culminates in cellular death. Studies performed during the past 30 years have identified several key pathophysiological events that lead to ischemic neuronal degeneration. The pathophysiological processes in stroke are complex and involve disruption of the blood–brain barrier (BBB), energy failure, loss of cell ion homeostasis, acidosis, increased intracellular calcium levels, excitotoxicity, free radical-mediated toxicity, generation of arachidonic acid products, cytokine-mediated apoptosis, activation of glial cells, and infiltration of leukocytes [39, 5, 6]. In focal cerebral ischemia, ischemic tissue is divided into an infarction core and penumbra. The core is the area of the brain where blood flow is reduced below 10–20% of its normal levels [55]. In the core, rapid anoxic depolarization causes immediate loss of membrane potential followed by the loss of membrane integrity and rapid necrotic cell death. The penumbra is the tissue surrounding the core where the blood flow is partly preserved due to collateral circulation and diffusion [55]. The decline in blood flow and the accompanying loss of oxygen supply result in a reduction of high-energy metabolites such as adenosine triphosphate (ATP) and phosphocreatine [39]. The combination of ATP breakdown and compensatory activation of anaerobic glycolysis during ischemia leads to an increase in the levels of inorganic phosphate, lactate, and Hþ formation causing cellular acidification. The decrease of cellular ATP levels impairs the ability of T.V. Arumugam (*) School of Biomedical Sciences, The University of Queensland, Brisbane, QLD 4072, Australia e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_1, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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membrane ion-motive ATPases to remove Naþ and Ca2þ from the cell [2]. This results in membrane depolarization, which promotes activation of synaptic glutamate receptors (Fig. 1). Neurons are normally exposed to brief pulses of glutamate because excess extracellular glutamate is actively returned to presynaptic terminals and glial cells. During ischemia, however, the energydependent mechanisms responsible for glutamate re-uptake are impaired, hence extracellular glutamate levels can reach 100 mM. Excessive accumulation of extracellular glutamate further activates glutamate receptors, resulting in massive Ca2þ influx through N-methyl-D-aspartate (NMDA) receptors, a-amino-3-hydroxy-5-methyl-4-isoxazolepropionate (AMPA) receptors, and voltage-dependent Ca2þ channels [43, 24]. This elevation of extracellular glutamate causes a prolonged and excessive activation of membrane glutamate receptors, further stimulating Ca2þ influx. Mitochondrial dysfunction occurs as the result of energy failure and disruption of cellular Ca2þ homeostasis.

Fig. 1 Ionic dysregulation during cerebral ischemia. The decrease of cellular ATP levels impairs the ability of membrane ion-motive ATPases to remove Naþ and Ca2þ from the cell. The increase in cytosolic Naþ causes cell swelling. Increased levels of intracellular Naþ can directly cause oxidative stress and cell death. When intracellular Naþ, Cl–, and H2O increase occur at the same time, cytotoxic edema occurs. Excessive Kþ efflux and intracellular Kþ depletion play a role in the apoptotic cascade. Kþ release from mitochondria contributes to the increased ROS production and ATP depletion. Ca2þ influx induces activation of two distinct cysteine proteases: caspases and calpains. These in turn cause degradation of cytoskeletal proteins, membrane receptors, and metabolic enzymes. Ca2þ also induces oxidative stress, which along with caspases induces mitochondrial dysfunction that results in cell death by apoptosis

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Increased production of free radicals results from mitochondrial dysfunction, Ca2þ overload, and activation of enzymes such as cyclooxygenase and nitric oxide synthase [39]. Free radicals damage cellular proteins, DNA, and membrane lipids. A particularly important aspect of oxidative stress in neurons is membrane lipid peroxidation, which results in the generation of toxic aldehydes such as 4-hydroxynonenal that impair the function of membrane ion-motive ATPases and glucose and glutamate transporters, thereby amplifying disruption of cellular calcium homeostasis [36]. The traditional view that glutamate-mediated disruption of cellular Ca2þ homeostasis in neurons has been expanded by a number of observations that implicate Cl channels and several types of non-channel transporter proteins, such as the Naþ/Ca2þ exchangers (NCXs), Kþ-dependent Naþ/Ca2þ exchangers (NCKXs), Naþ/Kþ/Ca2þ/Cl cotransporters (NKCCs), and Naþ/Hþ exchangers (NHEs) in the development of stroke lesion [2, 21, 11, 14]. Some of these ion transporters increase tissue damage by promoting pathological cell swelling and necrotic cell death, while others contribute to a long-term accumulation of cytoplasmic Ca2þ. Apart from the above-mentioned ion transporters, recently identified channels such as acid-sensing ionic channels (ASICs) [58], the sulfonylurea receptors (SUR1s) [52], TWIK-related potassium channels (TREKs) [8], and the transient receptor potential channels (TRPCs) [30] are also implicated in stroke pathology. This chapter summarizes the mechanisms of disruption of Ca2þ, Naþ, Kþ, and Cl homeostasis in cerebral ischemia.

2 Disruption of Ca2þ Homeostasis in Cerebral Ischemia Studies of stroke patients and animal and cell culture models have provided a wealth of data supporting the involvement of alterations in Ca2þ regulation in the pathogenesis of stroke. Activation of glutamate receptors and voltagedependent Ca2þ channels (VDCCs) and impairment of NCXs and ion-motive ATPases contribute to neuronal Ca2þ overload and cell death after a stroke [22]. Therefore, stabilization of intracellular Ca2þ has been a major goal in the search for a therapeutic method of minimizing the brain injury that follows stroke. Neurons are excitable cells that rapidly transfer electrochemical signals in a highly controlled spatio-temporal manner. A major intracellular messenger that mediates many physiological responses of neurons is Ca2þ. When properly controlled, Ca2þ fluxes across the plasma membrane and between intracellular compartments play critical roles in fundamental functions of neurons, including the regulation of neurite outgrowth and synaptogensis, synaptic transmission and plasticity, and cell survival [34]. The influx of Ca2þ through voltagedependent and ligand-gated channels in the plasma membrane is a critical signal for the release of neurotransmitters from presynaptic terminals and for responses of the postsynaptic neuron [37]. Glutamate, the major excitatory neurotransmitter in the central nervous system (CNS), induces an increase in

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the concentration of cytoplasmic Ca2þ by directly activating AMPA and NMDA receptor channels and by indirectly activating VDCC [34, 48]. In addition, activation of metabotropic glutamate receptors coupled to the GTPbinding protein Gq11 stimulates the release of inositol triphosphate (IP3), which activates Ca2þ channels in the endoplasmic reticulum (ER) [17]. Ca2þ is removed from the cytoplasm by the activities of the plasma membrane NCXs, plasma membrane and ER Ca2þ-ATPases, and Ca2þ-binding proteins such as calbindin and parvalbumin [38, 40]. By buffering intracellular Ca2þ loads, Ca2þ-binding proteins such as calbindin may serve as endogenous antiexcitotoxic proteins [40]. Ca2þ can also be transported into and released from mitochondria [56]. During normal physiological activity, the intracellular Ca2þ concentration increases only transiently (seconds to a few minutes) and has no adverse effects on the neurons [53]. However, in cerebral ischemia the ability of neurons to control Ca2þ fluxes and recover from a Ca2þ load is compromised. Excessive Ca2þ levels induce neuronal cell death in several different crossamplifying cascades: (1) Ca2þ activates either directly or indirectly cysteine proteases called calpains and caspases that degrade a variety of substrates, including cytoskeletal proteins, membrane receptors, and metabolic enzymes [10]. Calpains may also play an important role in the triggering of apoptotic cascades by virtue of their ability to activate caspases [28]. (2) Ca2þ induces oxidative stress [35] through several different mechanisms. This includes activation of oxygenases, perturbation of mitochondrial Ca2þ and energy metabolism, and induction of membrane-associated oxidative stress (MAOS). The reactive oxygen species (ROS) generated in response to glutamate-induced Ca2þ influx includes superoxide anion radicals, hydrogen peroxide, hydroxyl radicals, nitric oxide, and peroxynitrite [32]. (3) Ca2þ triggers apoptosis, a form of programmed cell death [1]. This might occur by Ca2þ -mediated induction/activation of pro-apoptotic proteins such as Bax, Par-4, and p53 leading to mitochondrial membrane permeability changes, release of cytochrome c, and caspase activation [13, 12].

3 Disruption of Naþ Homeostasis in Cerebral Ischemia Maintenance of a low intracellular Naþ is critical for normal cell function. Under steady-state conditions, intracellular Naþ is generally maintained below 20 mM whereas the extracellular Naþ concentration is 140 mM. This sevenfold concentration difference, together with the negative membrane potential (Vm), constitutes a substantial inward driving force for Naþ, which is used ubiquitously to drive a wide variety of transport processes. Cell volume and net intracellular osmolyte concentration play a pivotal role in a wide range of cellular processes and consequently must be tightly regulated to maintain normal cell function. In healthy cells, intracellular Naþ is maintained relatively

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constant due to Naþ-Kþ-ATPase activity [4, 42]. Although excitotoxicity is frequently considered synonymous with Ca2þ-dependent cell death, early studies clearly showed neurotoxic effects of glutamate such as rapid toxicity that are determined mainly by the influx of Naþ, Cl–, as well as osmotic swelling, and a more delayed Ca2þ-dependent cell death [57, 23]. One form of excitotoxicity relies on the presence of extracellular Naþ and Cl but not Ca2þ, and manifests as rapidly forming dendritic varicosities followed by generalized somatic swelling and necrotic cell death. Replacement of extracellular Naþ or Cl with impermeant ion species prevents glutamate-induced cell swelling and strongly reduces cell death. Pharmacological evidence suggests that Naþ enters the cell mainly via the glutamate-gated AMPA and NMDA channels [42, 20]. Increases in intracellular Naþ during cerebral ischemia favor NCX-mediated Ca2þ influx and consequent Ca2þ-induced cell damage [29]. The rapid increases in intracellular Naþ and Ca2þ together with swelling of neurons, astrocytes, and endothelial cells are major causes of brain damage associated with limited perfusion. Uncontrolled cell swelling is often associated with necrosis and is harmful to neuronal and glial cells [44]. The extent of intracellular Naþ elevation and cell swelling varies depending on the location in the ischemic region, with the most pronounced effects occurring in the core of the ischemic zone and more moderate slower onset effects in the penumbral regions. During cerebral ischemia, brain edema forms by a process involving increased secretion of Naþ and water across an intact brain blood barrier (BBB) [49]. At the same time, ischemia stimulates NHE1and NKCC1-dependent uptake of Naþ Cl, and water, causing cytotoxic edema [29, 47, 33], a process likely facilitated by the increased BBB secretion of Naþ, Cl, and water into the brain interstitium. Apart from Naþ influx and accumulation in the intracellular space-inducing cell swelling, increased levels of intracellular Naþ can directly cause damage, oxidative stress, and cell death. When oxidative stress occurs together with a Naþ influx, the damaging effect of oxidative stress is greatly exacerbated. The basis for this is an increased utilization of ATP by the Naþ pump activated by a Naþ entry coupled with the inability of mitochondria to respond adequately with increasing ATP production. An increase in intracellular Naþ through voltage-sensitive sodium channels (VSSCs) during hypoxia contributes to apoptosis [7]. Hypoxia increases intracellular Naþ and induces neuronal apoptosis. Reducing Naþ influx with the VSSC blocker, tetrodotoxin (TTX), attenuates apoptotic neuronal death via a reduction in caspase-3 activation [7]. Activation of VSSCs during hypoxia causes altered expression of genes that are critical regulators of apoptotic cell death [7].

4 Disruption of Kþ Homeostasis in Cerebral Ischemia Extracellular Kþ can range within 2.5–3.5 mM under normal conditions to 50–80 mM under ischemic and spreading depression events. Sustained exposure to elevated Kþ has been shown to cause significant neuronal death even under

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conditions of abundant glucose supply [54]. Physiological concentrations of potassium prevent death receptor activation, cytochrome c release, apoptosome formation, caspase activation, and apoptotic nuclease activity [59]. Kþ channels are a major contributor to a cell’s resting potential and their activation helps to maintain a hyperpolarized resting membrane potential. Kþ efflux occurs prior to Naþ or Ca2þ influx [27], which does not occur until ATP levels have fallen by more than 50%. There are a number of different types of Kþ channels. The metabolic nature of an ischemic insult suggests that ATP-sensitive Kþ channels, which are activated by a decrease in ATP, would be one of the first channels to respond during ischemia [60]. Astrocytes attempt to buffer this increase in Kþ efflux by switching to anaerobic glycolysis and swell 5–10 times their normal size. Eventually, astrocytes are no longer able to cope with the increase in Kþ efflux and they lyse. It was shown that cortical neurons exposed to glutamate with reduced Naþ and Ca2þ substantially lose their intracellular Kþ and undergo apoptosis. Both Kþ loss and apoptosis can be attenuated by increasing extracellular Kþ, indicating that glutamate receptor-mediated Kþ efflux contributes to neuronal apoptosis after brain ischemia [60]. Excessive Kþ efflux and intracellular Kþ depletion have been hypothesized to be key steps in the apoptotic cascade of many cells, including central neurons. One possible factor limiting the benefit of enhancing Kþ channel activity in the ischemic brain might be enhancement of an apoptotic component of focal ischemic neuronal death. Potassium channel openers are neuroprotective against ischemic cell death in rodents, presumably as a result of membrane hyperpolarization and reduced membrane excitability [59]. Kþ channel openers may interfere with Ca2þ mobilization from intracellular stores; in addition, these openers may mimic preconditioning or alter KATP channels on mitochondrial membranes [59]. Kþ release from mitochondria, and subsequent efflux from the cell, occurs in neurons subjected to ischemic insults that induce apoptosis. The ability of 5-hydroxydecanoate (5HD), an inhibitor of mitochondrial Kþ channels, to reduce ischemic brain injury in vivo and apoptosis of cultured neurons suggests a role for modulation of mitochondrial KATP (Mito-KATP) in neuronal death cascades. The protective effect of 5HD against focal ischemic brain injury was only observed when 5HD was administered prior to the onset of ischemia, suggesting that opening of Mito-KATP soon after the onset of ischemia plays an important role in the cell death cascade. During ischemia, Mito-KATP channels may be open and contribute to the increased ROS production and ATP depletion [31].

5 Disruption of Cl Homeostasis in Cerebral Ischemia Anion channels play an electrogenic role in excitable cells, shifting the membrane potential. Recent studies have demonstrated more general roles for anion channels. These include roles in cell volume regulation, cell proliferation, and cell death

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[26, 25]. In adult mammalian neurons, Clconcentration difference between the extracellular milieu (110–150 mM) and the intracellular space (7–30 mM) generates a gradient that is maintained by several Cl extrusion mechanisms, including Kþ–Cl cotransporters, Naþ-dependent Cl/HCO3– exchangers, ClATPase, and VDCCs [51]. Following cerebral ischemia, intracellular Cl increases by several mechanisms, including passive influx to accompany the influx of Naþ, influx through GABA-gated Cl– channels, inhibition of the voltage-gated Cl channel, hypofunction or reversal of outward Clcotransporters, and activation of inward Clcotransporters [45, 51, 11]. In in vitro models of ischemia, an increase in intracellular Cl occurs in area CA1 cell as well as CA1 pyramidal neurons and interneurons early following the onset of re-oxygenation [50]. Similar increases in intracellular Cl have been observed in hypoglossal neurons subjected to anoxia [19]. GABA, an inhibitory neurotransmitter, is usually hyperpolarizing under cerebral ischemia, because the equilibrium reversal potential for Cl (ECl) is negative relative to the resting membrane potential. In some instances, intracellular Cl– levels are higher than normal and ECl shifts to more positive values. If ECl is more positive than the resting membrane potential, GABAA receptor activation causes Cl efflux and membrane depolarization [51]. The increase in intracellular Cl following energy deprivation is detrimental to the neurons. For example, reduced GABAA responses in area CA1 pyramidal neurons are observed following the oxygen–glucose deprivation–induced rise in intracellular Cl [50, 18]. These in vitro findings are supported by experiments with animal models of cerebral ischemia. In rats subjected to focal cerebral ischemia, there is a depolarizing shift in the reversal potential for GABAA-mediated inhibitory postsynaptic potentials [41]. When ATP levels are low during cerebral ischemia, Cl–ATPase fails to transport Cl into the extracellular space. This may result in ATPdependent rundown of GABAA currents in neurons [16]. Increase in extracellular Kþ levels during ischemia may drive both Kþ and Cl back into the neuron [46]. Elevations in extracellular Kþ have been shown to reduce the GABA-mediated inhibitory postsynaptic potential due to a positive shift in the ECl [9] and to stimulate the Naþ/Kþ/Cl cotransporter to increase intracellular Cl [51].

6 Conclusion Cell culture and animal models of ischemic brain injury have proven invaluable in elucidating the cellular and molecular mechanisms of neuronal cell death in stroke. The failure of ionic homeostasis is a hallmark of ischemic brain damage. Cell surface signaling events include the activities of voltage-gated Kþ, Naþ, and Ca2þ channels and ligand-gated glutamate and GABA receptors and channels. Ca2þ channels have received a lot of attention in studies of cerebral ischemia because Ca2þ influx and the disruption of Ca2þ homeostasis play an important role in ischemic cell death. Intracellular signaling events include

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alterations in cytosolic and subcellular Ca2þ dynamics, Ca2þ-dependent kinases, and immediate early genes whereas intercellular mechanisms include mitochondrial dysfunction and free radical formation. Kþ channels are major contributors to a cell’s resting potential and their activation helps to maintain a hyperpolarized resting membrane potential. Naþ channels also play an important role in neuronal excitability and they are as widely expressed as Kþ channels. Failure of both Kþ and Naþ channels in cerebral ischemia causes significant neuronal death. In addition intracellular Cl increases by number of different mechanisms. Pathological accumulation of Naþ and Cl plays a very important role in the development of ischemic cell and tissue injury. The damaging effects of high cytosolic Naþ and Cl are realized via pathological cell swelling, the impact of Naþ overload on Ca2þ homeostasis, and an increased metabolic burden that is placed on the cell due to enhanced work of the Naþ, Kþ, and Ca2þ pumps. Effective stroke therapies may indeed require targeting several points in the neurotoxic cascade in a specific temporal sequence. Such a combination therapy may begin with the immediate administration of clot-dissolving tPA, followed by a Ca2þ channel blocker, as well as inhibitors blocking Naþ, Kþ, and Cl transport pathways.

References 1. Ankarcrona M, Dypbukt JM, Bonfoco E, Zhivotovsky B, Orrenius S, Lipton SA, and Nicotera P. Glutamate-induced neuronal death: a succession of necrosis or apoptosis depending on mitochondrial function. Neuron 15: 961–973, 1995. 2. Annunziato L, Pannaccione A, Cataldi M, Secondo A, Castaldo P, Di Renzo G, and Taglialatela M. Modulation of ion channels by reactive oxygen and nitrogen species: a pathophysiological role in brain aging? Neurobiol Aging 23: 819–834, 2002. 3. Annunziato L, Pignataro G, Boscia F, Sirabella R, Formisano L, Saggese M, Cuomo O, Gala R, Secondo A, Viggiano D, Molinaro P, Valsecchi V, Tortiglione A, Adornetto A, Scorziello A, Cataldi M, and Di Renzo GF. ncx1, ncx2, and ncx3 gene product expression and function in neuronal anoxia and brain ischemia. Ann N Y Acad Sci 1099: 413–426, 2007. 4. Aperia A. New roles for an old enzyme: Na,K-ATPase emerges as an interesting drug target. J Intern Med 261: 44–52, 2007. 5. Arumugam TV, Granger DN, and Mattson MP. Stroke and T-cells. Neuromolecular Med 7: 229–242, 2005. 6. Arumugam TV, Selvaraj PK, Woodruff TM, and Mattson MP. Targeting ischemic brain injury with intravenous immunoglobulin. Expert Opin Ther Targets 12: 19–29, 2008. 7. Banasiak KJ, Burenkova O, and Haddad GG. Activation of voltage-sensitive sodium channels during oxygen deprivation leads to apoptotic neuronal death. Neuroscience 126: 31–44, 2004. 8. Buckler KJ and Honore´ E. The lipid-activated two-pore domain K+ channel TREK-1 is resistant to hypoxia: implication for ischaemic neuroprotection. J Physiol 562: 213–222, 2005. 9. Chamberlin NL and Dingledine R. GABAergic inhibition and the induction of spontaneous epileptiform activity by low chloride and high potassium in the hippocampal slice. Brain Res 445: 12–18, 1988.

Basis of Ionic Dysregulation in Cerebral Ischemia

9

10. Chan SL and Mattson MP. Caspase and calpain substrates: roles in synaptic plasticity and cell death. J Neurosci Res 58: 167–190, 1999. 11. Chen H and Sun D. The role of Na-K-Cl co-transporter in cerebral ischemia. Neurol Res 27: 280–286, 2005. 12. Culmsee C and Mattson MP. p53 in neuronal apoptosis. Biochem Biophys Res Commun 331: 761–777, 2005. 13. Culmsee C, Zhu Y, Krieglstein J, and Mattson MP. Evidence for the involvement of Par-4 in ischemic neuron cell death. J Cereb Blood Flow Metab 21: 334–343, 2001. 14. Cuomo O, Gala R, Pignataro G, Boscia F, Secondo A, Scorziello A, Pannaccione A, Viggiano D, Adornetto A, Molinaro P, Li XF, Lytton J, Di Renzo G, and Annunziato L. A critical role for the potassium-dependent sodium-calcium exchanger NCKX2 in protection against focal ischemic brain damage. J Neurosci 28: 2053–2063, 2008. 15. Donnan GA, Fisher M, Macleod M, and Davis SM. Stroke. Lancet 371: 1612–1623, 2008. 16. Harata N, Wu J, Ishibashi H, Ono K, and Akaike N. Run-down of the GABAA response under experimental ischaemia in acutely dissociated CA1 pyramidal neurones of the rat. J Physiol 500 (Pt 3): 673–688, 1997. 17. Hartmann J and Konnerth A. Determinants of postsynaptic Ca2+ signaling in Purkinje neurons. Cell Calcium 37: 459–466, 2005. 18. Inglefield JR and Schwartz-Bloom RD. Activation of excitatory amino acid receptors in the rat hippocampal slice increases intracellular Cl– and cell volume. J Neurochem 71: 1396–1404, 1998. 19. Jiang C, Agulian S, and Haddad GG. Cl– and Na+ homeostasis during anoxia in rat hypoglossal neurons: intracellular and extracellular in vitro studies. J Physiol 448: 697–708, 1992. 20. Kiedrowski L. NCX and NCKX operation in ischemic neurons. Ann N Y Acad Sci 1099: 383–395, 2007. 21. Kitayama J, Kitazono T, Yao H, Ooboshi H, Takaba H, Ago T, Fujishima M, and Ibayashi S. Inhibition of Na+/H+ exchanger reduces infarct volume of focal cerebral ischemia in rats. Brain Res 922: 223–228, 2001. 22. Kristia´n T and Siesjo¨ BK. Calcium in ischemic cell death. Stroke 29: 705–718, 1998. 23. Kumar V, Naik RS, Hillert M, and Klein J. Effects of chloride flux modulators in an in vitro model of brain edema formation. Brain Res 1122: 222–229, 2006. 24. Kwak S and Weiss JH. Calcium-permeable AMPA channels in neurodegenerative disease and ischemia. Curr Opin Neurobiol 16: 281–287, 2006. 25. Lang F. Mechanisms and significance of cell volume regulation. J Am Coll Nutr 26: 613S–623S, 2007. 26. Lang F, Foller M, Lang KS, Lang PA, Ritter M, Gulbins E, Vereninov A, and Huber SM. ¨ Ion channels in cell proliferation and apoptotic cell death. J Membr Biol 205: 147–157, 2005. 27. Leis JA, Bekar LK, and Walz W. Potassium homeostasis in the ischemic brain. Glia 50: 407–416, 2005. 28. Leist M, Volbracht C, Ku¨hnle S, Fava E, Ferrando-May E, and Nicotera P. Caspasemediated apoptosis in neuronal excitotoxicity triggered by nitric oxide. Mol Med 3: 750–764, 1997. 29. Lenart B, Kintner DB, Shull GE, and Sun D. Na-K-Cl cotransporter-mediated intracellular Na+ accumulation affects Ca2+ signaling in astrocytes in an in vitro ischemic model. J Neurosci 24: 9585–9597, 2004. 30. Lipski J, Park TI, Li D, Lee SC, Trevarton AJ, Chung KK, Freestone PS, and Bai JZ. Involvement of TRP-like channels in the acute ischemic response of hippocampal CA1 neurons in brain slices. Brain Res 1077: 187–199, 2006. 31. Liu D, Slevin JR, Lu C, Chan SL, Hansson M, Elmer E, and Mattson MP. Involvement of mitochondrial K+ release and cellular efflux in ischemic and apoptotic neuronal death. J Neurochem 86: 966–979, 2003.

10

T.V. Arumugam et al.

32. Love S. Oxidative stress in brain ischemia. Brain Pathol 9: 119–131, 1999. 33. Luo J, Kintner DB, Shull GE, and Sun D. ERK1/2-p90RSK-mediated phosphoryl tion of Na+/H+ exchanger isoform 1. A role in ischemic neuronal death. J Biol Chem 282: 28274–28284, 2007. 34. Mattson MP. Neurotransmitters in the regulation of neuronal cytoarchitecture. Brain Res 472: 179–212, 1988. 35. Mattson MP. Modification of ion homeostasis by lipid peroxidation: roles in neuronal degeneration and adaptive plasticity. Trends Neurosci 21: 53–57, 1998. 36. Mattson MP. Excitotoxic and excitoprotective mechanisms: abundant targets for the prevention and treatment of neurodegenerative disorders. Neuromolecular Med 3: 65–94, 2003. 37. Mattson MP. Calcium and neurodegeneration. Aging Cell 6: 337–350, 2007. 38. Mattson MP, Guthrie PB, and Kater SB. A role for Na+-dependent Ca2+ extrusion in protection against neuronal excitotoxicity. FASEB J 3: 2519–2526, 1989. 39. Mattson MP, Culmsee C, and Yu ZF. Apoptotic and antiapoptotic mechanisms in stroke. Cell Tissue Res 301: 173–187, 2000. 40. Mattson MP, Rychlik B, Chu C, and Christakos S. Evidence for calcium-reducing and excito-protective roles for the calcium-binding protein calbindin-D28k in cultured hippocampal neurons. Neuron 6: 41–51, 1991. 41. Mittmann T, Qu¨ M, Zilles K, and Luhmann HJ. Long-term cellular dysfunction after focal cerebral ischemia: in vitro analyses. Neuroscience 85: 15–27, 1998. 42. Mongin AA. Disruption of ionic and cell volume homeostasis in cerebral ischemia: The perfect storm. Pathophysiology 14: 183–193, 2007. 43. Nakamichi N, Oikawa H, Kambe Y, and Yoneda Y. Relevant modulation by ferrous ions of N-methyl-D-aspartate receptors in ischemic brain injuries. Curr Neurovasc Res 1: 429–440, 2004. 44. Okada Y, Maeno E, Shimizu T, Dezaki K, Wang J, and Morishima S. Receptor-mediated control of regulatory volume decrease (RVD) and apoptotic volume decrease (AVD). J Physiol 532: 3–16, 2001. 45. Okada Y, Shimizu T, Maeno E, Tanabe S, Wang X, and Takahashi N. Volume-sensitive chloride channels involved in apoptotic volume decrease and cell death. J Membr Biol 209: 21–29, 2006. 46. Payne JA. Functional characterization of the neuronal-specific K-Cl cotransporter: implications for [K+]o regulation. Am J Physiol 273: C1516–C1525, 1997. 47. Pedersen SF, O’Donnell ME, Anderson SE, and Cala PM. Physiology and pathophysiology of Na+/H+ exchange and Na+-K+-2Cl– cotransport in the heart, brain, and blood. Am J Physiol Regul Integr Comp Physiol 291: R1–R25, 2006. 48. Pinheiro PS and Mulle C. Presynaptic glutamate receptors: physiological functions and mechanisms of action. Nat Rev Neurosci 9: 423–436, 2008. 49. Rosenberg GA. Ischemic brain edema. Prog Cardiovasc Dis 42: 209–216, 1999. 50. Sah R and Schwartz-Bloom RD. Optical imaging reveals elevated intracellular chloride in hippocampal pyramidal neurons after oxidative stress. J Neurosci 19: 9209–9217, 1999. 51. Schwartz-Bloom RD and Sah R. Gamma-aminobutyric acid(A) neurotransmission and cerebral ischemia. J Neurochem 77: 353–371, 2001. 52. Simard JM, Tarasov KV, and Gerzanich V. Non-selective cation channels, transient receptor potential channels and ischemic stroke. Biochim Biophys Acta 1772: 947–957, 2007. 53. Takahashi A, Camacho P, Lechleiter JD, and Herman B. Measurement of intracellular calcium. Physiol Rev 79: 1089–1125, 1999. 54. Wei L, Yu SP, Gottron F, Snider BJ, Zipfel GJ, and Choi DW. Potassium channel blockers attenuate hypoxia- and ischemia-induced neuronal death in vitro and in vivo. Stroke 34: 1281–1286, 2003. 55. Weinstein PR, Hong S, and Sharp FR. Molecular identification of the ischemic penumbra. Stroke 35: 2666–2670, 2004.

Basis of Ionic Dysregulation in Cerebral Ischemia

11

56. Werth JL and Thayer SA. Mitochondria buffer physiological calcium loads in cultured rat dorsal root ganglion neurons. J Neurosci 14: 348–356, 1994. 57. Won SJ, Kim DY, and Gwag BJ. Cellular and molecular pathways of ischemic neuronal death. J Biochem Mol Biol 35: 67–86, 2002. 58. Xiong ZG, Chu XP, and Simon RP. Ca2+-permeable acid-sensing ion channels and ischemic brain injury. J Membr Biol 209: 59–68, 2006. 59. Yu SP. Regulation and critical role of potassium homeostasis in apoptosis. Prog Neurobiol 70: 363–386, 2003. 60. Yu SP, Yeh C, Strasser U, Tian M, and Choi DW. NMDA receptor-mediated K+ efflux and neuronal apoptosis. Science 284: 336–339, 1999.

Why have Ionotropic and Metabotropic Glutamate Antagonists Failed in Stroke Therapy? Gianfranco Di Renzo, Giuseppe Pignataro, and Lucio Annunziato

1 Introduction The concept of ‘‘excitotoxicity’’ was introduced in 1969 when Olney and Sharpe first demonstrated that neurons exposed to their own neurotransmitter glutamate were destined to die [49]. Later on, in 1985, glutamate toxicity was associated with anoxic cell death, since anoxic depolarization resulted in the release of glutamate into the extracellular compartments [42, 54]. Similarly, in 1987, Choi indicated that glutamate was a remarkably potent and rapidly acting neurotoxin able to mediate neurotoxic effects by inducing Ca2þ influx through glutamate receptor activation, and he supported the theory that glutamate can be considered a key neurotransmitter in developing many neurological diseases [13, 14, 15]. Since then, glutamate receptors have been the most studied channels involved in ischemic stroke pathophysiology. Glutamate can exert its effects by interacting with both ionotropic glutamate receptors (iGluRs), also referred to as ligand-gated ion channels, and metabotropic glutamate G-protein-coupled receptors. The group of iGluRs comprises three major classes, the a-amino-3hydroxy-5-methyl-4-isoxazolepropionate (AMPA) receptors, kainate (KA) receptors, and N-methyl-D-aspartate (NMDA) receptors, named according to their selective agonists [62]. Glutamatergic synapses frequently harbor both AMPA and NMDA receptors. Characteristically, both classes of receptors differ in their response kinetics upon presynaptic glutamate release. Indeed, AMPA receptors mediate fast glutamate-gated postsynaptic responses, even at very negative potentials or in the absence of action potentials. The fast desensitization of AMPA receptors leads to short excitatory postsynaptic currents (EPSCs). In contrast, NMDA receptors contain an agonist-binding site, i.e., a glycine modulatory site, and other binding sites within the ion channel, where magnesium exerts a voltage-dependent block [21]. Acting as detectors of G. Di Renzo (*) Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, Via Pansini 5, 80131, Naples, Italy e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_2, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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membrane depolarization and ligand-gated channel activation, NMDA receptors require the removal of the Mg2þ block by an increase in membrane potential to allow cation permeation through the receptor pore [62]. The excitotoxicity theory encountered a great favor for almost 40 years, because of a discrete success of glutamate receptors antagonists in treating experimentally induced stroke. Unfortunately, however, over the last two decades, all stroke clinical trials with agents acting on these receptors have been disappointing [16, 17, 45, 46, 48]. The disappointing experience derived from the use of pharmacological agents interfering with glutamate-mediated mechanisms in stroke pathophysiology has been paid by many scientists working in the field with the reluctance of government and private associations to grant projects on the development of new drugs to treat stroke. Thus, the discouraging results have redirected scientists’ attention to other channels and membrane transporters able to control neuronal ionic homeostasis. The purpose of this chapter is to review some of the evidence in the field in an attempt to explain why drugs acting on glutamate receptors have resulted ineffective in treating stroke.

2 Glutamate and Stroke It is widely accepted that a critical factor in determining neuronal and glial death during cerebral ischemia is the progressive accumulation of intracellular Naþ([Naþ]i) and Ca2þ([Ca2þ]i) ions, which can precipitate necrosis and apoptosis of vulnerable neurons. Whereas the detrimental action of [Naþ]i increase is attributable to both cell swelling and microtubular disorganization – two phenomena that lead to cell necrosis [59] – a rise in [Ca2þ]i has been shown to be a key factor in ischemic brain damage, for it modulates several death pathways, including oxidative and nitrosative stress, mitochondrial dysfunction, and protease activation. Since Olney’s seminal work first suggested that excitatory amino acids could elicit neurotoxicity [49], a vast amount of data have demonstrated that glutamate extracellular concentrations briskly rise during acute brain injury, thus triggering an influx of Ca2þ and Naþ ions into neurons through glutamate receptors [13]. This evidence has led to the elaboration of the paradigm of glutamate excitotoxicity, a theory that explained ischemic neuronal cell death as a mere consequence of Naþ and Ca2þ influx through glutamate receptors [13]. Although this paradigm has been guiding basic research in the field of neurodegeneration for almost three decades, more recently it has become the object of serious criticism and re-assessment. What has aroused such skepticism among researchers has been the fact that although first-, second-, and third-generation glutamate receptor antagonists have long yielded promising results in animal models of brain ischemia, they have failed to elicit a neuroprotective action in stroke and traumatic brain injury in humans. Therefore, the theory of excitotoxicity, though a fascinating paradigm, can only

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explain some of the events occurring in the acute phase of anoxic insult but cannot be seen as a major target for developing new therapeutic avenues for brain ischemia. In fact, the energy failure occurring during the course of a stroke episode triggers many events that are strictly dependent on several factors: 1. the type of ischemic event, i.e., focal vs global ischemia 2. the region of the central nervous system (CNS) involved in the mechanism of ischemic damage 3. the distance from the area primarily affected by the ischemic event, i.e., ischemic core vs ischemic penumbra 4. the duration of the ischemic event, and 5. the time elapsing between the onset of the ischemic event and the evaluation of the severity of the insult. Furthermore, the fact that after stroke onset Ca2þ concentrations follow a triphasic response that is not equally sensitive to NMDA receptor antagonists [52, 66] should be taken into account. In fact, during exposure to excitatory neurotransmitters, intracellular calcium increase is followed by a transient return to basal levels. After a free interval of a few hours, a gradually progressing secondary increase occurs. The initial Ca2þ influx can be blocked by NMDA receptor antagonists or by the removal of Ca2þ from the extracellular medium but not by antagonists of non-NMDA receptors [52, 66]. The second delayed Ca2þ increase can be counteracted by extracellular Ca2þ removal but not by the application of an NMDA or a non-NMDA receptor antagonist. Interestingly, this second Ca2þ increase can be accelerated by increasing extracellular Ca2þ or by blocking the plasma membrane of Naþ/Ca2þ exchanger [2, 66]. The changes in [Ca2þ]i probably produce other leak currents that result in irreversible disturbances in ion homeostasis and, eventually, in cell death. Although the dominant mediator of the delayed injury has not yet been established, it is likely that a complex chain of events, including the activation of the Ca2þ-dependent catabolic process, the early damage by Naþ overload that induces oncotic cell swelling and ionic edema, and the dysregulation of the ionic homeostasis of other ions, such as zinc and magnesium, may act to destroy cellular integrity. This indicates that not only glutamate antagonists but also a great variety of other drugs that interfere with the other processes might improve neuronal survival after ischemic damage. In addition, when the features of global and focal ischemia are analyzed in association with glutamate levels, several substantial findings challenge the excitotoxic hypothesis of ischemic injury. First, global ischemia induces a damage that is confined to well-defined vulnerable areas although glutamate is released in the same amount throughout the brain [6, 57]. Second, ischemic damage after global ischemia is delayed for days whereas glutamate levels increase immediately after the onset of the ischemic event. Third, since neuronal death in the core region is mainly due to energy depletion, glutamate toxicity should be considered important only for

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the damage occurring in penumbra, the peripheral region of the ischemic lesion. However, in focal ischemia extracellular glutamate increases up to 100 times above the basal level in the ischemic core, reaches peak values 1–2 hours later, and declines slowly thereafter. By contrast, in the ischemic penumbra, it increases only up to 25 times above the basal level and may return to normal within 30 min even if blood flow does not improve [20]. Fourth, the hypothesis that the possible excitotoxicity is induced by glutamate release is also challenged by the fact that the release of excitatory amino acids can be counteracted by other ischemia-related factors such as acidosis and by the release of hyperpolarizing inhibitory neurotransmitters, which during stroke are released at concentrations higher than those of glutamate [40]. Finally, microdialysis studies of glutamate levels after stroke have clearly demonstrated that although the extracellular glutamate concentration increases sharply and rapidly, reaching concentrations 10–100 times higher than preinsult values [7], this increase lasts only for 10–30 min [7]. Therefore, although glutamate may be involved in the acute neurodestructive phase that occurs immediately after ischemic injury, its normal physiological functions, including the promotion of neuronal survival, are resumed directly after this phase [37]. In addition, delayed treatment with NMDA antagonists suppresses neurogenesis, triggered by focal cerebral ischemia, in the hippocampus [4]. These findings suggest that in addition to damaging neurons immediately after the injury, glutamate may also facilitate repair shortly thereafter. Interestingly, whereas excitotoxic effects of glutamate are short lasting, repair mechanisms appear to be long lasting. The interference with neuronal survival means that NMDA antagonists are unsuitable neuroprotective drugs for stroke therapy. The only way to provide pharmacological neuroprotection with NMDA antagonists would be to administer them before the insult and for a very short period (even minutes) after the injury, circumstances that would be virtually impossible in a clinical emergency setting. Thus, when designing novel therapies, researchers need to consider and respect the physiological role of glutamate in the brain. By focusing on the destructive effects of glutamate after injury and by ignoring its physiological functions, many patients were unnecessarily exposed to glutamate NMDA antagonists. In addition, when we consider that overlooking the potential detrimental role of glutamate has resulted in years of long painstaking but unpromising research, as well as in unprofitable investments from pharmaceutical companies, the need to re-evaluate the physiological role of glutamate and thus to invest human and financial resources in this or other related lines of research becomes apparent. In the last few years, several seminal experimental works have markedly changed the scenario in this field. In fact, it has been shown that some integral plasma membrane proteins, involved in the control of Ca2þ, Naþ, Kþ, and Cl– ion influx or efflux and, therefore, responsible for maintaining the homeostasis of these cations and anions, might function as crucial players in the brain

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ischemic process [1, 43, 50, 64]. Indeed, these proteins may provide the molecular basis underlying glutamate-independent ionic homeostasis dysregulation in neuronal ischemic cell death and, most important, may represent more suitable molecular targets for therapeutic intervention.

3 The Reasons of the Failure of Excitotoxicity Theory In the last decade, several lines of evidence have accumulated against the concept that high levels of extracellular glutamate are associated with neurological disorders and thus may contribute to neuronal death. The reasons for that can be summarized in the following six ‘‘Hossmann postulates’’ [32]: 1. The type of metabolic and biochemical response induced by glutamate exposure is considerably different from that evoked by ischemic injury. In fact, in vitro and in vivo studies clearly demonstrated that a transient suppression of energy metabolism, as well as changes in protein synthesis, occurs after experimental ischemia [19]. In contrast, exposure to a high dose of glutamate changes neither energy state nor protein synthesis rate. Changes in protein synthesis are a consistent and probably fatal event occurring in all kinds of ischemic cell death [31]. Preservation of protein synthesis after glutamate exposure is, consequently, a strong indication that glutamate-induced damage is different from ischemic-induced damage. 2. Microdialysis and other techniques, which allow to measure glutamate levels in the brain after experimentally induced cerebral ischemia, clearly indicate that increases in glutamate after stroke are not necessarily required for induction of the pathological process. In fact, in models mimicking focal cerebral ischemia, threshold determinations of glutamate release clearly show that glutamate rises to high levels in the ischemic core but not in the penumbra [47, 56]. However, the only reason for cell necrosis in the core is energy depletion; therefore, although glutamate can contribute to cell death, it cannot be considered the main player in this mechanism. Furthermore, although the increase in extracellular glutamate is much lower in the penumbra than in the infarct core, a neurotoxic effect cannot be excluded. However, the probability of such an effect is rather small. On the other hand, in global ischemia, the measurement of glutamate in the different brain regions of rats subjected to 4-Vessel Occlusion (4-VO) have revealed that during ischemia the increases in glutamate levels are almost identical to those found in the hippocampus, striatum, cortex, and thalamus [23]. In addition, even within the hippocampus, glutamate release is the same in all affected regions. By contrast, global ischemia induces a region-specific damage and the different hippocampal regions show a selective vulnerability to the global ischemic insult. The difference is even more evident when different durations of ischemia are compared. In fact, despite the varying concentrations of glutamate in the different regions, the pattern of vulnerability does not

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change [24]. This hypothesis is further supported by the fact that ischemia induces glutamate release in regions spared from histopathological damage in the brain [24]. Since neuronal death occurring in the core region is mainly due to energy depletion, glutamate toxicity should be important only for the damage occurring in penumbra. If so, it is possible to hypothesize that glutamate neurotoxicity in vivo is less toxic than in vitro. However, owing to the activation of anaerobic glycolysis, the penumbral region suffers from a substantial degree of tissue acidosis, a condition that has been shown to alleviate glutamate toxicity in vitro [38]. The ischemic penumbra is not affected by its trophic environment because the structural integrity of this region is fully preserved. Therefore, excitotoxicity induced by glutamate is not plausible in this ischemic area. Glutamate toxicity in vitro is a delayed phenomenon that requires 24 hours to for complete evolution [15]. By contrast, the ischemic penumbra survives no longer than 6 hours. At this time point, in fact, the ischemic penumbra becomes the ischemic core [5] Besides glutamate, other ischemia-related factors such as inhibitory neurotransmitters are released and can be expected to reduce excitotoxicity [38, 40, 66]. For instance, GABA is released at the same blood flow level as glutamate and adenosine even at higher flow thresholds. Thus, the relative increase in GABA is more pronounced than that of glutamate [40].

To these six postulates it should be added that, in the whole brain, the level of glutamate can be maintained at a low threshold by all those systems, which being localized on the membrane of neurons and glial cells are able to induce glutamate re-uptake. More important, it must not be forgotten that recent experimental evidence has suggested a protective role for glutamate in the pathophysiology of the ischemic event [37]. In fact, whereas synaptic transmission mediated by NMDA receptors is essential for neuronal survival, blockade of NMDA receptors triggers apoptosis in the developing brain [35, 51]. Environmental enrichment, which stimulates synaptic activity, inhibits spontaneous apoptosis in the hippocampus and is neuroprotective [65]. Accordingly, when NMDA receptor antagonists are administered during slowly progressing neurodegeneration, they markedly exacerbate damage in the adult brain [36]. In addition, NMDA receptor antagonists cause apoptosis in primary hippocampal cultures and can exacerbate apoptosis induced by staurosporine [29]. By contrast, NMDA-receptor-mediated synaptic activation is neuroprotective in vitro and diminishes apoptosis induced by staurosporine [29]. Activation of pro-survival transcription factors, such as cAMP response element-binding protein (CREB), accompanies NMDA-mediated neuroprotection in vitro [29]. In particular, the Ca2þ pool in the immediate vicinity of synaptic NMDA receptors is able to trigger signaling from the synapse to the nucleus via the extracellular signal-regulated kinase (ERK1/2) (Fig. 1) [27]. One important function of this Ca2þ microdomain, which is located near NMDA

Failure of Ionotropic and Metabotropic Glutamate Antagonists Fig. 1 Activation of prosurvival factors by NMDA receptor agonists

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CREB Nucleus Activation of prosurvival factors

NEUROPROTECTION

receptors, is to prolong CREB phosphorylation induced by synaptic stimulation, thereby enhancing CREB-mediated gene expression. CREB controls transcription of pro-survival genes such as the brain-derived neurotrophic factor (BDNF), the vasoactive intestinal peptide (VIP), bcl-2, and mcl-1 [53, 61]. Thus, the survival-promoting properties of NMDA receptor could derive from the transcription of such pro-survival genes. In addition, NMDA activation induces an increase of cerebral blood flow NO-dependent [22], thus ameliorating brain ischemic conditions. Indeed, neurons in the penumbra that survive ischemic insults are characterized by high concentrations of BDNF, and bcl-2, and activated CREB [60], hence suggesting a sustained induction of pro-survival signals. These findings lead to the logical conclusion that suppression of survival signals promoted by NMDA receptor activation may facilitate cell death. Taken together, there is sufficient evidence to suggest that synaptic activity mediated by NMDA receptors might promote neuronal survival. Blockade of NMDA-mediated synaptic transmission could therefore be detrimental in situations when support by endogenous measures is required, as occurs after stroke or in chronic neurodegenerative disorders. From all the above-mentioned considerations it is possible to state that it is time to conclude that NMDA antagonists

20

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have failed and the attention of researchers working in the field has to be redirected to other cation channels, pumps, and ionic transporters.

4 Conclusion: Beyond NMDA Receptors The NMDA receptor has long been viewed as a major player in inducing excitotoxic cell death. For instance, Simon et al. [58] proposed that the inhibition of the receptor would protect neurons from this death. However, further animal trials demonstrated that the neuroprotective effect of NMDA receptor blockers, such as MK-801, was in fact due to a hypothermia induced by the agent and not by a specific action of the drug [9, 10, 11]. From a number of clinical trials, the surfacing of certain adverse effects has deeply discouraged scientists from fulfilling further research goals in the field. Indeed, NMDA antagonists cause a number of physiological perturbations, including an increase in blood pressure, as well as tachycardia [12]. In particular, achieving serum concentrations high enough to equal those neuroprotective ones obtained in rodents has been challenging, for competitive NMDA receptor antagonists cause many toxic side effects when given at the recommended doses [11]. As suggested by Hoyte [34], there is actually ‘‘a loss in translation’’ when moving from animal models to clinical trials. Intriguingly, the latest literature has highlighted that the activation of NMDA receptors can result in either cell survival or cell death depending on whether the receptor is synaptic or extrasynaptic [22, 26, 28, 30]. The activation of synaptic NMDA receptors promotes cell survival by activation of the CaM kinase and Ras–ERK1/2 (extracellular signal-regulated kinase) pathways and subsequent expression of BDNF [28]. In contrast, the activation of extrasynaptic NMDA receptors inactivates the CREB pathway and downregulates BDNF [28]. Beyond the synapse, different glutamate mechanisms might also operate in other parts of the neuron. In particular, in the axon, the release of glutamate and the subsequent activation of AMPA are both initiated by a large Naþ influx and the reverse of Naþ-dependent glutamate transporters [39]. Under some experimental conditions, imbalances in sodium might even be more important than calcium in axonal compartments. In neuronal dendrites, overactivation of NMDA receptors is damaging. However, activation of kainate-type receptors, which are closely related to AMPA receptors, might actually promote growth and remodeling [44]. Ultimately, glutamergic signaling is mediated not just by NMDA and AMPA currents. Many other glutamate receptors and transporters exist in the brain. Careful targeting of other neuronal glutamate receptors and transporters, including the five metabotropic glutamate (mGlu) receptor subtypes and the excitatory amino acid transporter 2 (EAAT2), could also prove fruitful, on the basis of similar variations in their capacity to trigger death or survival [8, 55]. It should be underlined that beyond the glutamateassociated channels, many other ionic channels also carry large ionic currents in

Failure of Ionotropic and Metabotropic Glutamate Antagonists

21

damaged neurons [25]. Altogether, it is important to realize that NMDA–AMPA pathways comprise only a subset of the multiple routes of ionic imbalance that are induced in brain injury and neurodegeneration. Another potential limitation of the standard NMDA–AMPA model is the focus on neurons alone. Indeed, some types of glial cells, including astrocytes and oligodendrocytes, play crucial roles in glutamate regulation, and, thus their roles ought also to be considered alongside neurons. More specifically, glutamate uptake by astrocytes via GLAST and GLT-1, the rodent form of the EAAT2, normally keeps extracellular glutamate below toxic levels. If these mechanisms are impaired by ischemia, neuronal excitotoxicity can be amplified [41]. As both astrocytes and oligodendrocytes express NMDA and AMPA/ kainate receptors, they are also vulnerable to high levels of glutamate [18, 41]. But as already seen in neurons, the processes of oligodendrocytes and astrocytes might respond differently from the cell body. It has been proposed that NMDA receptors comprising NR2C and NR3A subunits mediate injury in the glial processes whereas damages to the glial cell bodies are mediated by AMPA/ kainate receptors [63]. Additionally, glia express many different subtypes of mGlu receptors that exhibit a variety of modulatory functions [18]. Because of all these complexities in glial glutamate regulation and signaling, it might perhaps be difficult to design a single NMDA or AMPA antagonist that would protect all neurons and glial cells after stroke. In the final analysis, the standard NMDA–AMPA model of excitotoxicity is perhaps oversimplistic and does not take into account the complex interactions with other parallel routes of ionic entry and imbalance within the injured cell. Accordingly, several other emerging mechanisms of ionic imbalance deserve further attention as we continue searching for neuroprotectants against stroke and brain injury. Some of the potentially promising ones could include sodium/ potassium ATPase, sodium–calcium exchangers (NCXs), sodium–proton exchangers (NHEs), sodium–potassium–chloride (NKCC), Kþ channels, Naþ channels, acid-sensing ion channels (ASICs), transient receptor potential channels (TRPs), and other non-selective cation channels [3]. Finally, we hope that future research projects have to prize glutamate lesson highly in order to be on the straight and narrow path for setting up new effective strategies in stroke therapy.

5 Future Perspectives Two obvious factors emerge from the discussion presented above: (a) Virtually, all preclinical studies have suggested that NMDA antagonists are effectively protective in focal ischemia only if administered immediately after the insult. This property obviously constitutes a clear drawback in using NMDA antagonists in humans, for as highlighted in clinical trials, longer periods of time necessarily elapse between the onset of the insult and the actual administration of the first treatment.

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(b) The possibility exists that NMDA antagonism, though potentially protective in focal ischemia, is also deleterious in that it adversely affects endogenous NMDA-receptor-mediated neuronal-survival mechanisms [33]. However, intriguing futuristic possibilities are receiving attention and new signs of hope appear over the horizon. In fact, although the disappointment regarding the failure of NMDA antagonists is high, the neuroprotective potential of NMDA isoform-specific antagonists acting extrasynaptically remains to be explored. Furthermore, by combining glutamate, non-glutamatergic receptors, pump, and ionic transporter modulators with antioxidants, as well as anti-apoptotic and antiinflammatory agents, it will be possible to cover the time course of stroke development and eventually develop effective treatments for stroke.

References 1. Aarts M, Iihara K, Wei WL, Xiong ZG, Arundine M, Cerwinski W, MacDonald JF, and Tymianski M. A key role for TRPM7 channels in anoxic neuronal death. Cell 115: 863–877, 2003. 2. Andreeva N, Khodorov B, Stelmashook E, Cragoe E Jr., and Victorov I. Inhibition of Na+/Ca2+ exchange enhances delayed neuronal death elicited by glutamate in cerebellar granule cell cultures. Brain Res 548: 322–325, 1991. 3. Annunziato L, Cataldi M, Pignataro G, Secondo A, and Molinaro P. Glutamate-independent calcium toxicity: introduction. Stroke 38: 661–664, 2007. 4. Arvidsson A, Kokaia Z, and Lindvall O. N-methyl-D-aspartate receptor-mediated increase of neurogenesis in adult rat dentate gyrus following stroke. Eur J Neurosci 14: 10–18, 2001. 5. Back T, Kohno K, and Hossmann KA. Cortical negative DC deflections following middle cerebral artery occlusion and KCl-induced spreading depression: effect on blood flow, tissue oxygenation, and electroencephalogram. J Cereb Blood Flow Metab 14: 12–19, 1994. 6. Benveniste H. Brain microdialysis. J Neurochem 52: 1667–1679, 1989. 7. Benveniste H, Drejer J, Schousboe A, and Diemer NH. Elevation of the extracellular concentrations of glutamate and aspartate in rat hippocampus during transient cerebral ischemia monitored by intracerebral microdialysis. J Neurochem 43: 1369–1374, 1984. 8. Besancon E, Guo S, Lok J, Tymianski M, and Lo EH. Beyond NMDA and AMPA glutamate receptors: emerging mechanisms for ionic imbalance and cell death in stroke. Trends Pharmacol Sci 29: 268–275, 2008. 9. Buchan A, Li H, and Pulsinelli WA. The N-methyl-D-aspartate antagonist, MK-801, fails to protect against neuronal damage caused by transient, severe forebrain ischemia in adult rats. J Neurosci 11: 1049–1056, 1991. 10. Buchan A and Pulsinelli WA. Hypothermia but not the N-methyl-D-aspartate antagonist, MK-801, attenuates neuronal damage in gerbils subjected to transient global ischemia. J Neurosci 10: 311–316, 1990. 11. Buchan AM. Do NMDA antagonists protect against cerebral ischemia: are clinical trials warranted? Cerebrovasc Brain Metab Rev 2: 1–26, 1990. 12. Buchan AM, Slivka A, and Xue D. The effect of the NMDA receptor antagonist MK-801 on cerebral blood flow and infarct volume in experimental focal stroke. Brain Res 574: 171–177, 1992. 13. Choi DW. Glutamate neurotoxicity and diseases of the nervous system. Neuron 1: 623–634, 1988.

Failure of Ionotropic and Metabotropic Glutamate Antagonists

23

14. Choi DW. Ionic dependence of glutamate neurotoxicity. J Neurosci 7: 369–379, 1987. 15. Choi DW, Maulucci-Gedde M, and Kriegstein AR. Glutamate neurotoxicity in cortical cell culture. J Neurosci 7: 357–368, 1987. 16. Davis SM, Albers GW, Diener HC, Lees KR, and Norris J. Termination of acute stroke studies involving selfotel treatment. ASSIST Steering Committed. Lancet 349: 32, 1997. 17. Davis SM, Lees KR, Albers GW, Diener HC, Markabi S, Karlsson G, and Norris J. Selfotel in acute ischemic stroke: possible neurotoxic effects of an NMDA antagonist. Stroke 31: 347–354, 2000. 18. Deng W, Wang H, Rosenberg PA, Volpe JJ, and Jensen FE. Role of metabotropic glutamate receptors in oligodendrocyte excitotoxicity and oxidative stress. Proc Natl Acad Sci USA 101: 7751–7756, 2004. 19. Djuricic B, Rohn G, Paschen W, and Hossmann KA. Protein synthesis in the hippocampal slice: transient inhibition by glutamate and lasting inhibition by ischemia. Metab Brain Dis 9: 235–247, 1994. 20. Fabricius M, Jensen LH, and Lauritzen M. Microdialysis of interstitial amino acids during spreading depression and anoxic depolarization in rat neocortex. Brain Res 612: 61–69, 1993. 21. Ginsberg MD. Neuroprotection for ischemic stroke: past, present and future. Neuropharmacology 55: 363–389, 2008. 22. Girouard H, Wang G, Gallo EF, Anrather J, Zhou P, Pickel VM, and Iadecola C. NMDA receptor activation increases free radical production through nitric oxide and NOX2. J Neurosci 29: 2545–2552, 2009. 23. Globus MY, Busto R, Dietrich WD, Martinez E, Valdes I, and Ginsberg MD. Effect of ischemia on the in vivo release of striatal dopamine, glutamate, and gamma-aminobutyric acid studied by intracerebral microdialysis. J Neurochem 51: 1455–1464, 1988. 24. Globus MY, Busto R, Martinez E, Valdes I, and Dietrich WD. Ischemia induces release of glutamate in regions spared from histopathologic damage in the rat. Stroke 21: III43–46, 1990. 25. Gribkoff VK and Winquist RJ. Voltage-gated cation channel modulators for the treatment of stroke. Expert Opin Investig Drugs 14: 579–592, 2005. 26. Habas A, Kharebava G, Szatmari E, and Hetman M. NMDA neuroprotection against a phosphatidylinositol-3 kinase inhibitor, LY294002 by NR2B-mediated suppression of glycogen synthase kinase-3beta-induced apoptosis. J Neurochem 96: 335–348, 2006. 27. Hardingham GE, Arnold FJ, and Bading H. A calcium microdomain near NMDA receptors: on switch for ERK-dependent synapse-to-nucleus communication. Nat Neurosci 4: 565–566, 2001. 28. Hardingham GE and Bading H. The Yin and Yang of NMDA receptor signalling. Trends Neurosci 26: 81–89, 2003. 29. Hardingham GE, Fukunaga Y, and Bading H. Extrasynaptic NMDARs oppose synaptic NMDARs by triggering CREB shut-off and cell death pathways. Nat Neurosci 5: 405–414, 2002. 30. Hetman M and Kharebava G. Survival signaling pathways activated by NMDA receptors. Curr Top Med Chem 6: 787–799, 2006. 31. Hossmann KA. Disturbances of cerebral protein synthesis and ischemic cell death. Prog Brain Res 96: 161–177, 1993. 32. Hossmann KA. Excitotoxic mechanisms in focal ischemia. Adv Neurol 71: 69–74, 1996. 33. Hoyte L, Barber PA, Buchan AM, and Hill MD. The rise and fall of NMDA antagonists for ischemic stroke. Curr Mol Med 4: 131–136, 2004. 34. Hoyte L, Kaur J, and Buchan AM. Lost in translation: taking neuroprotection from animal models to clinical trials. Exp Neurol 188: 200–204, 2004. 35. Ikonomidou C, Bosch F, Miksa M, Bittigau P, Vockler J, Dikranian K, Tenkova TI, Stefovska V, Turski L, and Olney JW. Blockade of NMDA receptors and apoptotic neurodegeneration in the developing brain. Science 283: 70–74, 1999.

24

G. Di Renzo et al.

36. Ikonomidou C, Stefovska V, and Turski L. Neuronal death enhanced by N-methyl-Daspartate antagonists. Proc Natl Acad Sci USA 97: 12885–12890, 2000. 37. Ikonomidou C and Turski L. Why did NMDA receptor antagonists fail clinical trials for stroke and traumatic brain injury? Lancet Neurol 1: 383–386, 2002. 38. Kaku DA, Giffard RG, and Choi DW. Neuroprotective effects of glutamate antagonists and extracellular acidity. Science 260: 1516–1518, 1993. 39. Li S and Stys PK. Mechanisms of ionotropic glutamate receptor-mediated excitotoxicity in isolated spinal cord white matter. J Neurosci 20: 1190–1198, 2000. 40. Matsumoto K, Graf R, Rosner G, Taguchi J, and Heiss WD. Elevation of neuroactive substances in the cortex of cats during prolonged focal ischemia. J Cereb Blood Flow Metab 13: 586–594, 1993. 41. Matute C, Alberdi E, Ibarretxe G, and Sanchez-Gomez MV. Excitotoxicity in glial cells. Eur J Pharmacol 447: 239–246, 2002. 42. Meldrum B. Possible therapeutic applications of antagonists of excitatory amino acid neurotransmitters. Clin Sci (Lond) 68: 113–122, 1985. 43. Molinaro P, Cuomo O, Pignataro G, Boscia F, Sirabella R, Pannaccione A, Secondo A, Scorziello A, Adornetto A, Gala R, Viggiano D, Sokolow S, Herchuelz A, Schurmans S, Di Renzo G, and Annunziato L. Targeted disruption of Na+/Ca2+ exchanger 3 (NCX3) gene leads to a worsening of ischemic brain damage. J Neurosci 28: 1179–1184, 2008. 44. Monnerie H and Le Roux PD. Glutamate receptor agonist kainate enhances primary dendrite number and length from immature mouse cortical neurons in vitro. J Neurosci Res 83: 944–956, 2006. 45. Obrenovitch TP. High extracellular glutamate and neuronal death in neurological disorders. Cause, contribution or consequence? Ann N Y Acad Sci 890: 273–286, 1999. 46. Obrenovitch TP. Neuroprotective strategies: voltage-gated Na+-channel down-modulation versus presynaptic glutamate release inhibition. Rev Neurosci 9: 203–211, 1998. 47. Obrenovitch TP, Urenjak J, Richards DA, Ueda Y, Curzon G, and Symon L. Extracellular neuroactive amino acids in the rat striatum during ischaemia: comparison between penumbral conditions and ischaemia with sustained anoxic depolarisation. J Neurochem 61: 178–186, 1993. 48. Obrenovitch TP, Urenjak J, Zilkha E, and Jay TM. Excitotoxicity in neurological disorders – the glutamate paradox. Int J Dev Neurosci 18: 281–287, 2000. 49. Olney JW and Sharpe LG. Brain lesions in an infant rhesus monkey treated with monsodium glutamate. Science 166: 386–388, 1969. 50. Pignataro G, Gala R, Cuomo O, Tortiglione A, Giaccio L, Castaldo P, Sirabella R, Matrone C, Canitano A, Amoroso S, Di Renzo G, and Annunziato L. Two sodium/ calcium exchanger gene products, NCX1 and NCX3, play a major role in the development of permanent focal cerebral ischemia. Stroke 35: 2566–2570, 2004. 51. Pohl D, Bittigau P, Ishimaru MJ, Stadthaus D, Hubner C, Olney JW, Turski L, and Ikonomidou C. N-Methyl-D-aspartate antagonists and apoptotic cell death triggered by head trauma in developing rat brain. Proc Natl Acad Sci USA 96: 2508–2513, 1999. 52. Randall RD and Thayer SA. Glutamate-induced calcium transient triggers delayed calcium overload and neurotoxicity in rat hippocampal neurons. J Neurosci 12: 1882–1895, 1992. 53. Riccio A, Ahn S, Davenport CM, Blendy JA, and Ginty DD. Mediation by a CREB family transcription factor of NGF-dependent survival of sympathetic neurons. Science 286: 2358–2361, 1999. 54. Rothman SM and Olney JW. Glutamate and the pathophysiology of hypoxic – ischemic brain damage. Ann Neurol 19: 105–111, 1986. 55. Rothstein JD, Patel S, Regan MR, Haenggeli C, Huang YH, Bergles DE, Jin L, DykesHoberg M, Vidensky S, Chung DS, Toan SV, Bruijn LI, Su ZZ, Gupta P, and Fisher PB. Beta-lactam antibiotics offer neuroprotection by increasing glutamate transporter expression. Nature 433: 73–77, 2005.

Failure of Ionotropic and Metabotropic Glutamate Antagonists

25

56. Shimada N, Graf R, Rosner G, Wakayama A, George CP, and Heiss WD. Ischemic flow threshold for extracellular glutamate increase in cat cortex. J Cereb Blood Flow Metab 9: 603–606, 1989. 57. Shimizu H, Graham SH, Chang LH, Mintorovitch J, James TL, Faden AI, and Weinstein PR. Relationship between extracellular neurotransmitter amino acids and energy metabolism during cerebral ischemia in rats monitored by microdialysis and in vivo magnetic resonance spectroscopy. Brain Res 605: 33–42, 1993. 58. Simon RP, Swan JH, Griffiths T, and Meldrum BS. Blockade of N-methyl-D-aspartate receptors may protect against ischemic damage in the brain. Science 226: 850–852, 1984. 59. Syntichaki P and Tavernarakis N. The biochemistry of neuronal necrosis: rogue biology? Nat Rev Neurosci 4: 672–684, 2003. 60. Walton MR and Dragunow I. Is CREB a key to neuronal survival? Trends Neurosci 23: 48–53, 2000. 61. Wang JM, Chao JR, Chen W, Kuo ML, Yen JJ, and Yang-Yen HF. The antiapoptotic gene mcl-1 is up-regulated by the phosphatidylinositol 3-kinase/Akt signaling pathway through a transcription factor complex containing CREB. Mol Cell Biol 19: 6195–6206, 1999. 62. Wollmuth LP and Sobolevsky AI. Structure and gating of the glutamate receptor ion channel. Trends Neurosci 27: 321–328, 2004. 63. Wong R. NMDA receptors expressed in oligodendrocytes. Bioessays 28: 460–464, 2006. 64. Xiong ZG, Pignataro G, Li M, Chang SY, and Simon RP. Acid-sensing ion channels (ASICs) as pharmacological targets for neurodegenerative diseases. Curr Opin Pharmacol 8: 25–32, 2008. 65. Young D, Lawlor PA, Leone P, Dragunow M, and During MJ. Environmental enrichment inhibits spontaneous apoptosis, prevents seizures and is neuroprotective. Nat Med 5: 448–453, 1999. 66. Zinkand WC, Thompson C, Salama AI, and Patel J. Excitatory amino acid-evoked calcium influx and calcium-dependent neurotoxicity in rat cortical cultures. Ann N Y Acad Sci 648: 355–357, 1992.

Mitochondrial Channels as Potential Targets for Pharmacological Strategies in Brain Ischemia Rosemary H. Milton and Michael R. Duchen

1 Introduction Ischemic stroke is a major cause of death and disability urgently requiring novel therapeutic approaches. Deprivation of oxygen and respiratory substrate during an ischemic episode has its most immediate impact on mitochondria, the body’s primary oxygen consumers. Mitochondrial function is in turn critically dependent on a range of carriers and channels, most of which remain inadequately understood at the genetic and molecular level. As mitochondrial channels are involved in energy production, calcium handling, generation of reactive oxygen species (ROS) and cell-death related signaling, it is evident that increased understanding of these processes and their regulation will help the search for novel rational approaches to therapy. Most mitochondrial ion channels and transporters have been less extensively characterised than their plasma-membrane-based counterparts, in part because they are more difficult to study directly. Much of the research we discuss here has been undertaken on mitochondria from organs other than the brain; these data must be extrapolated with care. This chapter will focus on the roles of the mitochondrial ATP-sensitive potassium channel (mitoKATP), uncoupling proteins (UCPs) and the permeability transition pore (PTP), although there are also significant but perhaps less intensively studied roles for other mitochondrial channels, transporters, and pumps (see Table 1).

M.R. Duchen (*) Department of Physiology, University College London, London, WC1E 6BT, UK e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_3, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

27

11q13

6p11.2-q12

Xq24

Uncoupling Protein 2 UCP2

Uncoupling Protein 4 UCP4

Uncoupling Protein 5 UCP5 (or BCMP1)

6 TM helices

6 TM helices

Controversial, reducing ROS production

Controversial, reducing ROS production

Controversial. Regulation of Kþ homeostasis, osmotic pressure and therefore structural integrity Controversial, reducing ROS production

Unknown, maybe plasma membrane KCa channel

Unknown, maybe plasma membrane KCa channel

Mitochondrial Ca sensitive potassium channel mitoKCa

6 TM helices

Controversial. Regulation of Kþ homeostasis, osmotic pressure and therefore structural integrity

Unknown, maybe plasma membrane KATP channel

Unknown, maybe plasma membrane KATP channel

Mitochondrial ATP sensitive potassium channel, mitoKATP

Other UCPs implicated and UCP4 expressed in brain

Other UCPs implicated and UCP4 expressed in brain

Implicated in brain I-R injury

Implicated in heart I-R injury, expressed in brain

Strongly implicated in brain I-R injury, in pre- and postconditioning Opened by cromokalim, pinacidil, diazoxide, BM10195. Blocked by glibenclamide, 5HD Opened by NS1619. Blocked by iberiotoxin ChTx and paxilline. Uncoupling can be simulated with protonophores such as FCCP Uncoupling can be simulated with protonophores such as FCCP Uncoupling can be simulated with protonophores such as FCCP

Table 1 Molecular and functional properties of ionic channels, uniporters, and exchangers in the mitochondria Selected current Channel, pump or Role in mitochondrial Implicated in ischemiapharmacological transporter Gene Structure function reperfusion? tools

[10, 13, 14]

[10–12]

[8–10]

[6;7]

[1–5]

Selected references

28 R.H. Milton and M.R. Duchen

VDAC1 5q31, VDAC2 10q22 VDAC3 8p11.2

SLC25A1 22q11.21, 2 5q31, 3 12q23 4 4q35 22q13.31

Voltage dependent anion channel VDAC

Adenine Nucleotide Translocator ANT

Unknown

Unknown (in mitochondria)

Calcium uniporter

Sodium-calcium exchanger

Translocator Protein TSPO (Peripheral Benzodiazepine Receptor, PBR)

10q21–10q23

Gene

Cyclphilin D CypD

Channel, pump or transporter

Unknown (in mitochondria)

Unknown

5 TM helices

Eight b-strands, two a-helices, and one 310 helix Beta-barrel with 13 betastrands and one alpha helix 6 TM helices

Structure

Role in calcium uptake by mitochondria Regulation of ionic concentrations

Putative role in PTP opening

Regulation of metabolism by regulating flux of metabolites, role in opening of PTP Exchanges ADP for ATP, role in opening of PTP

Role in opening of PTP

Role in mitochondrial function

Table 1 (continued)

Implicated in brain I-R injury, implicated at plasma membrane

PTP in general is implicated in brain I-R injury, TSPO has been ascribed a role in PTP opening and specifically in renal ischemia Implicated in heart I-R injury

PTP in general is implicated in brain I-R injury

PTP in general is implicated in brain I-R injury

Strongly implicated in brain I-R injury

Implicated in ischemiareperfusion?

Blocked by ruthenium red and Ru-360 Blocked by CGP37157

PK11195, diazepam, DPA-714

Blocked by bongkrekic acid

Blocked by Cyclosporine A

Selected current pharmacological tools

[25–27]

[23, 24]

[21, 22]

[20]

[19]

[15–18]

Selected references

Mitochondrial Channels in Brain Ischemia 29

Unknown (in mitochondria)

Sodium-proton exchanger

Unknown (in mitochondria)

Structure Regulation of ionic concentrations

Role in mitochondrial function Implicated in heart I-R injury, implicated at plasma membrane

Implicated in ischemiareperfusion? Blocked by cariporide, amiloride

Selected current pharmacological tools [28]

Selected references

1. Lauritzen, I., De, W., Jr., and Lazdunski, M. (1997) J. Neurochem. 69, 1570–1579 2. Nistico, R., Piccirilli, S., Sebastianelli, L., Nistico, G., Bernardi, G., and Mercuri, N. B. (2007) Int. Rev. Neurobiol. 82, 383–395 3. Bajgar, R., Seetharaman, S., Kowaltowski, A. J., Garlid, K. D., and Paucek, P. (2001) J. Biol. Chem. 276, 33369–33374 4. Gaspar, T., Snipes, J. A., Busija, A. R., Kis, B., Domoki, F., Bari, F., and Busija, D. W. (2008) J. Cereb. Blood Flow Metab. 28, 1090–1103 5. Lee, J. J., Li, L., Jung, H. H., and Zuo, Z. (2008) Anesthesiology 108, 1055–1062 6. Piwonska, M., Wilczek, E., Szewczyk, A., and Wilczynski, G. M. (2008) Neuroscience 153, 446–460 7. Xu, W., Liu, Y., Wang, S., McDonald, T., Van Eyk, J. E., Sidor, A., and O’Rourke, B. (2002) Science 298, 1029–1033 8. Mattiasson, G., Shamloo, M., Gido, G., Mathi, K., Tomasevic, G., Yi, S., Warden, C. H., Castilho, R. F., Melcher, T., Gonzalez-Zulueta, M., Nikolich, K., and Wieloch, T. (2003) Nat. Med. 9, 1062–1068 9. de Bilbao, F., Arsenijevic, D., Vallet, P., Hjelle, O. P., Ottersen, O. P., Bouras, C., Raffin, Y., Abou, K., Langhans, W., Collins, S., Plamondon, J., ves-Guerra, M. C., Haguenauer, A., Garcia, I., Richard, D., Ricquier, D., and Giannakopoulos, P. (2004) J. Neurochem. 89, 1283–1292 10. Pandya, J. D., Pauly, J. R., Nukala, V. N., Sebastian, A. H., Day, K. M., Korde, A. S., Maragos, W. F., Hall, E. D., and Sullivan, P. G. (2007) J. Neurotrauma 24, 798–811 11. Mao, W., Yu, X. X., Zhong, A., Li, W., Brush, J., Sherwood, S. W., Adams, S. H., and Pan, G. (1999) FEBS Lett. 443, 326–330 12. Liu, D., Chan, S. L., de Souza-Pinto, N. C., Slevin, J. R., Wersto, R. P., Zhan, M., Mustafa, K., de Cabo, R., and Mattson, M. P. (2006) Neuromolecular. Med. 8, 389–414 13. Sanchis, D., Fleury, C., Chomiki, N., Goubern, M., Huang, Q., Neverova, M., Gregoire, F., Easlick, J., Raimbault, S., Levi-Meyrueis, C., Miroux, B., Collins, S., Seldin, M., Richard, D., Warden, C., Bouillaud, F., and Ricquier, D. (1998) J. Biol. Chem. 273, 34611–34615 14. Nakase, T., Yoshida, Y., and Nagata, K. (2007) Neuropathology. 27, 442–447 15. Shiga, Y., Onodera, H., Matsuo, Y., and Kogure, K. (1992) Brain Res. 595, 145–148 16. Zhang, W. H., Wang, H., Wang, X., Narayanan, M. V., Stavrovskaya, I. G., Kristal, B. S., and Friedlander, R. M. (2008) Stroke 39, 455–462 17. Korde, A. S., Pettigrew, L. C., Craddock, S. D., Pocernich, C. B., Waldmeier, P. C., and Maragos, W. F. (2007) J. Neurotrauma 24, 895–908 18. Muramatsu, Y., Furuichi, Y., Tojo, N., Moriguchi, A., Maemoto, T., Nakada, H., Hino, M., and Matsuoka, N. (2007) Brain Res. 1149, 181–190 19. Shoshan-Barmatz, V., Israelson, A., Brdiczka, D., and Sheu, S. S. (2006) Curr. Pharm. Des 12, 2249–2270 20. Klingenberg, M. (2008) Biochim. Biophys. Acta

Gene

Channel, pump or transporter

Table 1 (continued)

30 R.H. Milton and M.R. Duchen

Table 1 (continued) 21. Doucet, C., Zhang, K., Desurmont, T., Hebrard, W., Scepi, M., Nadeau, C., Cau, J., Leyre, P., Febrer, G., Carretier, M., Richer, J. P., Papadopoulos, V., Hauet, T., Burucoa, C., and Goujon, J. M. (2007) Nephron Exp. Nephrol. 107, e1–11 22. Papadopoulos, V. (2004) Endocr. Res. 30, 677–684 23. Garcia-Rivas, G. J., Carvajal, K., Correa, F., and Zazueta, C. (2006) Br. J. Pharmacol. 149, 829–837 24. Zhang, S. Z., Gao, Q., Cao, C. M., Bruce, I. C., and Xia, Q. (2006) Life Sci. 78, 738–745 25. Zhang, Y. and Lipton, P. (1999) J. Neurosci. 19, 3307–3315 26. Santo-Domingo, J., Vay, L., Hernandez-Sanmiguel, E., Lobaton, C. D., Moreno, A., Montero, M., and Alvarez, J. (2007) Br. J. Pharmacol. 151, 647–654 27. Motegi, K., Tanonaka, K., Takenaga, Y., Takagi, N., and Takeo, S. (2007) Br. J. Pharmacol. 151, 963–978 28. Toda, T., Kadono, T., Hoshiai, M., Eguchi, Y., Nakazawa, S., Nakazawa, H., Higashijima, N., and Ishida, H. (2007) Am. J. Physiol Heart Circ. Physiol. 293, H3517–H3523

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2 Gene Structure, Structural Features, and Molecular Biology The molecular identity, structural features, and associated molecular biology of most mitochondrial channels remain unknown. All mitochondrial channels and transporters are thought to be encoded by nuclear DNA, however, so their identities should be accessible.

2.1 Mitochondrial ATP-Sensitive Kþ Channel, MitoKATP ATP-regulated Kþ channels, or Kþ flux, have been identified in isolated mitochondrial preparations by several groups. Evidence of a role for mitoKATP in ischemia-reperfusion injury is accumulating (for review see [1]), yet the molecular identity of the channel remains nebulous. While there is some indication that mitoKATP contains the same subunits as the plasma membrane KATP channel, the pharmacology differs (for review see [2]) and the mitochondrial channel has a smaller conductance [3]. The plasma membrane channel is well characterized; it is a hetero-octamer composed of four inwardly rectifying Kþ channels, either Kir6.1 or 6.2, and four sulfonylurea receptor subunits, either SUR1 or SUR2 in various combinations [4]. Kir6.1 and 6.2 and SUR2 have putative mitochondrial targeting sequences within their N-termini, and immunogold electron microscopy and Western blotting have suggested that they can be found in neuronal mitochondria [5]. However, there is doubt about the specificity of commercial antibodies to Kir6.1 [6], and Kir6.2 knockout mice have normal mitoKATP function in cardiac myocytes [7]. It remains possible, therefore, that mitoKATP is composed of entirely different proteins from the surface channel, or the same subunits but with some complex regulation or redundancy.

2.2 Uncoupling Proteins, UCPs In contrast to mitoKATP, the UCPs are a family of proteins with well-established molecular identities. UCPs 2–5 were named in light of their homology with the classical UCP of brown adipose tissue, UCP1, but analogous functions have proved more difficult to demonstrate. Respiring mitochondria export protons across the inner membrane using energy from substrate oxidation, which is ‘‘coupled’’ to the energy required for ATP synthesis, which occurs when these protons re-enter the mitochondria. UCP1 uncouples substrate oxidation from ATP synthesis by providing an alternative route for protons to re-enter the mitochondria (see Fig. 1). UCPs 2 and 3 were identified through their homology with UCP1 in 1997 [8, 9] and UCP4 and UCP5 (BCMP1) were identified in the next 2 years [10, 11]. All UCPs are members of the 35 strong family of mitochondrial anion carriers

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H+

33

H+ O2* Q

H+ H+

Intermembrane space

C

e-

e-

e-

eMatrix

O2 * + NADH NAD FADH2FAD

O2*

++

HO 4H+ + O2 2 ADP + Pi

I

II

III

H+

IV

H+

ATP

ATP Synthase

H+ H+

Q e-

C e-

e-

e-

+ ++ NADH NAD FADH2FAD

HO 4H+ + O2 2 ADP + Pi

I

II

III

IV

ATP

ATP Synthase Uncoupling

Fig. 1 Mitochondrial respiration. Mitochondrial production of ATP is dependent on the electron transport chain (complexes I-IV, and intermediaries including ubiquinone (Q) and cytochrome c (C)) and the ATP synthase. The electron transport chain transfers redox energy from NADH and FADH2 to oxygen in several stages. As electrons move down the chain, the release of energy is used to transport protons across the inner mitochondrial membrane. The translocation of protons generates an electrochemical gradient expressed primarily as a membrane potential. Protons then move down this electrochemical gradient through the ATP synthase, driving the generation of ATP from ADP and inorganic phosphate. Although electron transfer is generally efficient, electrons may transfer onto molecular oxygen, rather than onto the next stage of the chain, producing superoxide (O2*) which can then contribute to oxidative stress. Uncoupling mitochondria (lower panel) an alternative conductance for protons to re-enter the mitochondria, in competition with the proton path through the ATP synthase. The extrusion of protons by the electron transport chain is therefore uncoupled from ATP synthesis. To maintain the mitochondrial membrane potential and ATP generation, electrons are transferred more rapidly along the electron transport chain, so that the probability of electron leak is reduced. It has been proposed that such a mechanism may reduce oxidative stress.

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and contain three repeats of 100 amino acids, each containing two hydrophobic stretches which correspond to transmembrane helices. UCPs 2, 4 and 5 are expressed in the brain, and may therefore be of relevance to stroke. The ucp2ucp3 cluster in humans is 11q13 between DS11S916 and D11S911 [8]. The fact that the ucp2 gene is only 7 kb downstream from ucp3 possibly reflects a duplication event [12]. The transcription unit of ucp2 is made up of two untranslated exons followed by 6 exons encoding UCP2, which in humans is a 309 amino acid long protein with a mol. wt of 33218. UCP4 was identified through its 34% homology with UCP3, and has been mapped to 6p11.2–q12. Similar to the other UCPs, it has the characteristic 6 transmembrane domains, 3 mitochondrial transporter protein signature motifs, and a putative nucleotide-binding site [10]. UCP5 has 38% homology with UCP2, and is located on the X chromosome in man (Xq25–26) between the genetic markers DXS1206 and DXS1047 [13].

2.3 Mitochondrial Permeability Transition Pore, PTP PTP was first identified by Haworth and Hunter in a series of landmark papers [14–16] as a massive increase in permeability of the mitochondrial membrane following calcium overload and oxidative stress, reflecting the opening of a large conductance channel complex [17]. Until recently, it was held that this channel comprised a multipartite structure with several putative constituent proteins, including the adenine nucleotide translocase (ANT) and the voltagedependent anion channel (VDAC). Recent studies based on inactivation of the genes encoding the candidate proteins [18–20] have cast doubt on the roles of ANT and VDAC, but Cyclophilin D (CypD), the product of the Ppif gene, remains one of the more reliable candidates. The Ppif (peptidylprolyl isomerase) gene consists of 6 protein-encoding exons separated by five introns mapped to 10q21–q23 [21]. Mitochondria from CypD knockout mice do not exhibit typical PTP-like pore opening [18]. While VDAC has long been thought to be a constituent of PTP, mice lacking all 3 VDAC isoforms have now been shown to have intact PTP-like activity [19]. VDAC1 was originally mapped to the X chromosome in the interval Xq13–q21 and VDAC2 to chromosome 21, while VDAC3 maps to chromosome 12 [22]. However, others have mapped VDAC1 to chromosome 5q31 and VDAC2 to chromosome 10q22 [23]. VDAC forms barrels in the outer mitochondrial membrane, consisting of a transmembrane alpha helix and approximately 13 transmembrane beta strands, which enclose aqueous channels. ANT is thought to play a more regulatory role in PTP activity, since mitochondria from mice lacking ANT1 and 2 still exhibit pore opening [20]. The ANT has four isoforms in humans, all consisting of six transmembrane helices, with three repeats of two. Other proteins to have been implicated in PTP function are the peripheral benzodiazepine receptor (PBR; [24, 25]) and the

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mitochondrial phosphate carrier (PiC; [26]). The complex issue of PTP components and regulators will only be addressed fully when conditional knockout animals or siRNA are used.

3 Tissue and Cellular Distribution MitoKATP is expressed in the inner mitochondrial membrane. In situ hybridization and immunogold electron microscopy [5, 27] have suggested that it is expressed in neuronal mitochondria, but as mentioned, this is not incontrovertible. UCPs 2, 4 and 5 are expressed in mitochondria in the brain, with UCP2, interestingly, being strongly expressed in microglia [28]. The components of the PTP are also present in all cells, although there are variations in susceptibility to opening in different brain areas including throughout the brain [29]. All three VDAC isoforms are present in mitochondria from bovine, rabbit and rat brain [30], while ANT1 is thought to be the most expressed form in the brain [31].

4 Biophysical and Electrophysiological Properties and Regulatory Mechanisms MitoKATP was first described following patch clamping of mitoplasts [3]. The I–V relationship shows that current amplitude increases linearly with hyperpolarization in the negative voltage region and that the linear extrapolation shows a reversal potential at +27 mV, similar to EK which is +29.5 mV. Channel activation is not strongly voltage dependent, however. The conductance described in mitoplasts is consistent with the conductance described in channels reconstituted into artificial membranes [32]. The channels are inhibited by ATP, and activated by GTP and GDP, while modulated by PKC. Most interestingly, mitoKATP is possibly activated by oxidants, as would be generated during ischemia-reperfusion injury [33]. Patch clamping the inner mitochondrial membrane suggests PTP has a conductance of around 1.1 nS. The channel was activated by increased calcium concentrations, but not active in the presence of CsA or at voltages below –40 mV [34]. It has been suggested that the PTP channel has two different conductance states with different selectivities for anions and cations [35, 36] To the best of our knowledge, UCPs have never been subjected to electrophysiological analysis.

5 Physiological Properties MitoKATP is thought to play a role in regulation of mitochondrial Kþ concentration, and therefore mitochondrial volume regulation. Since the enzymes essential for mitochondrial function are present in the mitochondrial

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membrane, the maintenance of organelle volume and membrane configuration are crucial in respiratory function [37]. ‘Physiological’ functions of UCPs and PTP remain controversial, and both are more frequently spoken of in terms of pathology. UCPs are thought to reduce production of toxic ROS. It has also been proposed that UCPs 2 and 3 are involved in fatty acid export from mitochondria (for review see [38]), in the oxidative metabolism of glutamine [39] and in calcium uptake [40]. PTP is involved in cell death, whereby PTP opening could either dissipate the mitochondrial membrane potential and so impair production of ATP to the extent that the cell undergoes necrosis, or could cause the release of mitochondrial factors such as cytochrome c, which promote signaling pathways causing apoptosis. Less attention has been focused on less devastating roles for PTP; it has been suggested that it releases calcium accumulated by neuronal mitochondria during normal, non-toxic, cellular events [41]. The postulated components of PTP also have important physiological roles; ANT facilitates the exchange of ADP and ATP, while PBR is involved in cholesterol transport [42].

6 Pathophysiological Relevance in Stroke 6.1 MitoKATP Opening mitoKATP channels was initially thought to be protective following association with the phenomenon of ischemic pre-conditioning, wherein a transient and mild ischemia limits the extent of damage incurred following a more severe subsequent ischemia-reperfusion injury. In the heart, the mitoKATP channel openers, cromakalim and pinacidil, affect pre-conditioning and limit ischemiareperfusion injury [43], while physical (ischemic) pre-conditioning is prevented by blockade of mitoKATP channels with glibenclamide or 5-hydroxydecanoate (5HD) [44]. Confirming the pharmacology, it has also been shown that overexpressing Kir6.2 in mitochondria confers protection from hypoxia in HEK and HL-1 cells [45]. It has been argued on the basis of work in the heart that mitoKATP, rather than sarcolemmal KATP channels, is the target of these drugs, but the pharmacological selectivity is not absolute. It is thought, however, that expression of mitochondrial channels is likely to be more relevant in the brain [46]. In the brain, as in the heart, the mitoKATP channel opener cromakalim protects neurons in vitro against both oxygen and glucose deprivation (OGD) and glutamate excitotoxicity [47]. A more mitochondrial-specific opener of KATP channels (BMS 191095) reduced infarct volume in mice subjected to Middle Cerebral Artery Occlusion (MCAO) if given 24 h previously [48]. However, the role of these channels remains unclear as there is some suggestion that blocking, rather than opening, them is protective in hippocampal slices subjected to OGD (using glibenclamide) [49] and in mice subjected to MCAO (using 5HD) [50].

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The prevailing position, however, remains that opening mitoKATP channels prior to an ischemic injury is protective. MitoKATP channel opening could trigger pre-conditioning through the induction of ROS signaling pathways which have downstream effects responsible for protection, but it has also been argued that the channels themselves are the effectors of pre-conditioning and must remain open throughout ischemia (and perhaps reperfusion) to afford protection. It is still not clearly established whether channel openers cause an increase in ROS during pre-conditioning, and if they do, whether these effects are mediated through Kþ or an alternative action of the drug, perhaps directly on the mitochondrial respiratory chain [51–53]. The study most relevant to this chapter, using a mitochondria-specific KATP channel opener on neurons, found that the drug did not increase mitochondrial ROS production while inducing pre-conditioning, but did reduce the levels of ROS production experienced in subsequent glutamate excitotoxicity [54]. Moreover, mitoKATP-opening drugs could exert their effects through uncoupling, reducing ROS production and calcium influx into the mitochondria. Indeed, the mitoKATP channel openers, pinacidil and diazoxide, have mild uncoupling activity [46]. Conversely, it has been argued that the Kþ flux is not large enough to promote uncoupling, and if it were the mitochondria would rupture [55]. Confusingly, it has also been suggested that Kþ ions move in the opposite direction from that required for uncoupling, leaving neuronal mitochondria in the presence of diazoxide [50]. While pre-conditioning has proved a very instructive phenomenon, postconditioning is probably of greater interest to the patient who has just had a stroke. The observation that brief periods of ischemia following an initial and more severe ischemic episode can be protective was initially made in dog hearts [56] and has led to clinical trials in myocardial infarction [57] and replication of the principle in rat brain [58]. From a therapeutic perspective, routine anesthetics seem to offer an opportunity for pharmacological post-conditioning. Administration of isoflurane after ischemia in cortico-striatal slices reduced cell injury in a manner reversible by mitoKATP channel blockers [59]. The positive effects were seen when isoflurane was administered 10 min after OGD; it is obviously important to establish the window within which treatment was effective and examine these principles in vivo. Encouragingly, positive effects of post-conditioning 2 days after injury in rat brain have been reported [60].

6.2 UCPs Just as the endogenous protection afforded by mitoKATP could be exploited therapeutically, so the putative endogenous protective actions of UCPs might also be targeted. There is, however, a conceptual difference, in that the actions of UCPs are more likely to be mimicked than promoted. As mentioned, UCPs are widely believed to reduce ROS production by mitochondria by providing a

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leak pathway to protons, so reducing the mitochondrial membrane potential. Consequently, electrons move along the electron transport chain more rapidly, reducing the probability of electron leak to molecular oxygen to form radicals. However, uncoupling risks compromising ATP synthesis and could only be beneficial to the cell when ‘mild’ [61]. UCP2 is upregulated by pre-conditioning, and infarct volume following MCAO is reduced in mice over-expressing UCP2 [62]. Over-expression of UCP2 was linked to reduced levels of ROS within mitochondria. Interestingly, the converse study, in which UCP2 was knocked out, did not show the converse result. Mice lacking UCP2 are protected from ischemia-reperfusion injury in much the same way that mice over-expressing UCP2 are [28]. The authors attribute this to a compensation effect in these mice, which exhibited increased levels of anti-oxidant expression, again emphasizing the need for conditional knockout mice and siRNA, and conceptually complementary studies. It should also be remembered that other UCPs are expressed, and expressed non-uniformly, throughout the brain. Although there are limited agents available to modulate UCPs, synthetic protonophores exist. Mild uncoupling with the protonophore FCCP (carbonyl cyanide p-trifluoromethoxyphenylhydrazone) has been shown to be protective in ischemia-reperfusion injury in the heart and traumatic injury in brain [63, 64], but others have argued that uncoupling is protective neither in cultured neurons [65] nor in synaptosomes [66] since the level of uncoupling required to reduce ROS production causes ATP depletion to an injurious extent. Furthermore, uncoupling is likely to have major systemic side effects, unless it can be targeted to affected tissues, thus it seems that the process and consequences of uncoupling are currently not sufficiently understood to yield major drug targets.

6.3 PTP PTP opening has been most clearly implicated in the heart at reperfusion, which perhaps makes it more amenable to drug therapy than alternative sites, which need to be accessed earlier [67, 68]. The combination of calcium and ROS generation at reperfusion promotes opening of the PTP and cell death through apoptosis (via cytochrome c release) or necrosis. Cytochrome c release via PTP could also be involved in post-conditioning, since it is reduced in post-conditioned brain, relative to ischemic controls [68]. Since PTP activation requires ROS and calcium, it can be modulated through these triggers. Mitochondrially targeted anti-oxidants, such as NAC or SOD mimetics or mito-Q [69] show some promise as neuroprotective agents, but it is not clear if their presence is required before ischemia. One interesting means of reducing ROS involves small Scherzo-Shiller (SS) peptides, which contain ROS-scavenging tyrosine residues. SS-31 is protective against ischemiareperfusion injury in heart and brain [70]. Intriguingly, the authors claim the

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peptides might exert their effects not only through scavenging ROS, but through acting on PTP directly. Despite the potential problems with peptide treatments, these SS peptides are claimed to be suitable for administration by several routes and cross the blood brain barrier rapidly. PTP opening can also be curbed through limiting calcium overload by blocking the mitochondrial Naþ/Hþ exchanger. One such Naþ/Hþ blocker, cariporide, has been shown to be protective against OGD in neurons [71]. However, clinical trials of cariporide to reduce ischemic events in coronary bypass surgery actually showed these patients were at increased risk of stroke, accentuating the problems of translating principles from isolated preparations to patients, and from heart to brain [72]. The PTP can also be targeted directly. Cyclosporine A (CsA) has long been known to block PTP opening, acting on CypD. CypD is strongly implicated in stroke, since knockout mice have a significantly reduced infarct size following

2 K

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+

+

H

Outer mitochondrial membrane

H2O MitoKATP

UCP

PTP Inner mitochondrial membrane

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ROS production decreased

3

Membrane potential maintained, cytochromec release decreased

Fig. 2 Three potential mechanisms of neuroprotection. 1. Opening mitoKATP channels (using cromakalim, pinacidil, BMS191095). The result of this conductance is not clear, but could include maintenance of osmotic pressure, volume and therefore morphology and structural integrity. 2. Increasing proton conductance (mimicking UCPs; using FCCP). Increasing the membrane permeability to protons causes an increase in the speed at which electrons move along the electron transport chain. This reduces the probability of electron transfer to molecular oxygen to produce potentially harmful ROS. 3. Inhibiting PTP opening (using CsA) prevents collapse of the mitochondrial membrane potential and reduces cytochrome c release from mitochondria.

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MCAO [73] and CsA is protective in global cerebral ischemia in rats [74]. CsA is, interestingly, already a widely used immunosuppressant drug, yet it is not simple to apply its usage to stroke patients since it affects all cyclophilins. Nevertheless, it would be interesting to know if transplant patients treated with CsA had reduced risk of ischemia-reperfusion injury. Stavrovskaya and colleagues [75] screened drugs currently approved by the FDA as potential PTP inhibitors and found that promethazine was protective against OGD in neuronal cultures and MCAO in mice. A similar drug, nortriptyline, has now also been shown to have protective effects [76]. Given the roles of several preexisting drugs in blocking PTP, and the development of CsA derivatives such as FR901459 [77] one would hope that the theory could relatively rapidly be transferred to treatments (see Fig. 2).

7 Conclusions Mitochondria contain many proteins and processes which play a central role in defining the progression and outcome of ischemia-reperfusion injury, and so represent valuable potential therapeutic targets. Generally it is difficult to target intracellular channels without affecting processes throughout the cell, and if drugs are required to enter the mitochondria they must contend with the membrane potential [78]. So far, the most productive research on the role of mitochondria in ischemia-reperfusion has been performed in the heart, and heart and brain are not always as closely linked as we would like them to be. These problems, coupled with the general problems of delivering drugs across the blood–brain barrier and treating an acute condition like stroke with enough speed, have led to something of an impasse. However, the central role of mitochondria in ROS signaling, calcium regulation, energy production, and cell death pathways means there is likely to be some aspect of mitochondrial function which can be pharmacologically regulated to offer some hope to stroke patients. A variety of approaches, perhaps reviewing existing drugs with mitochondrial sites of action or perhaps synthesizing new peptides to target mitochondria, coupled with an increased understanding of the identities and functions of the vast range of mitochondrial channels, will hopefully one day lead to increased neuroprotection and preserved neurological function.

References 1. O’Rourke B. Evidence for mitochondrial K+ channels and their role in cardioprotection. Circ Res 94: 420, 2004. 2. Grover GJ and Garlid KD. ATP-Sensitive potassium channels: a review of their cardioprotective pharmacology. J Mol Cell Cardiol 32: 677, 2000. 3. Inoue I, Nagase H, Kishi K, and Higuti T. ATP-sensitive K+ channel in the mitochondrial inner membrane. Nature 352: 244, 1991.

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4. Aguilar-Bryan L and Bryan J. Molecular biology of adenosine triphosphate-sensitive potassium channels. Endocr Rev 20: 101, 1999. 5. Lacza Z, Snipes JA, Kis B, Szabo C, Grover G, and Busija DW. Investigation of the subunit composition and the pharmacology of the mitochondrial ATP-dependent K+ channel in the brain. Brain Res 994: 27, 2003. 6. Foster DB, Rucker JJ, and Marban E. Is Kir6.1 a subunit of mitoK(ATP)? Biochem Biophys Res Commun 366: 649, 2008. 7. Suzuki M, Sasaki N, Miki T, Sakamoto N, Ohmoto-Sekine Y, Tamagawa M, Seino S, Marban E, and Nakaya H. Role of sarcolemmal K(ATP) channels in cardioprotection against ischemia/reperfusion injury in mice. J Clin Invest 109: 509, 2002. 8. Fleury C, Neverova M, Collins S, Raimbault S, Champigny O, Levi-Meyrueis C, Bouillaud F, Seldin MF, Surwit RS, Ricquier D, and Warden CH. Uncoupling protein-2: a novel gene linked to obesity and hyperinsulinemia. Nat Genet 15: 269, 1997. 9. Boss O, Samec S, Paoloni-Giacobino A, Rossier C, Dulloo A, Seydoux J, Muzzin P, and Giacobino JP. Uncoupling protein-3: a new member of the mitochondrial carrier family with tissue-specific expression. FEBS Lett 408: 39, 1997. 10. Mao W, Yu XX, Zhong A, Li W, Brush J, Sherwood SW, Adams SH, and Pan G. UCP4, a novel brain-specific mitochondrial protein that reduces membrane potential in mammalian cells. FEBS Lett 443: 326, 1999. 11. Sanchis D, Fleury C, Chomiki N, Goubern M, Huang Q, Neverova M, Gregoire F, Easlick J, Raimbault S, Levi-Meyrueis C, Miroux B, Collins S, Seldin M, Richard D, Warden C, Bouillaud F, and Ricquier D. BMCP1, a novel mitochondrial carrier with high expression in the central nervous system of humans and rodents, and respiration uncoupling activity in recombinant yeast. J Biol Chem 273: 34611, 1998. 12. Pecqueur C, Cassard-Doulcier AM, Raimbault S, Miroux B, Fleury C, Gelly C, Bouillaud F, and Ricquier D. Functional organization of the human uncoupling protein-2 gene, and juxtaposition to the uncoupling protein-3 gene. Biochem Biophys Res Commun 255: 40, 1999. 13. Bouillaud F, Couplan E, Pecqueur C, and Ricquier D. Homologues of the uncoupling protein from brown adipose tissue (UCP1): UCP2, UCP3, BMCP1 and UCP4. Biochim Biophys Acta 1504: 107, 2001. 14. Hunter DR and Haworth RA. The Ca2+-induced membrane transition in mitochondria. I. The protective mechanisms. Arch Biochem Biophys 195: 453, 1979. 15. Hunter DR and Haworth RA. The Ca2+-induced membrane transition in mitochondria. III. Transitional Ca2+ release. Arch Biochem Biophys 195: 468, 1979. 16. Haworth RA and Hunter DR. The Ca2+-induced membrane transition in mitochondria. II. Nature of the Ca2+ trigger site. Arch Biochem Biophys 195: 460, 1979. 17. Zoratti M, Szabo I, and De Marchi U. Mitochondrial permeability transitions: how many doors to the house? Biochim Biophys Acta 1706: 40, 2005. 18. Baines CP, Kaiser RA, Purcell NH, Blair NS, Osinska H, Hambleton MA, Brunskill EW, Sayen MR, Gottlieb RA, Dorn GW, Robbins J, and Molkentin JD. Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature 434: 658, 2005. 19. Baines CP, Kaiser RA, Sheiko T, Craigen WJ, and Molkentin JD. Voltage-dependent anion channels are dispensable for mitochondrial-dependent cell death. Nat Cell Biol 9: 550, 2007. 20. Kokoszka JE, Waymire KG, Levy SE, Sligh JE, Cai J, Jones DP, MacGregor GR, and Wallace DC. The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature 427: 461, 2004. 21. Bowles KR, Zintz C, Abraham SE, Brandon L, Bowles NE, and Towbin JA. Genomic characterization of the human peptidyl-prolyl-cis-trans-isomerase, mitochondrial precursor gene: assessment of its role in familial dilated cardiomyopathy. Hum Genet 105: 582, 1999.

42

R.H. Milton and M.R. Duchen

22. Blachly-Dyson E, Baldini A, Litt M, McCabe ER, and Forte M. Human genes encoding the voltage-dependent anion channel (VDAC) of the outer mitochondrial membrane: mapping and identification of two new isoforms. Genomics 20: 62, 1994. 23. Messina A, Oliva M, Rosato C, Huizing M, Ruitenbeek W, van den Heuvel LP, Forte M, Rocchi M, and De Pinto V. Mapping of the human Voltage-Dependent Anion Channel isoforms 1 and 2 reconsidered. Biochem Biophys Res Commun 255: 707, 1999. 24. McEnery MW, Snowman AM, Trifiletti RR, and Snyder SH. Isolation of the mitochondrial benzodiazepine receptor: association with the voltage-dependent anion channel and the adenine nucleotide carrier. Proc Natl Acad Sci USA 89: 3170, 1992. 25. Azarashvili T, Grachev D, Krestinina O, Evtodienko Y, Yurkov I, Papadopoulos V, and Reiser G. The peripheral-type benzodiazepine receptor is involved in control of Ca2+induced permeability transition pore opening in rat brain mitochondria. Cell Calcium 42: 27, 2007. 26. Leung AW and Halestrap AP. Recent progress in elucidating the molecular mechanism of the mitochondrial permeability transition pore. Biochim Biophys Acta 1777: 946, 2008. 27. Zhou M, Tanaka O, Sekiguchi M, Sakabe K, Anzai M, Izumida I, Inoue T, Kawahara K, and Abe H. Localization of the ATP-sensitive potassium channel subunit (Kir6. 1/ uK(ATP)-1) in rat brain. Brain Res Mol Brain Res 74: 15, 1999. 28. de Bilbao F, Arsenijevic D, Vallet P, Hjelle OP, Ottersen OP, Bouras C, Raffin Y, Abou K, Langhans W, Collins S, Plamondon J, ves-Guerra MC, Haguenauer A, Garcia I, Richard D, Ricquier D, and Giannakopoulos P. Resistance to cerebral ischemic injury in UCP2 knockout mice: evidence for a role of UCP2 as a regulator of mitochondrial glutathione levels. J Neurochem 89: 1283, 2004. 29. Brustovetsky N, Brustovetsky T, Purl KJ, Capano M, Crompton M, and Dubinsky JM. Increased susceptibility of striatal mitochondria to calcium-induced permeability transition. J Neuropsychiatry 23: 4858, 2003. 30. Cesar MC and Wilson JE. All three isoforms of the voltage-dependent anion channel (VDAC1, VDAC2, and VDAC3) are present in mitochondria from bovine, rabbit, and rat brain. Arch Biochem Biophys 422: 191, 2004. 31. Doerner A, Pauschinger M, Badorff A, Noutsias M, Giessen S, Schulze K, Bilger J, Rauch U, and Schultheiss HP. Tissue-specific transcription pattern of the adenine nucleotide translocase isoforms in humans. FEBS Lett 414: 258, 1997. 32. Mironova GD, Skarga YY, Grigoriev SM, Negoda AE, Kolomytkin OV, and Marinov BS. Reconstitution of the mitochondrial ATP-dependent potassium channel into bilayer lipid membrane. J Bioenerg Biomembr 31: 159, 1999. 33. Fornazari M, de Paula JG, Castilho RF, and Kowaltowski AJ. Redox properties of the adenoside triphosphate-sensitive K+ channel in brain mitochondria. J Neurosci Nurs 86: 1548, 2008. 34. Loupatatzis C, Seitz G, Schonfeld P, Lang F, and Siemen D. Single-channel currents of the permeability transition pore from the inner mitochondrial membrane of rat liver and of a human hepatoma cell line. Cell Physiol Biochem 12: 269, 2002. 35. Campello S, De Marchi U, Szabo I, Tombola F, Martinou JC, and Zoratti M. The properties of the mitochondrial megachannel in mitoplasts from human colon carcinoma cells are not influenced by Bax. FEBS Lett 579: 3695, 2005. 36. De Marchi U, Basso E, Szabo I, and Zoratti M. Electrophysiological characterization of the Cyclophilin D-deleted mitochondrial permeability transition pore. Mol Membr Biol 23: 521, 2006. 37. Garlid KD and Paucek P. Mitochondrial potassium transport: the K(+) cycle. Biochim Biophys Acta 1606: 23, 2003. 38. Echtay KS. Mitochondrial uncoupling proteins – what is their physiological role? Free Radic Biol Med 43: 1351, 2007. 39. Criscuolo F, Mozo J, Hurtaud C, Nubel T, and Bouillaud F. UCP2, UCP3, avUCP, what do they do when proton transport is not stimulated? Possible relevance to pyruvate and glutamine metabolism. Biochim Biophys Acta 1757: 1284, 2006.

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40. Trenker M, Malli R, Fertschai I, Levak-Frank S, and Graier WF. Uncoupling proteins 2 and 3 are fundamental for mitochondrial Ca2+ uniport. Nat Cell Biol 9: 445, 2007. 41. Murchison D and Griffith WH. Mitochondria buffer non-toxic calcium loads and release calcium through the mitochondrial permeability transition pore and sodium/calcium exchanger in rat basal forebrain neurons. Brain Res 854: 139, 2000. 42. Papadopoulos V. In search of the function of the peripheral-type benzodiazepine receptor. Endocr Res 30: 677, 2004. 43. Grover GJ, McCullough JR, Henry DE, Conder ML, and Sleph PG. Anti-ischemic effects of the potassium channel activators pinacidil and cromakalim and the reversal of these effects with the potassium channel blocker glyburide. J Pharmacol Exp Ther 251: 98, 1989. 44. Gross GJ and Auchampach JA. Blockade of ATP-sensitive potassium channels prevents myocardial preconditioning in dogs. Circ Res 70: 223, 1992. 45. Ljubkovic M, Marinovic J, Fuchs A, Bosnjak ZJ, and Bienengraeber M. Targeted expression of Kir6.2 in mitochondria confers protection against hypoxic stress. J Physiol 577: 17, 2006. 46. Bajgar R, Seetharaman S, Kowaltowski AJ, Garlid KD, and Paucek P. Identification and properties of a novel intracellular (mitochondrial) ATP-sensitive potassium channel in brain. J Biol Chem 276: 33369, 2001. 47. Lauritzen I, De Weille JR, and Lazdunski M. The potassium channel opener (–)-cromakalim prevents glutamate-induced cell death in hippocampal neurons. J Neurochem 69: 1570, 1997. 48. Mayanagi K, Gaspar T, Katakam PV, Kis B, and Busija DW. The mitochondrial K(ATP) channel opener BMS-191095 reduces neuronal damage after transient focal cerebral ischemia in rats. J Cereb Blood Flow Metab 27: 348, 2007. 49. Nistico R, Piccirilli S, Sebastianelli L, Nistico G, Bernardi G, and Mercuri NB. The blockade of K(+)-ATP channels has neuroprotective effects in an in vitro model of brain ischemia, Int. Rev Neurobiol 82: 383, 2007. 50. Liu D, Slevin JR, Lu C, Chan SL, Hansson M, Elmer E, and Mattson MP. Involvement of mitochondrial K+ release and cellular efflux in ischemic and apoptotic neuronal death. J Neurochem 86: 966, 2003. 51. Andrukhiv A, Costa AD, West IC, and Garlid KD. Opening mitoKATP increases superoxide generation from complex I of the electron transport chain. Am J Physiol Heart Circ Physiol 291: H2067–H2074, 2006. 52. Facundo HT, Carreira RS, de Paula JG, Santos CC, Ferranti R, Laurindo FR, and Kowaltowski AJ. Ischemic preconditioning requires increases in reactive oxygen release independent of mitochondrial K+ channel activity. Free Radic Biol Med 40: 469, 2006. 53. Minners J, Lacerda L, Yellon DM, Opie LH, McLeod CJ, and Sack MN. Diazoxideinduced respiratory inhibition – a putative mitochondrial K(ATP) channel independent mechanism of pharmacological preconditioning. Mol Cell Biochem 294: 11, 2007. 54. Gaspar T, Snipes JA, Busija AR, Kis B, Domoki F, Bari F, and Busija DW. ROSindependent preconditioning in neurons via activation of mitoK(ATP) channels by BMS-191095. J Cereb Blood Flow Metab 28: 1090, 2008. 55. Garlid KD, Dos SP, Xie ZJ, Costa AD, and Paucek P. Mitochondrial potassium transport: the role of the mitochondrial ATP-sensitive K(+) channel in cardiac function and cardioprotection. Biochim Biophys Acta 1606: 1, 2003. 56. Zhao ZQ, Corvera JS, Halkos ME, Kerendi F, Wang NP, Guyton RA, and VintenJohansen J. Inhibition of myocardial injury by ischemic postconditioning during reperfusion: comparison with ischemic preconditioning. Am J Physiol Heart Circ Physiol 285: H579–H588, 2003. 57. Staat P, Rioufol G, Piot C, Cottin Y, Cung TT, L’Huillier I, Aupetit JF, Bonnefoy E, Finet G, Andre´-Foue¨t X, and Ovize M. Postconditioning the human heart. Circulation 112: 2143, 2005.

44

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58. Zhao H, Sapolsky RM, and Steinberg GK. Interrupting reperfusion as a stroke therapy: ischemic postconditioning reduces infarct size after focal ischemia in rats. J Cereb Blood Flow Metab 26: 1114, 2006. 59. Lee JJ, Li L, Jung HH, and Zuo Z. Postconditioning with isoflurane reduced ischemiainduced brain injury in rats. Anesthesiology 108: 1055, 2008. 60. Burda J, Danielisova V, Nemethova M, Gottlieb M, Matiasova M, Domorakova I, Mechirova E, Ferikova M, Salinas M, and Burda R. Delayed postconditionig initiates additive mechanism necessary for survival of selectively vulnerable neurons after transient ischemia in rat brain. Cell Mol Neurobiol 26: 1141, 2006. 61. Nicholls DG, Johnson-Cadwell L, Vesce S, Jekabsons M, and Yadava N. Bioenergetics of mitochondria in cultured neurons and their role in glutamate excitotoxicity. J Neurosci Nurs 85: 3206, 2007. 62. Mattiasson G, Shamloo M, Gido G, Mathi K, Tomasevic G, Yi S, Warden CH, Castilho RF, Melcher T, Gonzalez-Zulueta M, Nikolich K, and Wieloch T. Uncoupling protein-2 prevents neuronal death and diminishes brain dysfunction after stroke and brain trauma. Nat Med 9: 1062, 2003. 63. Brennan JP, Berry RG, Baghai M, Duchen MR, and Shattock MJ. FCCP is cardioprotective at concentrations that cause mitochondrial oxidation without detectable depolarisation. Cardiovasc Res 72: 322, 2006. 64. Pandya JD, Pauly JR, Nukala VN, Sebastian AH, Day KM, Korde AS, Maragos WF, Hall ED, and Sullivan PG. Post-Injury Administration of Mitochondrial Uncouplers Increases Tissue Sparing and Improves Behavioral Outcome following Traumatic Brain Injury in Rodents. J Neurotrauma 24: 798, 2007. 65. Johnson-Cadwell LI, Jekabsons MB, Wang A, Polster BM, and Nicholls DG. ‘Mild Uncoupling’ does not decrease mitochondrial superoxide levels in cultured cerebellar granule neurons but decreases spare respiratory capacity and increases toxicity to glutamate and oxidative stress. J Neurochem 101: 1619, 2007. 66. Tretter L and Adam-Vizi V. Uncoupling is without an effect on the production of reactive oxygen species by in situ synaptic mitochondria. J Neurochem 103: 1864, 2007. 67. Gomez L, Thibault H, Gharib A, Dumont JM, Vuagniaux G, Scalfaro P, Derumeaux G, and Ovize M. Inhibition of mitochondrial permeability transition improves functional recovery and reduces mortality following acute myocardial infarction in mice. Am J Physiol Heart Circ Physiol 293: H1654–H1661, 2007. 68. Wang JY, Shen J, Gao Q, Ye ZG, Yang SY, Liang HW, Bruce IC, Luo BY, and Xia Q. Ischemic postconditioning protects against global cerebral ischemia/reperfusion-induced injury in rats. Stroke 39: 983, 2008. 69. Sharma SS and Gupta S. Neuroprotective effect of MnTMPyP, a superoxide dismutase/ catalase mimetic in global cerebral ischemia is mediated through reduction of oxidative stress and DNA fragmentation. Eur J Pharmacol 561: 72, 2007. 70. Szeto HH. Mitochondria-targeted cytoprotective peptides for ischemia-reperfusion injury. Antioxid Redox Signal 10: 601, 2008. 71. Luo J, Chen H, Kintner DB, Shull GE, and Sun D. Inhibition of Na+/H+ exchanger isoform 1 attenuates mitochondrial cytochrome C release in cortical neurons following in vitro ischemia. Acta Neurochir Suppl 96: 244, 2006. 72. Mentzer RM Jr., Bartels C, Bolli R, Boyce S, Buckberg GD, Chaitman B, Haverich A, Knight J, Menasche P, Myers ML, Nicolau J, Simoons M, Thulin L, and Weisel RD. Sodium-hydrogen exchange inhibition by cariporide to reduce the risk of ischemic cardiac events in patients undergoing coronary artery bypass grafting: results of the EXPEDITION study. Ann Thorac Surg 85: 1261, 2008. 73. Schinzel AC, Takeuchi O, Huang Z, Fisher JK, Zhou Z, Rubens J, Hetz C, Danial NN, Moskowitz MA, and Korsmeyer SJ. Cyclophilin D is a component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia. Proc Natl Acad Sci USA 102: 12005, 2005.

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45

74. Akdemir G, Ergungor MF, Sezer M, Albayrak L, Daglioglu E, and Kilinc K. Therapeutic efficacy of intraventricular cyclosporine A and methylprednisolone on a global cerebral ischemia model in rats. Neurol Res 27: 827, 2005. 75. Stavrovskaya IG, Narayanan MV, Zhang W, Krasnikov BF, Heemskerk J, Young SS, Blass JP, Brown AM, Beal MF, Friedlander RM, and Kristal BS. Clinically approved heterocyclics act on a mitochondrial target and reduce stroke-induced pathology. J Exp Med 200: 211, 2004. 76. Zhang WH, Wang H, Wang X, Narayanan MV, Stavrovskaya IG, Kristal BS, and Friedlander RM. Nortriptyline protects mitochondria and reduces cerebral ischemia/ hypoxia injury. Stroke 39: 455, 2008. 77. Muramatsu Y, Furuichi Y, Tojo N, Moriguchi A, Maemoto T, Nakada H, Hino M, and Matsuoka N. Neuroprotective efficacy of FR901459, a novel derivative of cyclosporin A, in in vitro mitochondrial damage and in vivo transient cerebral ischemia models. Brain Res 1149: 181, 2007. 78. Yamada Y, Akita H, Kogure K, Kamiya H, and Harashima H. Mitochondrial drug delivery and mitochondrial disease therapy – an approach to liposome-based delivery targeted to mitochondria. Mitochondrion 7: 63, 2007.

Endoplasmic Reticulum Calcium Homeostasis and Neuronal Pathophysiology of Stroke Alexei Verkhratsky

1 Introduction Endoplasmic reticulum (ER) formed by the endomembrane is arguably the largest intracellular organelle. The ER is present in all types of neural cell, in neurones and in glia [1, 2]. Morphologically, the ER is represented by a system of microtubules and cisternae organised into a complex 3D network. In neurones, the ER extends from the nucleus and the soma to the dendritic arborisation and, through the axon, to presynaptic terminals. Although the lumen of the ER is internally continuous, the spatial segregation of enzymatic systems ascertains remarkable functional heterogeneity of intra-ER compartments. Indeed, the ER is a place where a multitude of critical cellular processes develop. The ER acts both as a cellular synthetic factory, where mRNA is translated into proteins and the latter undergo post-translational folding, and as an intracellular highway where many of these proteins are transported to their final destinations. In addition, the ER is the primary site of formation of phospholipids, glucosylphosphatidylinositols and leukotriens. Besides its synthetic and transporting functions, the ER acts as a complex signalling organelle, able to receive and produce signals, which are of crucial importance for cell activity and survival [3]. In particular, the ER is specifically involved in cellular Ca2+ signalling, being the largest dynamic store of Ca2+ that generates Ca2+ fluxes between the cytosol and the ER lumen in response to extracellular stimulation, thus producing and shaping cytosolic Ca2+ signals [4]. The Ca2+ movements within the ER are also of critical importance [5], as free Ca2+ concentration in the ER lumen ([Ca2+]L) control synthesis and folding of proteins and initiate many of ER output signals. A. Verkhratsky (*) The University of Manchester, Manchester, M13 9PT, UK; Institute of Experimental Medicine, ASCR, 142 20 Prague 4, Czech Republic e-mail: [email protected]

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The ER Ca2+ homeostasis and signalling (Fig. 1) is controlled by several molecular systems, which determine ion fluxes through the endomembrane [1, 4, 6]. These systems are (i) endomembrane Ca2+ pumps of the SERCA (Sarco(Endo)plasmic reticulum calcium ATP-ase) family; (ii) intracellular Ca2+ release channels (which include ryanodine receptors (RyRs), inositol-1,4,5-trisphosphate receptors (InsP3Rs), and may be some other Ca2+ permeable channels such as, e.g. NAADP receptors or TRPV1 channels); (iii) intra-ER Ca2+ buffers and (iv) other ion channels, which maintain the ER membrane potential by counter-balance currents produced by Ca2+ release from the store. The nature of these channels is largely unknown although recently cloned trimeric intracellular cation channels (TRICs) may be responsible for counter-balancing currents carried by monovalent cations [7].

Fig. 1 Molecular physiology of endoplasmic reticulum Ca2+ homeostasis/signalling. Calcium is taken up into the ER by energy-dependent transport accomplished by SERCA, thus building up concentration gradient between the ER lumen and the cytoplasm. Physiological stimulation opens Ca2+ release channels represented by ryanodine receptors (RyRs) and inositol 1,4,5-trisphosphate receptors (InsP3Rs) localised in the ER membrane. The ER membrane also contains cationic channels, which produce ion fluxes counter-balancing charge redistribution associated with Ca2+ release.

All components of the ER ion moving system are regulated by Ca2+ gradients across the ER membrane in a complex manner [8, 9]. The complement of the Ca2+-regulated channels and Ca2+ pumps converts the ER membrane into the excitable media, which is able to generate propagating waves of Ca2+ channels openings; these openings in turn underlie the well-known phenomena of propagating intra- and intercellular Ca2+ waves. In the following chapters I shall concentrate on the three components of Ca2+ homeostatic/signalling system of the ER, namely on SERCA pumps and Ca2+ release channels of the RyR and InsP3R types.

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2 Gene Structure 2.1 SERCA In vertebrates, the SERCA family is encoded by three distinct genes; each of the three genes has several splice variants. These genes are designated as ATP2A1–3; these produce SERCA1a/b, SERCA2a/b and SERCA3a/b/c/d/e isoforms [10].

2.2 Ryanodine Receptors Ca2+-gated Ca2+ release channels residing in the endomembrane are generally known as ryanodine receptors (RyRs) because plant alkaloid ryanodine was the first specific probe against these proteins. The RyR family is encoded by three major genes, the RyR1, RyR2 and RyR3 [11]; historically these subunits were also known as ‘‘skeletal muscle’’, ‘‘heart’’ and ‘‘brain’’ types, respectively. The RyR genes are very large, and comprise 100 exons. Diversity of RyRs also stems from splice variants of which 13 have been identified [12].

2.3 InsP3 Receptors The InsP3 receptor family comprises three homotetrameric isoforms encoded by distinct genes, InsP3R1, InsP3R2 and InsP3R3 [13]. Further diversity arises from alternative splicing of InsP3R1 (up to six variants) and heteromeric expression [14].

3 Structural Features and Molecular Biology 3.1 SERCA The overall structure of SERCA proteins includes four major domains (see [10] for details). The transmembrane or M-domain contains 10 transmembrane helices, which form Ca2+-binding sites of the pump; other three domains (A, P and N) are cytoplasmic. The A (or actuator) domain and P (phosphorylation) domains are coupled to the M domain whereas the N (or nucleotide-binding domain) is linked to the P domain.

3.2 Ryanodine Receptors Both Ca2+ release channels, the RyRs and the InsP3Rs, are large tetrameric channel proteins, which share a four-clover-leaf-like structure when observed

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by electron microscopy [11]. The RyRs are probably the largest channels discovered so far, the RyR1 monomers comprise 5032–5037 amino acid residues; RyR2 – 4969–4976 residues and RyR3 – 4872 residues, with the mol. wt. of each monomer around 500 kDa [15]. The tetrametic RyR channel complex has two portions: the large (80% of the whole protein) cytosolic (also known as ‘‘head’’) with dimensions of 29 29 12 nm and a central or transmembrane portion (comprised of four transmembrane domains), which extends by 7 nm from the cytosolic domain [16]. This part also forms a central hole of about 2 nm in diameter, where the actual channel pore supposedly resides [17].

3.3 InsP3 Receptors The overall topology of InsP3 receptor is very similar to RyRs; the InsP3Rs are homotetramers, although the monomers are much smaller with mol.wt. around 260 kDa. The InsP3R1 comprises 2749 amino acid residues, the InsP3R2 – 2701 and InsP3R3 – 2671 amino acid residues. The InsP3Rs also have a large cytoplasmic ‘‘head’’-like structure, which contains the InsP3-binding site and a relatively small channel-forming part constructed from six transmembrane domains [13, 18].

4 Tissue and Cellular Distribution 4.1 SERCA The SERCA2b isoform is ubiquitously expressed in all cellular elements of the nervous system, in central and peripheral neurones, in astro- and oligodendrocytes and in microglia [19]. The SERCA 2a and SERCA3 are found almost exclusively in cerebellar Purkinje neurones [1, 19]. The role of SERCA3 isoform remains unresolved; genetic deletion of SERCA3 does not result in any obvious phenotypic change, save minor deficits in relaxation of vascular and tracheal smooth muscles [20]. Within the cells the SERCA pumps are distributed throughout the ER membrane; this distribution can be visualised in living cells by using fluorescent markers such as fluorescent thapsigargin.

4.2 Ryanodine Receptors All three types of RyRs are present in neurones, although expression of the RyR2 subtype dominates. Many neurones co-express either two or three RyR subtypes. The RyR1 is heavily expressed in cerebellar Purkinje neurones whereas RyR3 is found in hippocampal structures, the corpus striatum and the diencephalon [21, 22, 23, 24, 25, 26]. Within the nerve cells the RyRs are

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expressed almost in every part: the RyRs were detected in neuronal somatas, in axons of hippocampal [22], in cerebellar mossy fibres [26] and in presynaptic terminals [27]. RyRs are also heavily expressed in the dendritic tree; their expression is particularly high in dendritic spines of hippocampal CA1 neurones, where they represent the dominant Ca2+ release channel [22]. The expression of RyRs is developmentally regulated. Throughout the embryonic stage, RyR1 mRNA levels are highest in the rostral cortical plate whereas RyR3 mRNA are most prominent in the caudal cortical plate and hippocampus. Relatively low expression of RyR2-specific mRNA was detected in the diencephalon and the brainstem. However, from postnatal day 7 onwards, RyR2 mRNA gradually became the major isoform in many brain regions, while RyR1 mRNA became prominent in the dentate gyrus and in the Purkinje cell layer. Postnatal down-regulation in the caudal cerebral cortex restricted RyR3 mRNA expression to the hippocampus, particularly the CA1 region [28].

4.3 InsP3 Receptors All three subtypes of InsP3Rs were identified throughout the brain, with particularly high expression in Purkinje neurones and in somatas of CA1 cells [18, 22]. The InsP3R1 is the most abundant subtype in the brain whereas InsP3R2 dominates in the spinal cord and is also highly expressed in glial cells. The InsP3R3 are expressed to a much lesser extend; being present in cerebellar granule layer and the medulla [29]. Interestingly, InsP3Rs display cell-specific intracellular distribution. For example, InsP3R1 is found in dendrites, dendritic spines, cell bodies, axons and axonal terminals of cerebellar Purkinje cells, but is mostly confined to somatic regions and proximal dendrites in other neurones [29]. The InsP3R3 is very much concentrated in the neuropil and neuronal terminals [29].

5 Biophysical and Electrophysiological Properties 5.1 Ryanodine Receptors All three RyRs are classical ligand-gated channels, which are activated by Ca2+ from the cytosolic side; the rank order of RyRs sensitivity to [Ca2+]i is RyR1 > RyR2 > RyR3 [30, 31]. The RyRs reconstituted in lipid bilayer membranes and studied under voltage clamp behaved as not perfectly selective Ca2+ channels (PCa2+/PK+ 6–7) with fast activation kinetics (t of activation 0.5–1 ms) and a large conductance (100–500 pS, depending on the nature of ion carrier[16]). Physiological unitary Ca2+ current through RyRs was estimated at 0.5–0.9 pA at 2 mM intraluminal Ca2+ concentration [32].

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5.2 InsP3 Receptors The InsP3 receptors are activated by second messenger InsP3 produced by phospholipase C; this enzyme in turn appears as a part of the phosphoinositide signal transduction cascades, coupled to an extended family of metabotropic receptors abundantly expressed in neural cells. The conductance of InsP3Rs has been estimated at 10–26 pS, being thus considerably smaller when compared with RyRs. The InsP3Rs were also reported to show multiple (up to four) conductance states [33, 34, 35]. The activation of InsP3Rs is synergistically regulated by both InsP3 and cytosolic Ca2+. The most important difference between InsP3R subtypes lies in their different sensitivity to cytosolic Ca2+ concentration: the InsP3R1 has the ‘‘classical’’ bell-shape dependence on [Ca2+]i with a maximal activation at 300–400 nM [33]; the InsP3R2 and InsP3R3 are also stimulated by increase in cytosolic Ca2+, but do not show inhibition at high [Ca2+]i [36].

6 Regulatory Mechanisms 6.1 SERCA Ca2+ ions in the ER lumen act as the main regulator of SERCA pump. Already early experiments on ER vesicles found that an increase in intra-vesicular Ca2+ effectively inhibits Ca2+ uptake [37, 38]. Experiments on cellular preparations when [Ca2+]L was monitored by fluorescent probes clearly demonstrated that a decrease in [Ca2+]L markedly increased the velocity of SERCA-dependent Ca2+ uptake [8, 39, 40]. This peculiar regulation involves interactions between intra-ER proteins calreticulin and Erp57 with SERCA pumps. At high [Ca2+]L the calreticulin–Erp57 complex binds to and inhibits the activity of SERCA whereas lowering of [Ca2+]L results in the protein complex dissociation from the pump, which in turn increased the activity of the latter [41, 42, 43].

6.2 Ryanodine Receptors Cyclic ADP-ribose (cADPR). In nerve cells, cADPR does not open RyRs directly, yet it does potentiate Ca2+-induced Ca2+ release. Intracellular injections of cADPR did not affect resting [Ca2+]i but increase [Ca2+]i transients induced by depolarization [44, 45, 46, 47]. This effect was blocked by high concentrations of ryanodine or by conditioning emptying of the ER stores by incubations with 10–20 mM caffeine, thus suggesting cADPR interaction with RyRs. The cADPR can act as a second messenger, production of which in neural cells could be controlled by muscarinic cholinoreceptors [48], b-adrenoreceprtors [49],

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metabotropic glutamate receptors [50] or NO/cGMP/cGMP-dependent protein kinase cascade [49], although the details of the cADPR-dependent signalling cascades are far from being fully characterized. Nicotinic acid adenine dinucleotide phosphate (NAADP). Although the main action of NAADP is believed to be mediated through specific receptors present in the ER membrane [51], there are indications that NAADP can directly activate RyRs [52,53]. Calexcitin. Calexcitin, a Ca2+ and GTP-binding protein isolated from neurones of conditionally trained Hermissenda [54], is an endogenous modulator of neuronal RyRs and CICR. Calexitin binds to RyRs in a Ca2+-dependent manner and facilitates Ca2+-induced Ca2+ release [55, 56]. Glutamate. Glutamate may influence the RyRs and CICR through direct interactions between metabotropic GluRs and RyRs, which also involved Homer proteins [57]. This mechanism hitherto was experimentally demonstrated in cerebellar granule neurones. In other type of neurones, in cells isolated from the avian nucleus magnocellularis, activation of mGluRs inhibited both RyRs and InsP3Rs by yet unidentified pathway [58]. Neurotrophins. Several neurotrophins were reported to affect Ca2+ release mediated through RyRs in neuronal preparations. For example, NGF was reported to activate RyR-dependent Ca2+ release in cultured cerebellar neurones through activation of p75 receptors [59]. Another neurotrophin, NT-3, potentiated Ca2+ release via RyR/InsP3Rs in neuromuscular junction [60] whereas its cousin NT-4 enhanced CICR in pyramidal neurones from the visual cortex [61]. Finally, neurotrophins may regulate the expression of RyRs; for example NGF markedly up-regulated RyRs in cultured adrenal chromaffin cells [62] whereas BDNF did the same to cerebellar granule neurones [63]. Nitric oxide. Nitric oxide can also act as an endogenous regulator of RyRs [64]. NO activated RyRs in hippocampal CA1 neurones [65]. Activation of RyR-mediated Ca2+ release may also have pathological relevance: for instance NO-mediated Ca2+ release via RyRs could be responsible for Il-1b-induced pyrogenic effects [66]; similarly NO-induced activation of Ca2+ release from a ryanodine-sensitive store can be involved in hypoxic [Ca2+]i rises in brain slices [67].

6.3 InsP3 Receptors Protein kinases and calcineurin. Regulation of InsP3Rs is generally achieved through phosphorylation/dephosphorylation of the receptor molecule. Phosphorylation of the InsP3R, which results in enhanced channel activity, is catalysed by protein kinases A, C and G [68, 69] and by tyrosine kinase [70]. Dephosphorylation of InsP3R reduces the channel activity; this is catalysed by the Ca2+/calmodulin-dependent phosphatase calcineurin [71], which also may regulate the expression of InsP3R at the transcriptional level [72].

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Calmodulin. Calmodulin specifically inhibits InsP3R type 1, which has a respective high-affinity-binding site [73]. Caldendrin. Cytoplasmic Ca2+-binding proteins caldendrins possess dual effect on InsP3Rs. A long-splice variant of caldendrin, when expressed in PC12 or HeLa cells, inhibited IICR triggered by activation of histamine and purinoceptors [74]. Another, shorter splice variant of caldendrin, however, activated the InsP3Rs, was shown to bind specifically to all three InsP3R subtypes. This binding activated InsP3Rs, suggesting the possibility of IICR initiation even in the absence of the natural agonist [75]. Immunophilin FK 506-binding protein 12 (FKBP12). FKBP12 directly associates with InsP3R1; this complex being disrupted upon the addition of the immunodepressants FK506 or rapamycin. Dissociation of InsP3R1–FKBP12 causes an increase of Ca2+ flux through the channel (by increasing channel open time) that can be reversed by adding FKBP12 [76, 77].

7 Physiological Properties 7.1 SERCA The ER plays a dual role in Ca2+ homeostasis and signalling being simultaneously a ‘‘sink and source’’ for Ca2+ ions [78]. This double function is essentially maintained by SERCA pumps, as their activity provide for high intra-ER Ca2+ content, which in turn forms Ca2+ concentration gradient between ER lumen and the cytosol. In turn, the level of [Ca2+]L determines the mode for ER function. Low levels of [Ca2+]L promote Ca2+ buffering whereas the high [Ca2+]L facilitates Ca2+ release [1].

7.2 Ryanodine Receptors The main function of RyRs is to amplify cytosolic Ca2+ signals arising either from plasmalemmal Ca2+ entry or through Ca2+ release via InsP3Rs; this RyR activation is manifested in Ca2+-induced Ca2+ release or CICR [79, 80]. On the molecular level, RyRs, similarly to other ion channels, demonstrate spontaneous openings, which produce elementary Ca2+ release events, generally referred to as ‘‘sparks’’ [81]. These sparks were observed in variety of muscle preparations and were also identified in neuronal cells [27, 82, 83]. Increase in [Ca2+]i or addition of caffeine increases the frequency of sparks until the threshold is reached and regenerative process of CICR ensues. The CICR has been identified and characterised in both peripheral and central neurones (see [1] for review). In various neurones the CICR can be either graded by Ca2+ entry, or can have an ‘‘all or none’’ regenerative behaviour [40, 84]. Importantly, the CICR process, although significantly

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amplifying cytosolic Ca2+ signals produces very minor changes in [Ca2+]L: the free Ca2+ concentration within the ER lumen is rarely reduced by more than 5–10% of the resting level [40]. Regenerative opening of RyRs also underlies propagating ER membrane excitation, which results in formation of intracellular Ca2+ waves [1].

7.3 InsP3 Receptors The InsP3Rs are part of signal transduction cascade, which comprises the plasmalemmal metabotropic receptors (represented by a wide variety of receptors, including mGluRs, P1 and P2Y purinoceptors, muscarinic cholinoreceptors and many more), coupled to phospholipase C (PLC) via G-proteins. Physiological activation of metabotropic receptors by appropriate neurotransmitters results in activation of PLC, that in turn produces second messengers diacylglycerol and InsP3 [85]. In neural cells metabotropic receptors are activated by neurotransmitters and hormones, which results in generation of InsP3induced Ca2+ release or IICR. The IICR in turn can be either spatially localised, which results in generation of microdomains of increased [Ca2+]i, or in certain conditions, trigger global [Ca2+]i response [1]. In neurones local [Ca2+]i microdomains produced by IICR play an important role in synaptic plasticity [86, 87]. In addition to InsP3, the InsP3Rs are regulated by [Ca2+]i and the increase in the latter facilitates the IICR. As a result local increases in [Ca2+]i may trigger Ca2+ release in the presence of constant levels of InsP3 or else assist in recruiting neighbouring InsP3Rs thus creating a propagating Ca2+ waves through the so-called CICR-assisted IICR [1, 88]. Importantly, this property may play a role of coincidence detector, when pairing of metabotropic receptors activation with plasmalemmal Ca2+ entry (resulting e.g. from backpropagating action potentials) synergistically increases the IICR and greatly potentiates Ca2+ signals [88, 89].

8 Pathophysiological Relevance in Stroke Conceptually, ischaemia and stroke trigger dysregulation of Ca2+ homeostasis in neural cells. As a result, cytoplasm became overloaded with Ca2+; an increased [Ca2+]i in turn triggers various death subroutines, which result in neuronal demise via either necrotic or apoptotic way [90]. Part of [Ca2+]i elevation in ischaemic conditions is due to ER Ca2+ release [91, 92]. For example, massive (30 mM) increases in [Ca2+]i following an interruption in blood flow observed in hippocampal neurones were substantially (5 times) reduced by inhibition of the ER with TG [93, 94]. In acute slices prepared from striatum hypoxia induced huge Ca2+ release from the ER [67].

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Similarly, in cultured hippocampal neurones treated with NaCN to mimic anoxic environment, a large [Ca2+]i increase, sensitive to dantrolene and ruthenium red, was observed [95]. Inhibition of RyRs by dantrolene significantly reduced neuronal death in the hippocampus in adult rats, subjected to global cerebral ischaemia [96]. In white matter Ca2+ release from the ER is also involved in producing the ischaemic damage. That is [Ca2+]i increase, which mediates axonal damage, persisted even when Ca2+ ions were removed from the external solutions; at the same time inhibition of RyRs had clear protective effect [97]. Changes in the ER Ca2+ homeostasis and partial depletion of the ER from free Ca2+ may also play an important role in defining the fate of neural cells after stroke and ischaemia. It is well known that decrease in [Ca2+]L can trigger the ER-stress response, which is characterised by an accumulation of unfolded proteins and generation of an array of death signals [3, 98, 99, 100]. The link between ischaemia-induced changes in ER Ca2+ homeostasis and intra-ER chaperone activity in neurones has been identified. It turned out that the oxygen-regulated protein of 150 kDa (orp150) is up-regulated in astrocytes following ischaemic stress, which may explain astroglia resistance to ischaemic conditions. Overexpression of orp150 in cultured rat cortical neurones protected them against ischaemia [101]. Transgenic mice expressing orp150 under the control of a platelet-derived growth factor B-chin promoter displayed high levels of orp150 in the cortex, hippocampus and cerebellum and were much more resistant to global cerebral ischaemia compared to wild-type controls [101].

9 Pharmacological Modulation 9.1 SERCA Pumps The most potent and specific (KD 20 nM), though irreversible SERCA inhibitor is thapsigargin [102]. The cyclopiazonic acid blocks SERCA at concentrations 20–50 mM, and its action is reversible [1].

9.2 Ryanodine Receptors Caffeine. Caffeine is the most popular pharmacological probe for studying RyR-mediated Ca2+ release in cellular preparations. Caffeine rapidly penetrates the plasmalemma, and equilibrates within intracellular compartments. Caffeine stimulates RyRs, at low (1–3 mM) by sensitising the channel to cytosolic Ca2+ whereas at higher (10–20 mM) concentrations caffeine opens the RyR channel and keeps them activated for as long as the drug is present in the system (see [1] for review).

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Ryanodine. Ryanodine action on the RyR is concentration dependent. At low concentrations (1–5 mM), ryanodine promotes RyR channel opening in a sub-conducting (40–60% as compared with a normal) state, which associated with significant (about 20 times) increase in the channel open time. At high concentrations (50 –100 mM) ryanodine completely blocks RyRs [103, 104]. Ruthenium red. Ruthenium red blocks RyRs at concentrations between 1 and 20 mM [105]. At the single channel level, ruthenium red reduces Po by prolonging the closed state of the RyR. It has to be noted that ruthenium red is not very specific as it also inhibits voltage-gated Ca2+ channels, epithelial Ca2+ channels ECaC1, TRPV channels and mitochondrial calcium uptake (see [1] for review). Dantrolene. Dantrolene reduces the open probability of RyRs; the effective drug concentrations are 10–90 mM [106]. General anaesthetics. Halothane and enflurane increase Po of RyR channels, and thus, enhance the Ca2+-induced Ca2+ release [105]. Local anaesthetics. Most of the local anaesthetics, such as procaine, tertracaine, lidocaine, prilocaine, the quaternary amines QX 572 and QX 314 and benzocaine, inhibit RyR-mediated Ca2+ release in millimolar concentrations [105]. Heparin. Heparin stimulates Ca2+ release [107] by increasing channel openings with EC50 0.23 mg/ml in a Ca2+-dependent manner [108, 109].

9.3 InsP3 Receptors Heparin. Heparin blocks the activity of InsP3-gated channels incorporated into lipid bilayers [108] and effectively inhibits IICR in many types of non-excitable and excitable cells, when administered intracellularly. Caffeine. Caffeine at 10–20 mM concentrations is extremely potent inhibitor of InsP3Rs [110]. Direct electrophysiological investigations of purified InsP3Rs demonstrated effective inhibition of unitary InsP3R current. The KD of caffeine action on InsP3R currents was 1.6 and 10 mM completely blocked unitary InsP3R currents [111]. 2-Aminoethoxydiphenyl borate (2-APB). 2-APB inhibits IICR with IC50 40 mM [112, 113]. At the same time, however, 2-APB lacks selectivity as it affects not only IICR but also Ca2+ pumps and store-operated Ca2+ entry [114, 113]. Xestospongines. Xestospongines (Xe’s) belong to a group of macrocyclic bis-1-oxaquinolizidines isolated from the Australian sponge, Xestospongia species [115]. Some of these agents, namely XeA, C and D are membrane-permeable blockers of IICR; of these, XeC was the most potent inhibitor with a KD of 360 nM. Further experiments have demonstrated that XeC blocks [Ca2+]i transients mediated by metabotropic pathways in the PC12 cell line and in primary rat cortical astrocytes. Incubation of cells with 5–20 mM XeC effectively abolished [Ca2+]i responses to bradykinin and carbachol [115]. At the same time XeC was also reported to inhibit SERCA pumps [116].

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Adenophostines. Adenophostines A and B, isolated from metabolites of fungi Penicillium brevicompactum [117], are extremely potent (10–100 times more potent than InsP3) agonists of InsP3Rs [118].

10 Conclusions Endoplasmic reticulum plays a central role in calcium homeostasis and signalling in neural cells. It acts as a dynamic calcium store, able to accumulate, store and release Ca2+ ions in response to physiological stimulation. These main functions of the ER are supported by highly coordinated molecular cascades that produce Ca2+ release (RyRs and InsP3Rs) and Ca2+ uptake (SERCA pumps). Activity of these molecular cascades is regulated by Ca2+ gradients across the ER membrane, which ascertains a great homeostatic potential of the ER-Ca2+-handling system. Intra-ER Ca2+ levels also control the activity of numerous enzymatic processes responsible for synthesis, post-translational modification and trafficking of proteins. Ischaemia and stroke disrupt ER Ca2+ homeostasis, triggering Ca2+-dependent death programmes and initiating ER stress. Although the involvement of the ER in pathophysiology of stroke is generally accepted, the pathological potential of ER Ca2+ homeostatic/signalling cascades remains virtually uncharacterised, while clinical pharmacology of these pathways is nonexistent. Further investigations of pathophysiology of ER Ca2+ homeostasis/signalling systems may open a conceptually new line for developing anti-stroke therapeutic strategies.

References 1. Verkhratsky A. Physiology and pathophysiology of the calcium store in the endoplasmic reticulum of neurons. Physiol Rev 85: 201–279, 2005. 2. Park MK, Choi YM, Kang YK, and Petersen OH. The endoplasmic reticulum as an integrator of multiple dendritic events. Neuroscientist 14: 68–77, 2008. 3. Berridge MJ. The endoplasmic reticulum: a multifunctional signaling organelle. Cell Calcium 32: 235–249, 2002. 4. Meldolesi J. Rapidly exchanging Ca2+ stores in neurons: molecular, structural and functional properties. Prog Neurobiol 65: 309–338, 2001. 5. Petersen OH and Verkhratsky A. Endoplasmic reticulum calcium tunnels integrate signalling in polarised cells. Cell Calcium 42: 373–378, 2007. 6. Petersen OH, Petersen CC, and Kasai H. Calcium and hormone action. Annu Rev Physiol 56: 297–319, 1994. 7. Yazawa M, Ferrante C, Feng J, Mio K, Ogura T, Zhang M, Lin PH, Pan Z, Komazaki S, Kato K, Nishi M, Zhao X, Weisleder N, Sato C, Ma J, and Takeshima H. TRIC channels are essential for Ca2+ handling in intracellular stores. Nature 448: 78–82, 2007. 8. Burdakov D, Petersen OH, and Verkhratsky A. Intraluminal calcium as a primary regulator of endoplasmic reticulum function. Cell Calcium 38: 303–310, 2005.

ER Calcium Homeostasis & Neuronal Pathophysiology of Stroke

59

9. Burdakov D and Verkhratsky A. Biophysical re-equilibration of Ca2+ fluxes as a simple biologically plausible explanation for complex intracellular Ca2+ release patterns. FEBS Lett 580: 463–468, 2006. 10. Wuytack F, Raeymaekers L, and Missiaen L. Molecular physiology of the SERCA and SPCA pumps. Cell Calcium 32: 279–305, 2002. 11. Hamilton SL. Ryanodine receptors. Cell Calcium 38: 253–260, 2005. 12. George CH, Rogers SA, Bertrand BM, Tunwell RE, Thomas NL, Steele DS, Cox EV, Pepper C, Hazeel CJ, Claycomb WC, and Lai FA. Alternative splicing of ryanodine receptors modulates cardiomyocyte Ca2+ signaling and susceptibility to apoptosis. Circ Res 100: 874–883, 2007. 13. Bezprozvanny I. The inositol 1,4,5-trisphosphate receptors. Cell Calcium 38: 261–272, 2005. 14. Nucifora FC Jr., Li SH, Danoff S, Ullrich A, and Ross CA. Molecular cloning of a cDNA for the human inositol 1,4,5-trisphosphate receptor type 1, and the identification of a third alternatively spliced variant. Brain Res Mol Brain Res 32: 291–296, 1995. 15. Kostyuk P and Verkhratsky A. Calcium Signalling in the Nervous System. Chichester: Wiley & Sons, 1995. 16. Fill M and Copello JA. Ryanodine receptor calcium release channels. Physiol Rev 82: 893–922, 2002. 17. Radermacher M, Wagenknecht T, Grassucci R, Frank J, Inui M, Chadwick C, and Fleischer S. Cryo-EM of the native structure of the calcium release channel/ryanodine receptor from sarcoplasmic reticulum. Biophys J 61: 936–940, 1992. 18. Furuichi T, Kohda K, Miyawaki A, and Mikoshiba K. Intracellular channels. Curr Opin Neurobiol 4: 294–303, 1994. 19. Baba-Aissa F, Raeymaekers L, Wuytack F, De Greef C, Missiaen L, and Casteels R. Distribution of the organellar Ca2+ transport ATPase SERCA2 isoforms in the cat brain. Brain Res 743: 141–153, 1996. 20. Liu LH, Paul RJ, Sutliff RL, Miller ML, Lorenz JN, Pun RY, Duffy JJ, Doetschman T, Kimura Y, MacLennan DH, Hoying JB, and Shull GE. Defective endothelium-dependent relaxation of vascular smooth muscle and endothelial cell Ca2+ signaling in mice lacking sarco(endo)plasmic reticulum Ca2+-ATPase isoform 3. J Biol Chem 272: 30538–30545, 1997. 21. Ouyang Y, Deerinck TJ, Walton PD, Airey JA, Sutko JL, and Ellisman MH. Distribution of ryanodine receptors in the chicken central nervous system. Brain Res 620: 269–280, 1993. 22. Sharp AH, McPherson PS, Dawson TM, Aoki C, Campbell KP, and Snyder SH. Differential immunohistochemical localization of inositol 1,4,5-trisphosphate- and ryanodine-sensitive Ca2+ release channels in rat brain. J Neurosci 13: 3051–3063, 1993. 23. Furuichi T, Furutama D, Hakamata Y, Nakai J, Takeshima H, and Mikoshiba K. Multiple types of ryanodine receptor/Ca2+ release channels are differentially expressed in rabbit brain. J Neurosci 14: 4794–4805, 1994. 24. Giannini G, Conti A, Mammarella S, Scrobogna M, and Sorrentino V. The ryanodine receptor/calcium channel genes are widely and differentially expressed in murine brain and peripheral tissues. J Cell Biol 128: 893–904, 1995. 25. Murayama T and Ogawa Y. Properties of Ryr3 ryanodine receptor isoform in mammalian brain. J Biol Chem 271: 5079–5084, 1996. 26. Ouyang Y, Martone ME, Deerinck TJ, Airey JA, Sutko JL, and Ellisman MH. Differential distribution and subcellular localization of ryanodine receptor isoforms in the chicken cerebellum during development. Brain Res 775: 52–62, 1997. 27. Llano I, Gonzalez J, Caputo C, Lai FA, Blayney LM, Tan YP, and Marty A. Presynaptic calcium stores underlie large-amplitude miniature IPSCs and spontaneous calcium transients. Nat Neurosci 3: 1256–1265, 2000.

60

A. Verkhratsky

28. Mori F, Fukaya M, Abe H, Wakabayashi K, and Watanabe M. Developmental changes in expression of the three ryanodine receptor mRNAs in the mouse brain. Neurosci Lett 285: 57–60, 2000. 29. Sharp AH, Nucifora FC Jr., Blondel O, Sheppard CA, Zhang C, Snyder SH, Russell JT, Ryugo DK, and Ross CA. Differential cellular expression of isoforms of inositol 1,4,5-triphosphate receptors in neurons and glia in brain. J Comp Neurol 406: 207–220, 1999. 30. Chen SR, Li X, Ebisawa K, and Zhang L. Functional characterization of the recombinant type 3 Ca2+ release channel (ryanodine receptor) expressed in HEK293 cells. J Biol Chem 272: 24234–24246, 1997. 31. Murayama T and Ogawa Y. Characterization of type 3 ryanodine receptor (RyR3) of sarcoplasmic reticulum from rabbit skeletal muscles. J Biol Chem 272: 24030–24037, 1997. 32. Mejia-Alvarez R, Kettlun C, Rios E, Stern M, and Fill M. Unitary Ca2+ current through cardiac ryanodine receptor channels under quasi-physiological ionic conditions. J Gen Physiol 113: 177–186, 1999. 33. Bezprozvanny I, Watras J, and Ehrlich BE. Bell-shaped calcium-response curves of Ins(1,4,5)P3- and calcium-gated channels from endoplasmic reticulum of cerebellum. Nature 351: 751–754, 1991. 34. Maeda N, Kawasaki T, Nakade S, Yokota N, Taguchi T, Kasai M, and Mikoshiba K. Structural and functional characterization of inositol 1,4,5-trisphosphate receptor channel from mouse cerebellum. J Biol Chem 266: 1109–1116, 1991. 35. Chadwick CC, Timerman AP, Saito A, Mayrleitner M, Schindler H, and Fleischer S. Structural and functional characterization of an inositol polyphosphate receptor from cerebellum. J Biol Chem 267: 3473–3481, 1992. 36. Hagar RE, Burgstahler AD, Nathanson MH, and Ehrlich BE. Type III InsP3 receptor channel stays open in the presence of increased calcium. Nature 396: 81–84, 1998. 37. Inesi G and de Meis L. Regulation of steady state filling in sarcoplasmic reticulum. Roles of back-inhibition, leakage, and slippage of the calcium pump. J Biol Chem 264: 5929–5936, 1989. 38. Favre CJ, Schrenzel J, Jacquet J, Lew DP, and Krause KH. Highly supralinear feedback inhibition of Ca2+ uptake by the Ca2+ load of intracellular stores. J Biol Chem 271: 14925–14930, 1996. 39. Mogami H, Tepikin AV, and Petersen OH. Termination of cytosolic Ca2+ signals: Ca2+ reuptake into intracellular stores is regulated by the free Ca2+ concentration in the store lumen. EMBO J 17: 435–442, 1998. 40. Solovyova N, Veselovsky N, Toescu EC, and Verkhratsky A. Ca2+ dynamics in the lumen of the endoplasmic reticulum in sensory neurons: direct visualization of Ca2+induced Ca2+ release triggered by physiological Ca2+ entry. Embo J 21: 622–630, 2002. 41. Camacho P and Lechleiter JD. Calreticulin inhibits repetitive intracellular Ca2+ waves. Cell 82: 765–771, 1995. 42. John LM, Lechleiter JD, and Camacho P. Differential modulation of SERCA2 isoforms by calreticulin. J Cell Biol 142: 963–973, 1998. 43. Li Y and Camacho P. Ca2+-dependent redox modulation of SERCA 2b by ERp57. J Cell Biol 164: 35–46, 2004. 44. Hua SY, Tokimasa T, Takasawa S, Furuya Y, Nohmi M, Okamoto H, and Kuba K. Cyclic ADP-ribose modulates Ca2+ release channels for activation by physiological Ca2+ entry in bullfrog sympathetic neurons. Neuron 12: 1073–1079, 1994. 45. Shmigol A, Eisner D, and Verkhratsky A. Cyclic ADP ribose enhances Ca2+induced Ca2+ release in isolated mouse sensory neurones. J Physiol (London) 483P: P, 1995. 46. Empson RM and Galione A. Cyclic ADP-ribose enhances coupling between voltagegated Ca2+ entry and intracellular Ca2+ release. J Biol Chem 272: 20967–20970, 1997.

ER Calcium Homeostasis & Neuronal Pathophysiology of Stroke

61

47. Budde T, Sieg F, Braunewell KH, Gundelfinger ED, and Pape HC. Ca2+-induced Ca2+ release supports the relay mode of activity in thalamocortical cells. Neuron 26: 483–492, 2000. 48. Higashida H, Yokoyama S, Hashii M, Taketo M, Higashida M, Takayasu T, Ohshima T, Takasawa S, Okamoto H, and Noda M. Muscarinic receptor-mediated dual regulation of ADP-ribosyl cyclase in NG108-15 neuronal cell membranes. J Biol Chem 272: 31272–31277, 1997. 49. Higashida H, Hashii M, Yokoyama S, Hoshi N, Asai K, and Kato T. Cyclic ADP-ribose as a potential second messenger for neuronal Ca2+ signaling. J Neurochem 76: 321–331, 2001. 50. Morikawa H, Khodakhah K, and Williams JT. Two intracellular pathways mediate metabotropic glutamate receptor-induced Ca2+ mobilization in dopamine neurons. J Neurosci 23: 149–157, 2003. 51. Galione A and Ruas M. NAADP receptors. Cell Calcium 38: 273–280, 2005. 52. Gerasimenko JV, Maruyama Y, Yano K, Dolman NJ, Tepikin AV, Petersen OH, and Gerasimenko OV. NAADP mobilizes Ca2+ from a thapsigargin-sensitive store in the nuclear envelope by activating ryanodine receptors. J Cell Biol 163: 271–282, 2003. 53. Galione A and Petersen OH. The NAADP receptor: new receptors or new regulation? Mol Interv 5: 73–79, 2005. 54. Neary JT, Crow T, and Alkon DL. Change in a specific phosphoprotein band following associative learning in Hermissenda. Nature 293: 658–660, 1981. 55. Nelson TJ, Cavallaro S, Yi CL, McPhie D, Schreurs BG, Gusev PA, Favit A, Zohar O, Kim J, Beushausen S, Ascoli G, Olds J, Neve R, and Alkon DL. Calexcitin: a signaling protein that binds calcium and GTP, inhibits potassium channels, and enhances membrane excitability. Proc Natl Acad Sci USA 93: 13808–13813, 1996. 56. Nelson TJ, Zhao WQ, Yuan S, Favit A, Pozzo-Miller L, and Alkon DL. Calexcitin interaction with neuronal ryanodine receptors. Biochem J 341: 423–433, 1999. 57. Fagni L, Chavis P, Ango F, and Bockaert J. Complex interactions between mGluRs, intracellular Ca2+ stores and ion channels in neurons. Trends Neurosci 23: 80–88, 2000. 58. Kato BM and Rubel EW. Glutamate regulates IP3-type and CICR stores in the avian cochlear nucleus. J Neurophysiol 81: 1587–1596, 1999. 59. Numakawa T, Nakayama H, Suzuki S, Kubo T, Nara F, Numakawa Y, Yokomaku D, Araki T, Ishimoto T, Ogura A, and Taguchi T. Nerve growth factor-induced glutamate release is via p75 receptor, ceramide, and Ca2+ from ryanodine receptor in developing cerebellar neurons. J Biol Chem 278: 41259–41269, 2003. 60. He X, Yang F, Xie Z, and Lu B. Intracellular Ca2+ and Ca2+/calmodulin-dependent kinase II mediate acute potentiation of neurotransmitter release by neurotrophin-3. J Cell Biol 149: 783–792, 2000. 61. Kato N, Tanaka T, Yamamoto K, and Isomura Y. Enhancement of activity-dependent calcium increase by neurotrophin-4 in visual cortex pyramidal neurons. Brain Res 832: 179–183, 1999. 62. Jimenez N and Hernandez-Cruz A. Modifications of intracellular Ca2+ signalling during nerve growth factor-induced neuronal differentiation of rat adrenal chromaffin cells. Eur J Neurosci 13: 1487–1500, 2001. 63. Mhyre TR, Maine DN, and Holliday J. Calcium-induced calcium release from intracellular stores is developmentally regulated in primary cultures of cerebellar granule neurons. J Neurobiol 42: 134–147, 2000. 64. Eu JP, Xu L, Stamler JS, and Meissner G. Regulation of ryanodine receptors by reactive nitrogen species. Biochem Pharmacol 57: 1079–1084, 1999. 65. Lu YF and Hawkins RD. Ryanodine receptors contribute to cGMP-induced late-phase LTP and CREB phosphorylation in the hippocampus. J Neurophysiol 88: 1270–1278, 2002.

62

A. Verkhratsky

66. Palmi M and Meini A. Role of the nitric oxide/cyclic GMP/Ca2+ signaling pathway in the pyrogenic effect of interleukin-1beta. Mol Neurobiol 25: 133–147, 2002. 67. Meini A, Benocci A, Frosini M, Sgaragli GP, Garcia JB, Pessina GP, Aldinucci C, and Palmi M. Potentiation of intracellular Ca2+ mobilization by hypoxia-induced NO generation in rat brain striatal slices and human astrocytoma U-373 MG cells and its involvement in tissue damage. Eur J Neurosci 17: 692–700, 2003. 68. Mikoshiba K, Furuichi T, and Miyawaki A. Structure and function of IP3 receptors. Semin Cell Biol 5: 273–281, 1994. 69. Mikoshiba K. The InsP3 receptor and intracellular Ca2+ signaling. Curr Opin Neurobiol 7: 339–345, 1997. 70. Jayaraman T, Ondrias K, Ondriasova E, and Marks AR. Regulation of the inositol 1,4,5trisphosphate receptor by tyrosine phosphorylation. Science 272: 1492–1494, 1996. 71. Cameron AM, Steiner JP, Roskams AJ, Ali SM, Ronnett GV, and Snyder SH. Calcineurin associated with the inositol 1,4,5-trisphosphate receptor-FKBP12 complex modulates Ca2+ flux. Cell 83: 463–472, 1995. 72. Genazzani AA, Carafoli E, and Guerini D. Calcineurin controls inositol 1,4,5-trisphosphate type 1 receptor expression in neurons. Proc Natl Acad Sci USA 96: 5797–5801, 1999. 73. Matifat F, Hague F, Brule G, and Collin T. Regulation of InsP3-mediated Ca2+ release by CaMKII in Xenopus oocytes. Pflugers Arch 441: 796–801, 2001. 74. Haynes LP, Tepikin AV, and Burgoyne RD. Calcium-binding protein 1 is an inhibitor of agonist-evoked, inositol 1,4,5-trisphosphate-mediated calcium signaling. J Biol Chem 279: 547–555, 2004. 75. Yang J, McBride S, Mak DO, Vardi N, Palczewski K, Haeseleer F, and Foskett JK. Identification of a family of calcium sensors as protein ligands of inositol trisphosphate receptor Ca2+ release channels. Proc Natl Acad Sci USA 99: 7711–7716, 2002. 76. Cameron AM, Steiner JP, Sabatini DM, Kaplin AI, Walensky LD, and Snyder SH. Immunophilin FK506 binding protein associated with inositol 1,4,5-trisphosphate receptor modulates calcium flux. Proc Natl Acad Sci USA 92: 1784–1788, 1995. 77. Dargan SL, Lea EJ, and Dawson AP. Modulation of type-1 Ins(1,4,5)P3 receptor channels by the FK506-binding protein, FKBP12. Biochem J 361: 401–407, 2002. 78. Friel DD and Tsien RW. A caffeine- and ryanodine-sensitive Ca2+ store in bullfrog sympathetic neurones modulates effects of Ca2+ entry on [Ca2+]i. J Physiol 450: 217–246, 1992. 79. Verkhratsky A and Shmigol A. Calcium-induced calcium release in neurones. Cell Calcium 19: 1–14, 1996. 80. Verkhratsky A. The endoplasmic reticulum and neuronal calcium signalling. Cell Calcium 32: 393–404, 2002. 81. Berridge MJ, Bootman MD, and Roderick HL. Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol 4: 517–529, 2003. 82. Koizumi S, Bootman MD, Bobanovic LK, Schell MJ, Berridge MJ, and Lipp P. Characterization of elementary Ca2+ release signals in NGF-differentiated PC12 cells and hippocampal neurons. Neuron 22: 125–137, 1999. 83. Melamed-Book N, Kachalsky SG, Kaiserman I, and Rahamimoff R. Neuronal calcium sparks and intracellular calcium ‘‘noise’’. Proc Natl Acad Sci USA 96: 15217–15221, 1999. 84. Usachev YM and Thayer SA. All-or-none Ca2+ release from intracellular stores triggered by Ca2+ influx through voltage-gated Ca2+ channels in rat sensory neurons. J Neurosci 17: 7404–7414, 1997. 85. Berridge MJ and Irvine RF. Inositol trisphosphate, a novel second messenger in cellular signal transduction. Nature 312: 315–321, 1984. 86. Finch EA and Augustine GJ. Local calcium signalling by inositol-1,4,5-trisphosphate in Purkinje cell dendrites. Nature 396: 753–756, 1998.

ER Calcium Homeostasis & Neuronal Pathophysiology of Stroke

63

87. Takechi H, Eilers J, and Konnerth A. A new class of synaptic response involving calcium release in dendritic spines. Nature 396: 757–760, 1998. 88. Nakamura T, Nakamura K, Lasser-Ross N, Barbara JG, Sandler VM, and Ross WN. Inositol 1,4,5-trisphosphate IP3-mediated Ca2+ release evoked by metabotropic agonists and backpropagating action potentials in hippocampal CA1 pyramidal neurons. J Neurosci 20: 8365–8376, 2000. 89. Nakamura T, Barbara JG, Nakamura K, and Ross WN. Synergistic release of Ca2+ from IP3-sensitive stores evoked by synaptic activation of mGluRs paired with backpropagating action potentials. Neuron 24: 727–737, 1999. 90. Nicotera P, Petersen OH, Melino G, and Verkhratsky A. Janus a god with two faces: death and survival utilise same mechanisms conserved by evolution. Cell Death Differ 14: 1235–1236, 2007. 91. Seta KA, Yuan Y, Spicer Z, Lu G, Bedard J, Ferguson TK, Pathrose P, Cole-Strauss A, Kaufhold A, and Millhorn DE. The role of calcium in hypoxia-induced signal transduction and gene expression. Cell Calcium 36: 331–340, 2004. 92. Toescu EC. Hypoxia sensing and pathways of cytosolic Ca2+ increases. Cell Calcium 36: 187–199, 2004. 93. Silver IA and Erecinska M. Intracellular and extracellular changes of [Ca2+] in hypoxia and ischemia in rat brain in vivo. J Gen Physiol 95: 837–866, 1990. 94. Grondahl TO, Hablitz JJ, and Langmoen IA. Depletion of intracellular Ca2+ stores or lowering extracellular calcium alters intracellular Ca2+ changes during cerebral energy deprivation. Brain Res 796: 125–131, 1998. 95. Dubinsky JM and Rothman SM. Intracellular calcium concentrations during ‘‘chemical hypoxia’’ and excitotoxic neuronal injury. J Neurosci 11: 2545–2551, 1991. 96. Nakayama R, Yano T, Ushijima K, Abe E, and Terasaki H. Effects of dantrolene on extracellular glutamate concentration and neuronal death in the rat hippocampal CA1 region subjected to transient ischemia. Anesthesiology 96: 705–710, 2002. 97. Ouardouz M, Nikolaeva MA, Coderre E, Zamponi GW, McRory JE, Trapp BD, Yin X, Wang W, Woulfe J, and Stys PK. Depolarization-induced Ca2+ release in ischemic spinal cord white matter involves L-type Ca2+ channel activation of ryanodine receptors. Neuron 40: 53–63, 2003. 98. Pahl HL and Baeuerle PA. The ER-overload response: activation of NF-kappa B. Trends Biochem Sci 22: 63–67, 1997. 99. Patil C and Walter P. Intracellular signaling from the endoplasmic reticulum to the nucleus: the unfolded protein response in yeast and mammals. Curr Opin Cell Biol 13: 349–355, 2001. 100. Lin JH, Walter P, and Yen TS. Endoplasmic reticulum stress in disease pathogenesis. Annu Rev Pathol 3: 399–425, 2008. 101. Tamatani M, Matsuyama T, Yamaguchi A, Mitsuda N, Tsukamoto Y, Taniguchi M, Che YH, Ozawa K, Hori O, Nishimura H, Yamashita A, Okabe M, Yanagi H, Stern DM, Ogawa S, and Tohyama M. ORP150 protects against hypoxia/ischemia-induced neuronal death. Nat Med 7: 317–323, 2001. 102. Thastrup O, Dawson AP, Scharff O, Foder B, Cullen PJ, Drobak BK, Bjerrum PJ, Christensen SB, and Hanley MR. Thapsigargin, a novel molecular probe for studying intracellular calcium release and storage. 1989. Agents Actions 43: 187–193, 1994. 103. Nagasaki K and Fleischer S. Ryanodine sensitivity of the calcium release channel of sarcoplasmic reticulum. Cell Calcium 9: 1–7, 1988. 104. Buck E, Zimanyi I, Abramson JJ, and Pessah IN. Ryanodine stabilizes multiple conformational states of the skeletal muscle calcium release channel. J Biol Chem 267: 23560–23567, 1992. 105. Zucchi R and Ronca-Testoni S. The sarcoplasmic reticulum Ca2+ channel/ryanodine receptor: modulation by endogenous effectors, drugs and disease states. Pharmacol Rev 49: 1–51, 1997.

64

A. Verkhratsky

106. Ohta T, Ito S, and Ohga A. Inhibitory action of dantrolene on Ca-induced Ca2+ release from sarcoplasmic reticulum in guinea pig skeletal muscle. Eur J Pharmacol 178: 11–19, 1990. 107. Ritov VB, Men’shikova EV, and Kozlov YP. Heparin induces Ca2+ release from the terminal cisterns of skeletal muscle sarcoplasmic reticulum. FEBS Lett 188: 77–80, 1985. 108. Bezprozvanny IB, Ondrias K, Kaftan E, Stoyanovsky DA, and Ehrlich BE. Activation of the calcium release channel (ryanodine receptor) by heparin and other polyanions is calcium dependent. Mol Biol Cell 4: 347–352, 1993. 109. Ehrlich BE, Kaftan E, Bezprozvannaya S, and Bezprozvanny I. The pharmacology of intracellular Ca2+-release channels. Trends Pharmacol Sci 15: 145–149, 1994. 110. Wakui M, Osipchuk YV, and Petersen OH. Receptor-activated cytoplasmic Ca2+ spiking mediated by inositol trisphosphate is due to Ca2+-induced Ca2+ release. Cell 63: 1025–1032, 1990. 111. Hirose K, Iino M, and Endo M. Caffeine inhibits Ca2+-mediated potentiation of inositol 1,4,5-trisphosphate-induced Ca2+ release in permeabilized vascular smooth muscle cells. Biochem Biophys Res Commun 194: 726–732, 1993. 112. Maruyama T, Kanaji T, Nakade S, Kanno T, and Mikoshiba K. 2APB, 2-aminoethoxydiphenyl borate, a membrane-penetrable modulator of Ins(1,4,5)P3-induced Ca2+ release. J Biochem (Tokyo) 122: 498–505, 1997. 113. Peppiatt CM, Collins TJ, Mackenzie L, Conway SJ, Holmes AB, Bootman MD, Berridge MJ, Seo JT, and Roderick HL. 2-aminoethoxydiphenyl borate (2-APB) antagonises inositol 1,4,5-trisphosphate-induced calcium release, inhibits calcium pumps and has a use-dependent and slowly reversible action on store-operated calcium entry channels. Cell Calcium 34: 97–108, 2003. 114. Bootman MD, Collins TJ, Mackenzie L, Roderick HL, Berridge MJ, and Peppiatt CM. 2-aminoethoxydiphenyl borate (2-APB) is a reliable blocker of store-operated Ca2+ entry but an inconsistent inhibitor of InsP3-induced Ca2+ release. FASEB J 16: 1145–1150, 2002. 115. Gafni J, Munsch JA, Lam TH, Catlin MC, Costa LG, Molinski TF, and Pessah IN. Xestospongins: potent membrane permeable blockers of the inositol 1,4,5-trisphosphate receptor. Neuron 19: 723–733, 1997. 116. Solovyova N, Fernyhough P, Glazner G, and Verkhratsky A. Xestospongin C empties the ER calcium store but does not inhibit InsP3-induced Ca2+ release in cultured dorsal root ganglia neurones. Cell Calcium 32: 49–52, 2002. 117. Takahashi M, Tanzawa K, and Takahashi S. Adenophostins, newly discovered metabolites of Penicillium brevicompactum, act as potent agonists of the inositol 1,4,5trisphosphate receptor. J Biol Chem 269: 369–372, 1994. 118. Wilcox RA, Primrose WU, Nahorski SR, and Challiss RA. New developments in the molecular pharmacology of the myo-inositol 1,4,5-trisphosphate receptor. Trends Pharmacol Sci 19: 467–475, 1998.

The Naþ/Ca2þ Exchanger: A Target for Therapeutic Intervention in Cerebral Ischemia Lucio Annunziato, Pasquale Molinaro, Agnese Secondo, Anna Pannaccione, Antonella Scorziello, Giuseppe Pignataro, Ornella Cuomo, Rossana Sirabella, Francesca Boscia, Alessandra Spinali, and Gianfranco Di Renzo

1 Introduction The Naþ/Ca2þ exchanger (NCX) belongs to the superfamily of Ca2þ/cation antiporter (CaCA) membrane proteins comprising the following members: (1) the NCX family, which exchanges three Naþ ions for one Ca2þ ion (Reeves and Hale, 1984; Fujioka et al., 2000; Kang and Hilgemann, 2004); (2) the Naþ/Ca2þ exchanger Kþ-dependent family (NCKX), which exchanges four Naþ for one Ca2þ plus one Kþ ion [1, 2]; (3) the bacterial family which probably promotes Ca2þ/Hþ exchange (YRBG) [3]; (4) the nonbacterial Ca2þ/Hþ exchange family (CAX), which is also the Ca2þ exchanger of yeast vacuoles [4]; and (5) the Ca2þ/ cation exchanger family (CCX), which contains the partially characterized molecule previously called Naþ/Ca2þ–Liþ exchanger (NCLX or NCKX6). Among these plasma membrane antiporters, the Kþ-independent NCX is a high-capacity and low-affinity ionic transporter that exchanges three Naþ ions for one Ca2þ ion. When intracellular Ca2þ concentrations ([Ca2þ]i) rise and the cells require the return to resting levels, this exchanger transport mechanism couples the uphill extrusion of Ca2þ ions to the entry of Naþ ions into the cells down their electrochemical gradient. This mode of operation, defined as forward mode or Ca2þ efflux or, more correctly, Ca2þ-exit mechanism, keeps the 104-fold difference in [Ca2þ]i across the cell membrane. In other physiological or pathophysiological conditions when the intracellular Naþ concentrations ([Naþ]i) rise or membrane depolarization occurs, thus reducing the transmembrane Naþ electrochemical gradient, NCX reverses its mode of operation and mediates the extrusion of Naþ and the entry of Ca2þ ions. This mode of operation is defined as reverse mode or Ca2þ-entry mechanism. The mode of operation of the antiporter depends on (1) the Naþ gradient, (2) the Ca2þ gradient, and (3) the membrane potential. Therefore NCX is a major player L. Annunziato (*) Division Pharmacology, Department of Neuroscience, School of Medicine, ‘‘Federico II’’ University of Naples, 80131 Naples, Italy e-mail: [email protected]

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in the regulation of physiological and pathophysiological responses to increases of [Ca2þ]i and [Naþ]i [5, 6, 7].

2 Gene Structure 2.1 NCX Family Three genes coding for the three different NCX1 [8], NCX2 [9], and NCX3 [10] proteins have been identified in mammals. These three genes appear to be dispersed, since NCX1, NCX2, and NCX3 have been mapped in human chromosomes 2, 19, and 14, respectively. The NCX1 gene includes 12 exons spread over a region of about 200 kb; however, the major part of the mature protein is coded by exons 2, 11, and 12. The different brain, kidney, and heart NCX1 promoters are partly located in exon 1, in a region comprised from 80 kb to 20 kb before the coding exon 2. Transcripts deriving from exons 3–8, labeled A through F, are subjected to alternative splicing generating different tissuespecific mature RNAs. In order to maintain an open reading frame, all splice variants must include either transcript of exon 3, named also A, or of exon 4, named also B, which are mutually exclusive in NCX1 mature RNA. At least 15 splice variants have been detected for NCX1, named NCX1.1–NCX1.15. Excitable tissues usually have splicing variants encoded by mRNAs, which include transcript of exon A whereas transcript of exon B predominates in other tissues. The NCX2 gene includes 10 exons spreading on a region of about 150 kb. To date, the promoter sequence and location are unknown and no alternative splicing has been demonstrated for this gene. The NCX3 gene includes nine exons spanning a region of about 150 kb. NCX3 minimal promoter spans for 300 bp, just upstream to exon 1. The exons 2–5 present analogies with exons 2–5 of the NCX1 gene [11]. Transcript regions derived from exons 3, 4, and 5 are subjected to alternative splicing. Thus, NCX3 gene has only three exons, instead of six in NCX1, that participate to alternative splicing of the mature transcript. Exons 6–9 of NCX3 correspond to homologous NCX1 exons 9–12. Six splice variants have been described for NCX3: NCX3.1, NCX3.2, NCX3.3, NCX3.4, NCX3-tN.1, and NCX3-tN.2. The physiological and functional significance of all these alternative splicing forms remains unclear; however, it is possible that tissue-specific pattern of alternative splicing is tailored to meet the specific needs of particular brain regions or neuronal population.

2.2 Gene Transcription Regulation The transcription of NCX1 gene is independent of [Ca2þ]i levels. By contrast, the regulation of NCX2 and NCX3 genes is strictly dependent on this bivalent

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cation, although in opposite directions [12, 13, 14]. Thus, changes in [Ca2þ]i levels in neurons directly regulate the amount of NCX2 and NCX3 produced proteins. The physiological and pathophysiological relevance of this elaborated cellular mechanism to modulate NCX protein diversity is nevertheless not fully understood.

2.3 NCX1 Transcription NCX1 gene has three tissue-specific transcription start sites regulated by heart, kidney, and brain promoters [13, 14]. NCX1 brain promoter is not provided with TATA box sequence, but contains several consensus sequences for specific protein 1 (Sp-1) and three binding sites for activator protein 2 (AP-2). The presence of these three AP-2-binding sites implies that NCX1 brain promoter has a housekeeping role and could also exhibit cell-specific properties similar to those observed for the nerve growth factor gene [15]. In addition, NCX1 brain promoter also contains putative binding sites for ubiquitous transcription factors such as nuclear factor-kB (NFkB) and hypoxia-inducible factor 1 (HIF-1).

2.4 NCX2 Transcription To date, no extensive data are available for NCX2 gene promoter and its transcription regulation. However, it has been demonstrated that NCX2 expression is rapidly downregulated after membrane depolarization in a Ca2þ-dependent phosphatase calcineurin manner [16]. Interestingly, the expression of NCX2 after transient middle cerebral artery occlusion results in a massive downregulation throughout the brain, both at the level of peri-infarct area as well as in non-ischemic surrounding brain regions [17]. These areas are subjected to repetitive spreading depression-like depolarization, originating in the ischemic core and propagating to the surrounding brain regions of the same hemisphere [18]. This depression-like depolarization might be responsible for the observed changes in NCX2 gene expression during ischemia [17].

2.5 NCX3 Transcription The NCX3 minimal promoter contains specific enhancers for both muscle and neuronal expression such as cAMP response element (CRE) sequence [19]. However, the complex constituted by CRE-binding protein (CREB) and CREB-binding protein (CBP), by itself, is not sufficient to initiate the NCX3 transcription activation in vivo, suggesting that other unidentified enhancers might be involved [20]. Interestingly, following permanent occlusion of middle

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cerebral artery in rats, NCX3 transcript increases in the peri-infarct region [17], a territory where pro-survival genes, driven by CRE enhancer, are upregulated [21]. In addition, in primary neuronal cultures, unlike NCX2, depolarization upregulates NCX3 mRNA in a calcineurin-independent manner [16]. NCX3 gene promoter also presents a repressor sequence constituted by a doublet of downstream regulatory element (DRE). This doublet of DRE sequences is bound by a transcriptional factor called Ca2þ-modulated transcriptional repressor downstream regulatory element antagonist modulator (DREAM), whose binding to DNA is inhibited by elevated [Ca2þ]i. When [Ca2þ]i is low, the affinity of DREAM for the DRE sequences increases and consequently NCX3 transcription is inhibited whereas when [Ca2þ]i is high, NCX3 transcription is increased [22]. Thus, the transcriptional regulation of NCX3 is dependent on Ca2þ level. This is a striking example of a feedback regulation of the Ca2þ signal in neurons.

3 Structural Features NCX has nine transmembrane segments [23] that can be divided into an Nterminal hydrophobic domain, composed of the first five TMS (1–5), and into a C-terminal hydrophobic domain, composed of the last four TMS (6–9) (Fig. 1). Amino acid sequence found between TMS2 and TMS3 is called a1 repeat whereas the region between TMS7 and TMS8 is named a2 repeat. Both a1

Fig. 1 Molecular topology of NCX with putative sites were drugs interfere with NCX activity.

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and a2 repeats are located on the extracellular and intracellular space, respectively, and may form a portion of the ion translocation pathway [24, 25]. The N-terminal hydrophobic domain (TMS1–5) is separated from the C-terminal hydrophobic domain (TMS6–9) through a large hydrophilic intracellular loop of 550 amino acids, named the f loop [23]. Although the f loop is not implicated in Naþ and Ca2þ translocation, it is responsible for the regulation of NCX activity elicited by several cytoplasmic messengers and transductional mechanisms, such as Hþ, Ca2þ, and Naþ ions, nitric oxide (NO), phosphatidylinositol 4,5 bisphosphate (PIP2), protein kinase C (PKC), protein kinase A (PKA), phosphoarginine (PA), and ATP. Furthermore, in the f loop, there are two stimulatory Ca2þ-binding domains named CBD1 and CBD2. Each CBD has an immunoglobulin-like fold that binds two Ca2þ ions [23, 26]. CBD1 displays a high affinity for Ca2þ with a Kd of 120–400 nM and is the primary Ca2þ sensor since it is able to detect small increases in cytosolic Ca2þ, thus inducing a large structural changes that activate the exchanger. In contrast, CBD2 displays a low affinity for Ca2þ with a Kd ranging from 820 nM to 8.6 mM and it binds Ca2þ only when [Ca2þ]i is elevated with a consequent modest structural alterations. These two different sensitivity thresholds may enable NCX to function dynamically over a wide range of Ca2þ concentrations and permit high Ca2þ exit or entry in excitable cells. In the C-terminal portion of the f loop, at the level of 562–679 aa, there is a putative sequence responsible for Naþ-dependent inactivation (I1) of NCX. In particular, when [Naþ]i rises above 15 mM this region might bind a 20-amino acid sequence, 219–238, named exchanger inhibitory peptide (XIP), located in the N-terminal portion of the same f loop, causing NCX inhibition [27]. In fact, the mutation of some amino acids of XIP sequence increases or removes NCX Naþ-dependent inactivation [28]. Interestingly, from the pharmacological point of view, an exogenous peptide, having the same amino acid sequence of XIP region in the f loop, exerts a potent inhibitory activity on NCX function [29, 30]. NCX2 and NCX3 proteins consist of 921 and 927 amino acids, respectively, and are characterized by molecular masses of 102 and 105 kDa, respectively. In addition, NCX2 displays a 65% sequence identity with NCX1 whereas NCX3 possesses a 73% sequence identity with NCX1 and 75% sequence identity with NCX2 [24]. All three NCX gene products share the same membrane topology with the amino-terminus located in the extracellular space, and the carboxylterminus is located intracellularly.

4 Molecular Biology NCX1 has an extracellular disulfide bond connecting Cys14 or Cys20, located in the extracellular N-terminal tail with Cys792, present in the loop between TMS6 and TMS7 [31]. Moreover, a1 and a2 repeats are

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speculatively modeled to be a re-entrant membrane loop similar to the poreforming region of ion channels. This arrangement is similar to that of the aquaporin water channels, which have also been found to have intramolecular repeats on opposite membrane surfaces [32]. The center of the hydrophobic segment of a2 repeat has a GIG sequence similar to the GYG motif characteristic of Kþ channel P loops. The expression of a mutated exchanger protein which lacks the C-terminal domain or the N-terminal domain does not show any antiporter activity, suggesting that both domains are required to exert exchanger activity [33].

5 Tissue and Cellular Distribution The NCX1 gene displays an ubiquitous expression and therefore is present in several tissues, including brain, heart, skeletal muscle, smooth muscle, kidney, eye, secretory, and blood cells whereas NCX2 and NCX3 gene products have been found exclusively in neuronal and skeletal muscle tissues [12]. The distribution of the three NCX isoforms in the mammalian brain gave useful insights into the physiological and pathophysiological role played by the different NCX isoforms in the regulation of neuronal function. In fact, NCX1, NCX2, and NCX3 are highly expressed in the mammalian brain, suggesting that NCX proteins may play a role in the regulation of neuronal function. Moreover, NCX1 and NCX3 have several splicing variants that appear to be selectively expressed in different regions and cellular populations of the brain [34, 35]. In cerebral cortex, NCX1 is intensively expressed in the pyramidal neurons of layers III and V within the molecular layer of the cerebral motor cortex. This area, which contains the terminal dendritic field of the pyramidal cells, displays a more intense NCX1 immunoreactivity than that of NCX2. In contrast, the somatosensory cortical area seems to express preferentially NCX2 transcripts. Such anatomical distribution reveals that the upper neurons of the motor system and the terminal neurons of the somatosensory system preferentially express distinct NCX isoforms [36, 37]. Within the hippocampus, the transcripts of the three NCX isoforms display an intense labeling of most neuronal populations. In particular, high levels of the three NCX genes have been detected in the granular cell layers of the dentate gyrus and in the pyramidal cells of CA1, CA2, CA3, and CA4 subfields [37]. The three NCX protein isoforms also display high levels of expression within the hippocampus. Thus, in the orient and radiatum layers of the CA1, NCX3 protein is more intense than NCX1 and NCX2. NCX1 protein expression is particularly intense in the granule cell layer and in the hilum of the dentate gyrus, which constitutes the terminal field of the perforant pathway, the major excitatory input to the hippocampus originating from the enthorinal cortex. In the CA3 area, the NCX1 and NCX3 genes of the mossy fibers projecting from the granule cells located in

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the dentate gyrus are more intense than those of NCX2. This peculiar distribution suggests that distinct NCX isoforms may play a crucial role in controlling the intracellular Naþ and Ca2þ homeostasis of the major afferent, intrinsic, and efferent hippocampal projections. Such circuitries are crucial for the synaptic plasticity phenomena, such as those involved in long-term potentiation (LTP) and long-term depression (Madison et al., 1991). NCX isoforms are also expressed in crucial areas of the extrapyramidal control of motor coordination. In fact, NCX1 mRNA can be detected in the substantia nigra pars compacta, in which dopaminergic cell bodies are localized; the NCX1 protein isoform is present in the striatum, in which the terminal projection fields of dopaminergic nigrostriatal neurons are found. Interestingly, both the transcript and the protein, encoded by the three NCX genes, are abundantly expressed in the nucleus accumbens [36, 37], a brain region involved in the motivational control of motor coordination and damaged following middle cerebral artery occlusion.

6 Biophysical and Electrophysiological Properties Naþ/Ca2þ exchanger has a stoichiometry of the three Naþ for one Ca2þ [38]; recent evidence demonstrated that this ion flux ratio is 3.2 for maximal transport in either direction, and at low rate NCX1 can also transport one Ca2þ with one Naþ ion [39]. The Naþ/Ca2þ exchanger has about a 10-fold lower affinity for Ca2þ than ATP-driven Ca2þ pumps but, when it is fully activated, can have a very high rate of transport with an estimate turnover value comprised between 1000 and 5000 s–1 for the transport of both Naþ and Ca2þ using non-saturating ion concentrations. This value is 10- to 50-fold higher than that showed by ATPdriven Ca2þ pumps [40]. A density of 400 exchanger molecules per square mm of plasma membrane produces a current of 30 pA/F [41]. The direction and the amplitude of the Naþ/ Ca2þ exchanger current depend on (1) cellular membrane potential (VM) and (2) Naþ and Ca2þ gradients. These gradients set the reversal potential of NCX (ENCX) which represents the Vm at which the exchanger reverses its mode of operation. The direction of the NCX current (INCX) depends on the difference between ENCX and the cellular membrane potential. When VM is more negative than ENCX, NCX works in Ca2þ-exit mode, in contrast, when the VM value is more positive than reversal potential ENCX, NCX operates in Ca2þ-entry mode. Since in resting neurons VM is –60 mV and ENCX is –40 mV, NCX operates in Ca2þ-exit mode at a low rate. The amplitude of INCX depends on the difference between VM and ENCX values and on several regulatory factors, such as the presence of inhibitors, phosphorylation, inactivation, and interaction with other proteins.

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7 Regulation of Naþ/Ca2+ Exchanger Activity Several factors are involved in the regulation of Naþ/ Ca2þ exchanger activity: (i) the concentrations of the two transported cations, Naþ and Ca2þ; (ii) the intracellular pH; (iii) the metabolic-related compounds, ATP, PIP2, PKA, and PKC; and (iv) reactive oxygen species (ROS) and reactive nitrogen species (RNS). [Ca2þ]i regulates NCX activity through CBD (see page 69). An increase in [Naþ]i can also regulate the Naþ/Ca2þ exchanger. In particular, when [Naþ]i increases, it binds to the transport site of the exchanger molecule, and after this Naþ influx, an inactivation process of the exchanger occurs. This inactivation process, very similar to the phenomenon occurring in voltage-dependent ionic channels, is named Naþ-dependent inactivation. The intracellular pH can also regulate the exchanger. [Hþ] strongly inhibits NCX activity under steady-state conditions, in fact, reductions in pHi values, as little as 0.4, can induce a 90% inhibition of NCX activity. Such inhibitory action depends on the presence of intracellular Naþ ions; hence, the action exerted by Hþ ions is pathophysiologically relevant with regards to brain and heart ischemia. ATP, acting as a phosphoryl donor molecule, may increase the activity of the exchanger in a number of ways. First, by activating G-protein-coupled receptors, via endogenous and exogenous ligands. Second, ATP can stimulate the activity of the Naþ/Ca2þ exchanger through the pathway involving PKC or PKA activation, and each of the NCX isoforms has distinctive putative phosphorylation sites. Finally, the other mechanism by which ATP can activate NCX occurs through the production of the lipid PIP2. In fact, this lipid binding the XIP region of the f loop eliminates NCX inactivation and stimulates NCX function. Interestingly, ATP cellular depletion interferes differently on the three isoforms, as it inhibits NCX1 and NCX2 but does not affect NCX3 activity [42, 43]. Among the regulatory factors, Naþ/Ca2þ exchanger is sensitive to reactive oxygen species, since modifications of the cellular redox state can cause an increase of NCX activity [5].

8 Physiological Properties The Naþ/ Ca2þ exchanger protein is present in both neurons and glia, and may play a relevant function in different neurophysiological conditions. In neurons, the level of expression of NCX is particularly high in those sites where a large movement of Ca2þ ions occurs across the plasma membrane, as it happens at the level of synapses [36, 44]. Specifically, during an action potential or after glutamate-activated channel activity, Ca2þ massively enters the plasma membrane. Such phenomenon triggers the fusion of synaptic vesicles with the

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plasma membrane and promotes neurotransmitter exocytosis. After this event, outward Kþ currents repolarize the plasma membrane, thus leading to VGCC closure. According to the diffusion principle, Ca2þ ions are distributed in the cytosolic compartment, reversibly interacting with Ca2þ-binding proteins. Residual Ca2þ is then rapidly extruded by the plasma membrane Ca2þ ATPase and by NCX. The Naþ/Ca2þ exchanger becomes the dominant Ca2þ extrusion mechanism when [Ca2þ]i is higher than 500 nM, as it happens when a train of action potentials reaches the nerve terminals. In glial cells the Naþ/Ca2þ exchanger is also significantly expressed [45, 46, 47, 48], but the precise role of the exchanger has not yet been determined. To characterize the precise role of the different NCX genes, knockout mice have been generated lacking NCX1 or NCX2 or NCX3. Targeted deletion of NCX1 results in NCX1-null embryos that die before birth [49]. Mice lacking NCX2 gene, the major isoform expressed in the brain, exhibit an enhanced performance in several hippocampus-dependent learning and memory tasks, together with a significantly delayed clearance of elevated Ca2þ following depolarization [50]. In addition, the frequency threshold for LTP and longterm depression in the hippocampal CA regions was shifted to a lowered frequency favoring LTP in these knockout mice [50]. Interestingly, mice lacking ncx3 gene exhibit reduced motor activity, weakness of forelimb muscles, and fatigability in comparison with ncx3+/+ mice [51]. However, since NCX3 is expressed in the CNS and in peripheral nervous system, it cannot be established whether these symptoms can be attributed to CNS defects or to alterations at the neuronal muscular junctions and skeletal fibers levels.

9 Pharmacological Modulation More than 40 years ago, amiloride and amiloride analogs were described by Cragoe [52], by a biological screen process, as potent inhibitors of sodium channels in urinary epithelium thus acting as a potassium-sparing diuretics. Amiloride and its analogs were subsequently shown to be inhibitors of other membrane transporters. In the same years of amiloride synthesis, Baker and Blaustein [53] functionally discovered the existence of the ubiquitous plasma membrane NCX. Amiloride, at very high concentrations, was found to be an effective inhibitor of NCX. Since then, amiloride has been used by numerous investigators, as a probe to block NCX function [54]. Unfortunately, two major drawbacks have limited its use. First, millimolar concentrations were required for its NCX inhibitory activity; second, it lacked of specificity, for it can also inhibit both the epithelial Naþ channels (ENaC) at micromolar concentrations [55] and the Naþ/Hþ exchanger (NHE) in the millimolar range. In the effort to develop amiloride derivatives provided with greater selectivity, Cragoe synthesized compounds more specific for NCX and NHE, respectively. In particular, two classes of amiloride analogs have been developed. The amiloride analogs of

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the first class, such as 5-[N-methyl-N-(guanidinocarbonylmethyl)] amiloride, bear substituents on the 5-amino nitrogen atom of the pyrazine ring [56]. They lack inhibitory properties on ENaC and NCX, even though they display greater effectiveness in inhibiting NHE in the 1–10 mM range [57, 58, 59]. The compounds of the second class, having no inhibitory effect on the Naþ/Hþ exchanger, bear substituents on the terminal guanidino nitrogen atom and behave as specific inhibitors (Ki 1–10 mM) of ENaC and NCX. Among these compounds, CB–DMB appears to be the most specific inhibitor of NCX activity, for it has no inhibitory properties against NHE and the ENaC in excitable cells [54], such as neurons, in which the kidney ENaC are not expressed [56, 57] (Table 1). Table 1 Inhibitors and activators of NCX activity Pharmacological class Compond NCX Inhibitors 1. Amiloride Derivatives 2. Substituted Pyrrolidines 3. Isothiourea Derivatives 4. Ethoxyanilines 5. Benzofuran Derivatives 6. Quinazolinone Derivatives 7. Thiazolidine Derivatives 8. Phenoxypyridine Derivatives 9. Nicotinamide Derivatives 10. Piperidine Derivatives 11. Ylacetamide Derivatives 12. Benzyloxyphenyl Derivatives 13. Peptides 14. Small Interference RNA 15. Antisense Oligodeoxynucleotides 16. Inorganic Cations NCX Activators 1. Redox Agents 2. Organic Compounds 3. Inorganic Cations 4. Agonists of GPCRs 5. Peptides

CB-DMB, DCB, DMB Bepridil KB-R7943 SEA0400 Amiodarone SM-15811 SN-6 JP11092454 6-[4-[(3-fluorobenzyl)-oxy]phenoxy]nicotinamide YM-252077 YM-270951 YM-244769 XIP, Glu-XIP, FMRFa, FRCRCF siRNA-NCX1, siRNA-NCX1, siRNA-NCX3 AS-NCX1, AS-NCX2, AS-NCX3 Ni2þ, La2þ, Gd2þ DTT, GSH, Fe2þ, O2 and of Fe3þ, H2O2 Diethylpyrocarbonate (DEPC) Liþ a, b Agonists, Histamine, 5HT2c Agonists, Endothelin-1, Angiotensin-II Insulin, Concanavalin A

More recently, Shigekawa’s group screening a compound library for the inhibitions of Naþ-dependent Ca2þ uptake identified an isothiourea derivative named KB-R7943 which gained the reputation as a specific compound for inhibiting the antiporter and therefore as a probe to test NCX functional role [60]. KB-R7943 in the low micromolar range, is able to block the reverse mode

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operation of the antiporter whereas concentrations 30 times higher are needed to inhibit the forward mode [61, 62]. However, recent reports have obscured its reputation as a selective probe for NCX inhibition. Indeed, several studies showed that KB-R7943 also exerts an inhibitory effect on several other pharmacological targets such as NHE, dihydropyridine (DHP)-sensitive Ca2þ uptake, passive Naþ uptake, Ca2þ-ATPases and Naþ, Kþ-ATPase [59], and receptoroperated NMDA channels [62]. In addition, it has been reported that the NCX inhibitor KB-R7943 activates large-conductance Ca2þ-activated Kþ channels in endothelial and vascular smooth muscle cells [63]. On the other hand, in the last decade several studies reported CB–DMB as an effective probe to study the role of NCX in some pathophysiological mechanisms including platelet hyperactivity in diabetes [64], collagen-induced platelet activation [64], and in vitro and in vivo ischemic cerebral death mechanisms [30, 65, 66, 67, 68]. In 2001, a new compound belonging to the ethoxyanilines family, SEA0400, was reported as being the most potent NCX inhibitor available at the time, IC50 = 5–92 nM, with a predominant activity on NCX1, a lower affinity for NCX2 and no effect on NCX3. However, the specificity of SEA0400 on NCX activity has recently been questioned, since it can also interfere with Ca2þ movement across the cell membrane. In 2002, by screening benzyloxyphenyl derivatives, Iwamoto’s group discovered the new compound SN-6, which differs from KB-R7943 only in the substituent of phenyl moiety. The presence of this phenyl group confers to this antiporter inhibitor a more selective action on NCX1 rather than NCX3. Using chimeric analysis and subsequent site-directed mutagenesis some critical amino acid residues responsible of SN-6 inhibition in the XIP region of the antiporter have been identified. Interestingly, SN-6 preferentially acts on the exchanger under ATP-depleted conditions. Recently, the further screening of new benzyloxyphenyl derivatives revealed a highly potent NCX inhibitor, named YM-244769. This orally bioavailable compound is more potent in inhibiting NCX3 than NCX1 and NCX2 in the reverse mode, but it is not active on the forward mode of operation of the three antiporter isoforms [69]. Recently, two more selective strategies able to inhibit each of the three NCX gene products, has been introduced as pharmacological tool. These strategies consist in the antisense oligodeoxynucleotide technique and small interference RNA directed against mRNA sequence of NCX1, NCX2, and NCX3 [70, 71]. With this potentially therapeutic approach it has been possible to discriminate the role played by each of the three NCX gene products in preclinical models of pathological states in which this antiporter is involved. The availability of pharmacological agents capable of stimulating the activity of NCX, either in the reverse or in the forward mode of operation, may represent a useful strategy to adopt in some pathophysiological conditions, such as brain ischemia. The pharmacological stimulation of the antiporter could, in fact, contribute to the re-establishment of intracellular Naþ and Ca2þ ion homeostasis. Among NCX activators there are monovalent cations such as Litium that stimulates the Naþi-dependent Ca2þ uptake of all three NCX gene products with low affinity. Reducing agents such as GSH, DDT,

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Fe2þ, and O2– superoxide in presence of oxidating agents such as Fe3þ, H2O2, GSSG, and O2 are able to stimulate NCX activity [72]. Organic compounds, such as agonists of G-protein-coupled receptors, diethylpyrocarbonate, concavallin A, NGF, and insulin, are capable to enhance NCX activity at protein level by biochemical modifications or at transcriptional level by regulating the gene expression of NCX1, NCX2, and NCX3 (Fig. 2).

Fig. 2 PI3-K/Akt1 transductional pathway activated by NGF acting on ncx1 gene expression and on NCX3 post-transductional regulation.

10 Pathophysiological Relevance in Stroke 10.1 NCX Gene and Protein Expression During Ischemia During focal ischemia NCX gene products are differently regulated in the ischemic core, in the peri-infarct area, as well as in intact brain regions. After permanent middle cerebral artery occlusion (pMCAO) all three transcripts NCX1, NCX2, and NCX3 are downregulated by 90% in the ischemic core, including prefrontal cortex (PFC) and part of the striatum, although NCX2 reduction occurs earlier [17]. In the other brain regions, belonging to the peri-infarct zone and including the infralimbic and prelimbic cortices and in the tenia tecta, NCX1 and NCX3 mRNAs display an upregulation. By contrast, in the same subregions, pMCAO elicited a mitigated decrease in NCX2 mRNA expression. In conclusion,

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pMCAO seems to affect NCX gene expression in a differential manner, depending on the exchanger isoform and brain regions involved in the insult. Interestingly, the pattern of expression of these three NCX transcripts mirrors the decline of the respective proteins in the core region [71]. The upregulation of NCX3 that occurs in the peri-infarct region, as opposed to NCX2 downregulation, has been interpreted as a compensatory mechanism induced by NCX3 isoform to counterbalance the reduced activity of the NCX2 protein thus counteracting the dysregulation of [Naþ]i and [Ca2þ]i homeostasis. Evidence for NCX3 neuroprotective role relies in the remarkable broadening of the infarct volume occurring when NCX3 protein is knocked down with a selective antisense oligonucleotide, thereby worsening the neurologic deficits [71]. Accordingly, it has been recently showed in ischemic NCX3–/– mice that NCX3 exerts a neuroprotective effect [73]. The discrepancy of NCX2 and NCX3 expression in response to stroke suggests, once again, that when NCX2 expression decreases in response to hypoxic conditions, NCX3 may assume a replacing function. Consistently, biochemical studies have clearly demonstrated that whereas NCX2 activity is strictly dependent on ATP levels, which are lowered during the development of brain ischemia, NCX3 is the only NCX gene product that is ATP independent [42, 43].

10.2 NCX Activity During In Vitro Anoxia Recent studies performed with the help of cloned cells stably expressing each isoform of the Naþ/Ca2þ exchanger, demonstrate that BHK cells transfected with the brain-specific NCX3 gene (BHK–NCX3), unlike NCX1(BHK–NCX1) and NCX2- (BHK–NCX2) transfected cells, are able to maintain [Ca2þ]i homeostasis during hypoxia plus re-oxygenation-induced Ca2þ overload, although ATP concentrations are reduced. In fact, hypoxia plus reoxygenation induces a lower increase of [Ca2þ]i in BHK–NCX3 than in BHK–NCX1 and BHK–NCX2. In addition, BHK–NCX3 are more resistant to chemical hypoxia plus re-oxygenation than BHK–NCX1 and BHK–NCX2. Interestingly, such augmented resistance can be counteracted by the selective silencing of the NCX3 isoform and by amiloride-derivative inhibitor of NCX, CB–DMB. The peculiar capability of NCX3 isoform to maintain [Ca2þ]i in the physiological range during hypoxic conditions might be correlated with its ability to work in the forward mode of operation in the presence of reduced ATP levels. In this regard, it has been reported that the NCX isoforms display a different sensitivity to ATP levels in the reverse mode of operation [43, 74]. Moreover, chemical hypoxia plus re-oxygenation produces a loss of mitochondrial membrane potential (Dc), a parameter related to the mitochondrial calcium homeostasis, in BHK–NCX1 and BHK–NCX2, but not in BHK–NCX3.

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Altogether, these results indicate that the NCX3 isoform, which is selectively expressed in the brain, more significantly contributes to the maintenance of [Ca2þ]i homeostasis during experimental conditions mimicking ischemia, thereby preventing mitochondrial Dc collapses and cell death. On the other hand, if anoxia is produced directly in primary cortical neurons, by means of oxygen glucose deprivation (OGD), the expression and the electrophysiological activity of the ubiquitous isoform NCX1 increases via nuclear factor kappa B (NF-kB). This increased activity seems to exert a neuroprotective role since NCX1, working in reverse mode of operation, promotes Ca2þ refilling into ER, thus preventing store depletion and consequent ER stress, which occurs after OGD [75].

10.3 Effect of Pharmacological NCX Modulation on Brain Ischemia In in vivo models, reproducing human cerebral ischemia through the occlusion of the middle cerebral artery, the inhibition of NCX, induced by Bepridil and by the selective amiloride derivative inhibitors CB–DMB [30], aggravates brain infarct whereas the activation of the antiporter with redox agents reduces the cerebral infarctual area [71]. At variance with these data, the inhibition of NCX with the isothiurea derivative KB-R7943 and SEA0400 reduces brain injury in the model of transient middle cerebral artery occlusion [76]. However, KB-R7943, besides blocking the antiporter, also produces a remarkable and prolonged hypothermic effect [71] that exerts, by itself, a relevant neuroprotective action in cerebral ischemia. On the other hand, by inhibiting other cellular ionic transport mechanisms and receptors, such as NMDA receptors and L-type Ca2þ channels [76], the same drug may yield a neuroprotective effect (Table 2). An explanation of the worsening effect exerted by NCX inhibitors relies on the different mode of action of this antiporter in the ischemic core and in the penumbra regions (Fig. 3). In particular, it is conceivable that, since in the penumbral region pump ATPase activity is still preserved because ATP levels are partially affected, NCX may likely operate in the forward mode. As a result, by extruding Ca2þ ions, the exchanger favors the entry of Naþ ions. Therefore, the pharmacological inhibition of NCX in this area reduces the extrusion of Ca2þ ions, thus enhancing Ca2þ-mediated cell injury. In contrast, in the ischemic core region, in which ATP levels are remarkably low and Naþ/Kþ ATPase activity is completely compromised, intracellular Naþ ions massively accumulate because of Naþ/Kþ ATPase failure. Hence, the intracellular Naþ loading promotes NCX to operate in the reverse mode as a Naþ efflux–Ca2þ influx pathway. This modality of operation may help brain cells to survive to the excessive [Naþ]i and water overload [71]. In conclusion, the NCX pharmacological inhibition in this core region further worsens the necrotic lesion of the

8.1

7.3

1–2.5 >30 0.05–0.09

0.3–30

0.1–1

Bepridil

CB-DMB

KB-R7943

SN-6

Glu-XIP

SEA0400

IC50 (mM)

Drug

Forward > Reverse mode

Forward > Reverse mode Forward > Reverse mode Forward > Reverse mode Forward > Reverse mode Reverse mode

Mode selectivity

Ala809, Val820, Gln826, Gly833, Asn839 Phe213, Gly833, Tyr224, Tyr226, Tyr228, Tyr231 Phe213, Gly833 and Asn839, Val227 and Val238 445–455

Amino acid regions

TMS5, a-2 repeat, XIP region, a-2 repeat f loop

a-2 repeat

Naþ-binding site

f loop

Site of action

Table 2 Molecular pharmacology of NCX inhibitors

NCX1, NCX2, NCX3

NCX1 > NCX3 > NCX2

NCX1 > NCX2

NCX1 = NCX3 > NCX2 NCX3 > NCX2 > NCX1

Isoform specificity

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Fig. 3 Hypothetical mode of operation of NCX in the penumbra and core regions of ischemic brain.

surviving glial and neuronal cells whereas the activation of NCX contributes to ameliorate ischemic damage.

10.4 Effect of NCX Knocking-Down and Knocking-Out on Brain Ischemia In rats bearing pMCAO, oligodeoxynucleotides blocking NCX1, NCX2, and NCX3, can induce different effects on the development of brain ischemia [71]. In fact, after pMCAO, the neuroprotective effect exerted by NCX is prevalently due to NCX1 and NCX3 products [71]. To determine the precise role of the different NCX genes in brain ischemia, genetically modified mice, lacking NCX1 or NCX2 or NCX3 have been generated. Unfortunately, targeted deletion of NCX1 results in NCX1-null embryos that do not have a spontaneous beating heart and die in utero. Studies conducted in NCX1 heterozygous mice that survive until adultness, showed that these animals did not exhibit increased vulnerability to ischemic damage, despite a reduction of NCX1 protein by almost 50% and of NCX activity by 60% [77]. On the other hand, mice deficient for NCX2 in the brain exhibit a worsening in the ischemic brain lesion induced through transient occlusion of MCA [78]. In in vitro ischemia, a significantly slower recovery in population spike amplitudes, a sustained elevation of [Ca2þ]i and an increased membrane

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depolarization were developed in the NCX2-deficient hippocampus [78]. The third isoform of the antiporter, NCX3, which is expressed selectively in the brain and in the muscle and is able to work also in absence of ATP, plays a relevant role in the ischemic process. In fact, it has been recently shown that ncx3 gene ablation leads to a worsening of brain damage after focal ischemia [73]. In addition, cortical neurons and hippocampal organotypic cultures from ncx3–/– mice display a massive neuronal death when exposed to OGD plus re-oxygenation [73].

11 Therapeutic Perspectives Nevertheless, the great effort to find new and more effective therapeutic interventions in ischemic stroke, the only available pharmacological treatment remains the administration of rTPA that must be administered within the therapeutical window of 3 h from stroke onset. The rationale of targeting NCX activity and/or expression lies on the possibility to pharmacologically modulate this crucial regulator of intracellular Ca2þ and Naþ homeostasis, in order to help neurons and glial cells to survive in the critical penumbra area and to save the major number of cells in the ischemic core. Indeed, most of the preclinical studies specifically targeting NCX isoforms and performed with cellular and animal models reproducing human ischemia suggest that an enhancement of the expression and the activity of CNS NCX isoforms may contribute to reduce the ischemic damage. In fact, their inhibition increases brain vulnerability to the insult. In the last 40 years, most of the medicinal chemistry studies have been devoted to the development of compounds inhibiting global NCX activity in the hope of obtaining compounds active in heart ischemia. However, the new data obtained in cerebral ischemia suggest that pharmacological activators rather than NCX inhibitors may help neurons and glial cells to overcome the stroke-mediated ionic dysregulation. This strategy may be pursued acting at the protein, transductional or transcriptional levels (Fig. 4). First, at protein level, it would be conceivable to identify a functional moiety of drugs working as selective activator of those cerebral NCX isoforms involved in the process of brain damage. This could be achieved by means of quantitative structure relationship activity approach starting from the molecular structure of already available NCX antagonists. Another approach could be pursued by developing cargo compounds consisting of a carrier able to cross the blood–brain barrier and of a molecule provided with neurotrophic factor properties, such as NGF. This new chemical entity could activate NGF/TrkA/Shc/PI3-K/Akt transductional pathway in the brain by stimulating tyrosine kinase receptors, thus inducing an increased expression and activity of NCX1 and NCX3 isoforms [67].

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Fig. 4 Potential steps in the pharmacological regulation of NCX isoform expression and activity.

12 Conclusions An increasing amount of evidence seems to suggest that NCX plays a pivotal role in maintaining intracellular Naþ and Ca2þ homeostasis during pathophysiological conditions in the brain. Indeed, it still remains to be fully clarified whether it is the suppression or the activation of the exchanger that yields potentially beneficial effects on a number of neurodegenerative diseases, such as ischemia, Alzheimer’s disease, aging, and white matter trauma. A number of conflicting results on the modulating effects of the exchanger have highlighted such differences. For instance, the severity and extension of brain injury may vary depending on whether the exchanger is activated or inhibited. Specifically, in animal models of cerebral ischemia, consequent to permanent vascular occlusion, the pharmacological activation of NCX reduces brain damage whereas drugs provided with inhibitory properties aggravate the infarct lesion as well as transgenic mice lacking ncx2 or ncx3 gene. Data derived from drug strategy and knockout mice seem to indicate that the possible development of agents able to selectively activate each of the three NCX gene products might achieve a more promising therapeutic goal. Unfortunately, although efforts to synthesize organic compounds that target the different NCX gene products have been made, no highly specific agents have yet been available. A further obstacle underlying the pharmacological modulation of brain NCX activity is represented by the difficulty of compounds to cross the blood–brain barrier. In this regard, promising strategies entail the conjugation of organic and

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peptidergic drugs with carrier molecules that are substrates for transporters present on the cell membranes of blood–brain barrier and neurons [79]. In conclusion, although the studies concerning the role played by NCX in the pathological mechanisms of brain injury during neurodegenerative diseases have had a late beginning compared with those concerning heart disease, the availability of pharmacological agents able to selectively modulate each NCX subtype activity and antiporter mode of operation will enable researchers to gain fundamental insights into its pathophysiological role and, thus, to obtain more tangible perspectives on how to treat such neurological disorders.

References 1. Schnetkamp PP. Na-Ca or Na-Ca-K exchange in rod photoreceptors. Prog Biophys Mol Biol 54: 1–29, 1989. 2. Lytton J. How many sodium ions does it take to turn an exchanger? J Physiol 545: 335, 2002. 3. Cunningham KW and Fink GR. Calcineurin inhibits VCX1-dependent H+/Ca2+ exchange and induces Ca2+ ATPases in Saccharomyces cerevisiae. Mol Cell Biol 16: 2226–2237, 1996. 4. Pozos TC, Sekler I, and Cyert MS. The product of HUM1, a novel yeast gene, is required for vacuolar Ca2+/H+ exchange and is related to mammalian Na+/Ca2+ exchangers. Mol Cell Biol 16: 3730–3741, 1996. 5. Annunziato L, Pignataro G, and Di Renzo GF. Pharmacology of brain Na+/Ca2+ exchanger: from molecular biology to therapeutic perspectives. Pharmacol Rev 56: 633–654, 2004. 6. Philipson KD and Nicoll DA. Sodium-calcium exchange: a molecular perspective. Annu Rev Physiol 62: 111–133, 2000. 7. Philipson KD, Nicoll DA, Ottolia M, Quednau BD, Reuter H, John S, and Qiu Z. The Na+/Ca2+ exchange molecule: an overview. Ann N Y Acad Sci 976: 1–10, 2002. 8. Nicoll DA, Longoni S, and Philipson KD. Molecular cloning and functional expression of the cardiac sarcolemmal Na+-Ca2+ exchanger. Science 250: 562–565, 1990. 9. Li Z, Matsuoka S, Hryshko LV, Nicoll DA, Bersohn MM, Burke EP, Lifton RP, and Philipson KD. Cloning of the NCX2 isoform of the plasma membrane Na+-Ca2+ exchanger. J Biol Chem 269: 17434–17439, 1994. 10. Nicoll DA, Quednau BD, Qui Z, Xia YR, Lusis AJ, and Philipson KD. Cloning of a third mammalian Na+-Ca2+ exchanger, NCX3. J Biol Chem 271: 24914–24921, 1996. 11. Kraev A, Chumakov I, and Carafoli E. The organization of the human gene NCX1 encoding the sodium-calcium exchanger. Genomics 37: 105–112, 1996. 12. Lee SL, Yu AS, and Lytton J. Tissue-specific expression of Na+-Ca2+ exchanger isoforms. J Biol Chem 269: 14849–14852, 1994. 13. Nicholas SB, Yang W, Lee SL, Zhu H, Philipson KD, and Lytton J. Alternative promoters and cardiac muscle cell-specific expression of the Na+/Ca2+ exchanger gene. Am J Physiol 274: H217–H232, 1998. 14. Scheller T, Kraev A, Skinner S, and Carafoli E. Cloning of the multipartite promoter of the sodium-calcium exchanger gene NCX1 and characterization of its activity in vascular smooth muscle cells. J Biol Chem 273: 7643–7649, 1998. 15. Sehgal A, Patil N, and Chao M. A constitutive promoter directs expression of the nerve growth factor receptor gene. Mol Cell Biol 8: 3160–3167, 1988. 16. Li L, Guerini D, and Carafoli E. Calcineurin controls the transcription of Na+/Ca2+ exchanger isoforms in developing cerebellar neurons. J Biol Chem 275: 20903–20910, 2000.

84

L. Annunziato et al.

17. Boscia F, Gala R, Pignataro G, de Bartolomeis A, Cicale M, Ambesi-Impiombato A, Di Renzo G, and Annunziato L. Permanent focal brain ischemia induces isoform-dependent changes in the pattern of Na+/Ca2+ exchanger gene expression in the ischemic core, periinfarct area, and intact brain regions. J Cereb Blood Flow Metab 26: 502–517, 2006. 18. Hossmann KA. Excitotoxic mechanisms in focal ischemia. Adv Neurol 71: 69–74, 1996. 19. Gabellini N, Bortoluzzi S, Danieli GA, and Carafoli E. Control of the Na+/Ca2+ exchanger 3 promoter by cyclic adenosine monophosphate and Ca2+ in differentiating neurons. J Neurochem 84: 282–293, 2003. 20. Gabellini N. Transcriptional regulation by cAMP and Ca2+ links the Na+/Ca2+ exchanger 3 to memory and sensory pathways. Mol Neurobiol 30: 91–116, 2004. 21. Sugiura S, Kitagawa K, Omura-Matsuoka E, Sasaki T, Tanaka S, Yagita Y, Matsushita K, Storm DR, and Hori M. CRE-mediated gene transcription in the peri-infarct area after focal cerebral ischemia in mice. J Neurosci Res 75: 401–407, 2004. 22. Gomez-Villafuertes R, Torres B, Barrio J, Savignac M, Gabellini N, Rizzato F, Pintado B, Gutierrez-Adan A, Mellstrom B, Carafoli E, and Naranjo JR. Downstream regulatory element antagonist modulator regulates Ca2+ homeostasis and viability in cerebellar neurons. J Neurosci 25: 10822–10830, 2005. 23. Nicoll DA, Ottolia M, Lu L, Lu Y, and Philipson KD. A new topological model of the cardiac sarcolemmal Na+-Ca2+ exchanger. J Biol Chem 274: 910–917, 1999. 24. Nicoll DA, Hryshko LV, Matsuoka S, Frank JS, and Philipson KD. Mutation of amino acid residues in the putative transmembrane segments of the cardiac sarcolemmal Na+-Ca2+ exchanger. J Biol Chem 271: 13385–13391, 1996. 25. Ottolia M, Nicoll DA, and Philipson KD. Mutational analysis of the alpha-1 repeat of the cardiac Na+-Ca2+ exchanger. J Biol Chem 280: 1061–1069, 2005. 26. Hilge M, Aelen J, and Vuister GW. Ca2+ regulation in the Na+/Ca2+ exchanger involves two markedly different Ca2+ sensors. Mol Cell 22: 15–25, 2006. 27. Maack C, Ganesan A, Sidor A, and O’Rourke B. Cardiac sodium-calcium exchanger is regulated by allosteric calcium and exchanger inhibitory peptide at distinct sites. Circ Res 96: 91–99, 2005. 28. Matsuoka S, Nicoll DA, He Z, and Philipson KD. Regulation of cardiac Na+-Ca2+ exchanger by the endogenous XIP region. J Gen Physiol 109: 273–286, 1997. 29. Li Z, Nicoll DA, Collins A, Hilgemann DW, Filoteo AG, Penniston JT, Weiss JN, Tomich JM, and Philipson KD. Identification of a peptide inhibitor of the cardiac sarcolemmal Na+-Ca2+ exchanger. J Biol Chem 266: 1014–1020, 1991. 30. Pignataro G, Tortiglione A, Scorziello A, Giaccio L, Secondo A, Severino B, Santagada V, Caliendo G, Amoroso S, Di Renzo G, and Annunziato L. Evidence for a protective role played by the Na+/Ca2+ exchanger in cerebral ischemia induced by middle cerebral artery occlusion in male rats. Neuropharmacology 46: 439–448, 2004. 31. Santacruz-Toloza L, Ottolia M, Nicoll DA, and Philipson KD. Functional analysis of a disulfide bond in the cardiac Na+-Ca2+ exchanger. J Biol Chem 275: 182–188, 2000. 32. Walz T, Hirai T, Murata K, Heymann JB, Mitsuoka K, Fujiyoshi Y, Smith BL, Agre P, and Engel A. The three-dimensional structure of aquaporin-1. Nature 387: 624–627, 1997. 33. Ottolia M, John S, Qiu Z, and Philipson KD. Split Na+-Ca2+ exchangers. Implications for function and expression. J Biol Chem 276: 19603–19609, 2001. 34. Quednau BD, Nicoll DA, and Philipson KD. Tissue specificity and alternative splicing of the Na+/Ca2+ exchanger isoforms NCX1, NCX2, and NCX3 in rat. Am J Physiol 272: C1250–C1261, 1997. 35. Yu SP and Choi DW. Na+-Ca2+ exchange currents in cortical neurons: concomitant forward and reverse operation and effect of glutamate. Eur J Neurosci 9: 1273–1281, 1997. 36. Canitano A, Papa M, Boscia F, Castaldo P, Sellitti S, Taglialatela M, and Annunziato L. Brain distribution of the Na+/Ca2+ exchanger-encoding genes NCX1, NCX2, and NCX3 and their related proteins in the central nervous system. Ann N Y Acad Sci 976: 394–404, 2002.

The Naþ/Ca2þ Exchanger

85

37. Papa M, Canitano A, Boscia F, Castaldo P, Sellitti S, Porzig H, Taglialatela M, and Annunziato L. Differential expression of the Na+-Ca2+ exchanger transcripts and proteins in rat brain regions. J Comp Neurol 461: 31–48, 2003. 38. Reeves JP and Hale CC. The stoichiometry of the cardiac sodium-calcium exchange system. J Biol Chem 259: 7733–7739, 1984. 39. Kang TM and Hilgemann DW. Multiple transport modes of the cardiac Na+/Ca2+ exchanger. Nature 427: 544–548, 2004. 40. Hilgemann DW, Nicoll DA, and Philipson KD. Charge movement during Na+ translocation by native and cloned cardiac Na+/Ca2+ exchanger. Nature 352: 715–718, 1991. 41. Blaustein MP and Lederer WJ. Sodium/calcium exchange: its physiological implications. Physiol Rev 79: 763–854, 1999. 42. Secondo A, Staiano RI, Scorziello A, Sirabella R, Boscia F, Adornetto A, Valsecchi V, Molinaro P, Canzoniero LM, Di Renzo G, and Annunziato L. BHK cells transfected with NCX3 are more resistant to hypoxia followed by reoxygenation than those transfected with NCX1 and NCX2: Possible relationship with mitochondrial membrane potential. Cell Calcium 42: 521–535, 2007. 43. Linck B, Qiu Z, He Z, Tong Q, Hilgemann DW, and Philipson KD. Functional comparison of the three isoforms of the Na+/Ca2+ exchanger (NCX1, NCX2, NCX3). Am J Physiol 274: C415–C423, 1998. 44. Juhaszova M, Shimizu H, Borin ML, Yip RK, Santiago EM, Lindenmayer GE, and Blaustein MP. Localization of the Na+-Ca2+ exchanger in vascular smooth muscle, and in neurons and astrocytes. Ann N Y Acad Sci 779: 318–335, 1996. 45. Finkbeiner SM. Glial calcium. Glia 9: 83–104, 1993. 46. Goldman WF, Yarowsky PJ, Juhaszova M, Krueger BK, and Blaustein MP. Sodium/ calcium exchange in rat cortical astrocytes. J Neurosci 14: 5834–5843, 1994. 47. Holgado A and Beauge L. Effects of external monovalent cations on Na+-Ca2+ exchange in cultured rat glial cells. Ann N Y Acad Sci 779: 279–281, 1996. 48. Takuma K, Matsuda T, Hashimoto H, Asano S, and Baba A. Cultured rat astrocytes possess Na+-Ca2+ exchanger. Glia 12: 336–342, 1994. 49. Reuter H, Henderson SA, Han T, Matsuda T, Baba A, Ross RS, Goldhaber JI, and Philipson KD. Knockout mice for pharmacological screening: testing the specificity of Na+-Ca2+ exchange inhibitors. Circ Res 91: 90–92, 2002. 50. Jeon D, Yang YM, Jeong MJ, Philipson KD, Rhim H, and Shin HS. Enhanced learning and memory in mice lacking Na+/Ca2+ exchanger 2. Neuron 38: 965–976, 2003. 51. Sokolow S, Manto M, Gailly P, Molgo J, Vandebrouck C, Vanderwinden JM, Herchuelz A, and Schurmans S. Impaired neuromuscular transmission and skeletal muscle fiber necrosis in mice lacking Na/Ca exchanger 3. J Clin Invest 113: 265–273, 2004. 52. Cragoe EJ Jr., Woltersdorf OW, Jr, Bicking JB, Kwong SF, and Jones JH. Pyrazine diuretics. II. N-amidino-3-amino-5-substituted 6-halopyrazinecarboxamides. J Med Chem 10: 66–75, 1967. 53. Baker PF and Blaustein MP. Sodium-dependent uptake of calcium by crab nerve. Biochim Biophys Acta 150: 167–170, 1968. 54. Sharikabad MN, Cragoe EJ Jr., and Brors O. Inhibition by 5-N-(4-chlorobenzyl)-2’, 4’-dimethylbenzamil of Na+/Ca2+ exchange and L-type Ca2+ channels in isolated cardiomyocytes. Pharmacol Toxicol 80: 57–61, 1997. 55. Tewes F, Munnier E, Antoon B, Ngaboni Okassa L, Cohen-Jonathan S, Marchais H, Douziech-Eyrolles L, Souce M, Dubois P, and Chourpa I. Comparative study of doxorubicin-loaded poly(lactide-co-glycolide) nanoparticles prepared by single and double emulsion methods. Eur J Pharm Biopharm 66: 488–492, 2007. 56. Taglialatela M, Amoroso S, Canzoniero LM, Di Renzo GF, and Annunziato L. Membrane events and ionic processes involved in dopamine release from tuberoinfundibular neurons. II. Effect of the inhibition of the Na+-Ca++ exchange by amiloride. J Pharmacol Exp Ther 246: 689–694, 1988.

86

L. Annunziato et al.

57. Taglialatela M, Canzoniero LM, Cragoe EJ, Jr, Di Renzo G, and Annunziato L. Na+-Ca2+ exchange activity in central nerve endings. II. Relationship between pharmacological blockade by amiloride analogues and dopamine release from tuberoinfundibular hypothalamic neurons. Mol Pharmacol 38: 393–400, 1990. 58. Amoroso S, Taglialatela M, Canzoniero LM, Cragoe EJ, Jr, di Renzo G, and Annunziato L. Possible involvement of Ca++ ions, protein kinase C and Na+-H+ antiporter in insulin-induced endogenous dopamine release from tuberoinfundibular neurons. Life Sci 46: 885–894, 1990. 59. Watano T, Kimura J, Morita T, and Nakanishi H. A novel antagonist, No. 7943, of the Na+/Ca2+ exchange current in guinea-pig cardiac ventricular cells. Br J Pharmacol 119: 555–563, 1996. 60. Iwamoto T and Shigekawa M. Differential inhibition of Na+/Ca2+ exchanger isoforms by divalent cations and isothiourea derivative. Am J Physiol 275: C423–C430, 1998. 61. Iwamoto T, Watano T, and Shigekawa M. A novel isothiourea derivative selectively inhibits the reverse mode of Na+/Ca2+ exchange in cells expressing NCX1. J Biol Chem 271: 22391–22397, 1996. 62. Sobolevsky AI and Khodorov BI. Blockade of NMDA channels in acutely isolated rat hippocampal neurons by the Na+/Ca2+ exchange inhibitor KB-R7943. Neuropharmacology 38: 1235–1242, 1999. 63. Liang GH, Kim JA, Seol GH, Choi S, and Suh SH. The Na+/Ca(2+) exchanger inhibitor KB-R7943 activates large-conductance Ca2+-activated K+ channels in endothelial and vascular smooth muscle cells. Eur J Pharmacol 582: 35–41, 2008. 64. Li Y, Woo V, and Bose R. Platelet hyperactivity and abnormal Ca2+ homeostasis in diabetes mellitus. Am J Physiol Heart Circ Physiol 280: H1480–H1489, 2001. 65. Amoroso S, Tortiglione A, Secondo A, Catalano A, Montagnani S, Di Renzo G, and Annunziato L. Sodium nitroprusside prevents chemical hypoxia-induced cell death through iron ions stimulating the activity of the Na+-Ca2+ exchanger in C6 glioma cells. J Neurochem 74: 1505–1513, 2000. 66. Secondo A, Staiano IR, Scorziello A, Sirabella R, Boscia F, Adornetto A, Canzoniero LM, Di Renzo G, and Annunziato L. The Na+/Ca2+ exchanger isoform 3 (NCX3) but not isoform 2 (NCX2) and 1 (NCX1) singly transfected in BHK cells plays a protective role in a model of in vitro hypoxia. Ann N Y Acad Sci 1099: 481–485, 2007. 67. Formisano L, Saggese M, Secondo A, Sirabella R, Vito P, Valsecchi V, Molinaro P, Di Renzo G, and Annunziato L. The two isoforms of the Na+/Ca2+ exchanger, NCX1 and NCX3, constitute novel additional targets for the prosurvival action of Akt/protein kinase B pathway. Mol Pharmacol 73: 727–737, 2008. 68. Tortiglione A, Picconi B, Barone I, Centonze D, Rossi S, Costa C, Di Filippo M, Tozzi A, Tantucci M, Bernardi G, Annunziato L, and Calabresi P. Na+/Ca2+ exchanger maintains ionic homeostasis in the peri-infarct area. Stroke 38: 1614–1620, 2007. 69. Yamashita J, Kita S, Iwamoto T, Ogata M, Takaoka M, Tazawa N, Nishikawa M, Wakimoto K, Shigekawa M, Komuro I, and Matsumura Y. Attenuation of ischemia/ reperfusion-induced renal injury in mice deficient in Na+/Ca2+ exchanger. J Pharmacol Exp Ther 304: 284–293, 2003. 70. Slodzinski MK, Juhaszova M, and Blaustein MP. Antisense inhibition of Na+/Ca2+ exchange in primary cultured arterial myocytes. Am J Physiol 269: C1340–C1345, 1995. 71. Pignataro G, Gala R, Cuomo O, Tortiglione A, Giaccio L, Castaldo P, Sirabella R, Matrone C, Canitano A, Amoroso S, Di Renzo G, and Annunziato L. Two sodium/ calcium exchanger gene products, NCX1 and NCX3, play a major role in the development of permanent focal cerebral ischemia. Stroke 35: 2566–2570, 2004. 72. Reeves JP, Bailey CA, and Hale CC. Redox modification of sodium-calcium exchange activity in cardiac sarcolemmal vesicles. J Biol Chem 261: 4948–4955, 1986. 73. Molinaro P, Cuomo O, Pignataro G, Boscia F, Sirabella R, Pannaccione A, Secondo A, Scorziello A, Adornetto A, Gala R, Viggiano D, Sokolow S, Herchuelz A, Schurmans S,

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74.

75.

76.

77.

78.

79.

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Di Renzo G, and Annunziato L. Targeted disruption of Na+/Ca2+ exchanger 3 (NCX3) gene leads to a worsening of ischemic brain damage. J Neurosci 28: 1179–1184, 2008. Condrescu M, Gardner JP, Chernaya G, Aceto JF, Kroupis C, and Reeves JP. ATPdependent regulation of sodium-calcium exchange in Chinese hamster ovary cells transfected with the bovine cardiac sodium-calcium exchanger. J Biol Chem 270: 9137–9146, 1995. Sirabella R, Secondo A, Pannaccione A, Scorziello A, Valsecchi V, Adornetto A, Bilo L, Di Renzo G, and Annunziato L. Anoxia-incuced NF-kB-dependent upregulation of NCX1 contributes to Ca2+ refilling into endoplasmic reticulum in cortical neurons. Stroke 40: 922–929, 2009. Matsuda T, Arakawa N, Takuma K, Kishida Y, Kawasaki Y, Sakaue M, Takahashi K, Takahashi T, Suzuki T, Ota T, Hamano-Takahashi A, Onishi M, Tanaka Y, Kameo K, and Baba A. SEA0400, a novel and selective inhibitor of the Na+-Ca2+ exchanger, attenuates reperfusion injury in the in vitro and in vivo cerebral ischemic models. J Pharmacol Exp Ther 298: 249–256, 2001. Luo J, Wang Y, Chen X, Chen H, Kintner DB, Shull GE, Philipson KD, and Sun D. Increased tolerance to ischemic neuronal damage by knockdown of Na+-Ca2+ exchanger isoform 1. Ann N Y Acad Sci 1099: 292–305, 2007. Jeon D, Chu K, Jung KH, Kim M, Yoon BW, Lee CJ, Oh U, and Shin HS. Na+/Ca2+ exchanger 2 is neuroprotective by exporting Ca2+ during a transient focal cerebral ischemia in the mouse. Cell Calcium 43: 482–491, 2008. Rochat B and Audus KL. Drug disposition and targeting. Transport across the bloodbrain barrier. Pharm Biotechnol 12: 181–200, 1999.

The ‘‘Loop’’ Diuretic Drug Bumetanide-Sensitive Naþ-Kþ-Cl– Cotransporter in Cerebral Ischemia Dandan Sun

1 Introduction The Naþ-Kþ-Cl– cotransporter (NKCC) belongs to the cation-chloride cotransporter superfamily (CCC), which mediates the coupled movement of Naþ and /or Kþ with Cl– across the plasma membrane [1–3]. Two isoforms of NKCC (NKCC1 and NKCC2) have been identified as two different gene products, human genes SLC12A1 (NKCC2) and SLC12A2 (NKCC1) [1,4]. Together with the thiazide-sensitive renal Naþ-Cl– cotransporter (NCC; SLC12A3), they constitute the Naþ-dependent branch of the CCC superfamily. NKCC1 and NKCC2 share 50–60% amino acid sequence homology and have distinct tissue distributions. NKCC1, which was first identified on the basolateral membranes of secretory epithelial cells, is referred to as the ‘‘secretory’’ or ‘‘house-keeping’’ NKCC [1,4]. In addition to secretory epithelial cells, NKCC1 is expressed in a wide variety of tissues including epithelial cells and nonepithelial cells, such as neurons and glia [2,5]. NKCC1 plays a role in salt secretion and absorption, cell volume regulation, and maintenance of intracellular Cl– concentration ([Cl–]i) [1,2,5]. NKCC1-null mice exhibit multiple tissue phenotype changes and functional disorders, including deafness, decreased blood pressure, intestinal bleeding, infertility, and salivary secretion reduction [5–7]. These findings imply that NKCC1 is important in cellular function in a wide variety of tissues. In contrast to NKCC1, NKCC2 expression is confined to the apical membrane of the epithelial cells of the thick ascending limb of Henle’s loop in kidney [1]. Thus, NKCC2 is important in regulation of extracellular fluid volume and osmolarity. NKCC2 mutation is associated with Bartter syndrome, a renal tubular disorder [5]. Loss of ionic homeostasis plays a central role in pathogenesis of ischemic cell damage. During ischemia, intracellular Kþ decreases, while intracellular Ca2+, D. Sun (*) Department of Neurological Surgery, University of Wisconsin School of Medicine and Public Health, Madison, WI, 53705, USA e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_6, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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Naþ, Cl–, along with H2O increase [8]. Ischemia-induced perturbation of ionic homeostasis leads to subsequent activation of proteases, phospholipases, and formation of oxygen and nitrogen free radicals. This cascade of signal transduction events results in long-term functional and structural changes in membrane and cytoskeletal integrity and eventual cell death. Results from in vitro and in vivo studies demonstrate a role for NKCC1 in ischemia-induced dissipation of ionic homeostasis. Therefore, the ion transport protein could provide a novel target for the future development of therapeutic treatment of ischemic cerebral damage. This chapter focuses on the role of NKCC1 in the central nervous system (CNS), especially under ischemic pathophysiological conditions. Interested readers are referred to a series of reviews that cover broader aspects of NKCC and Kþ-dependent chloride transporters [2,5,9].

2 NKCC1 Gene Structure and Structural Features Using cDNA probes, a NKCC1 human homolog has been identified in human colonic carcinoma line T84 cell [10]. Analysis of human NKCC1 (hNKCC1) gene SLC12A2 (3,498 kb) found that hNKCC1 is located in humans at chromosome 5q23 and composed of 1212 amino acids with a molecular mass of 132 kDa. The primary structure of hNKCC1 is 74% identical to the sequence of shark rectal gland Na-K-Cl cotransporter (sNKCC1) and 91% identical to a mouse Na-K-Cl cotransporter (mNKCC1) [10]. The molecular structure of NKCC1 has been proposed based on cDNA sequence analysis. The sequence analysis of sNKCC1 reveals that NKCC1 protein has 12 putative transmembrane domains (TM), flanked by large cytoplasmic amino and carboxyl termini, which contain regulatory phosphorylation sites [4,1] (Fig. 1). There are two potential N-linked glycosylation sites predicted on an extracellular loop between putative transmembrane segments 7 and 8 [4]. NKCC1 in different tissues have molecular weights that vary from 120 to 190 kDa [2]. They form homodimers in plasma membrane via COOH-terminus [11]. The proposed topology is supported by the recent experimental data using an in vitro translation system [12].

3 Molecular Biology Site-directed mutagenesis studies show that changes in Naþ and Rbþ (Kþ) affinity result from changes in TM2 whereas Cl– affinity was affected by changes in TM4–7 [13]. The affinity for bumetanide, a potent inhibitor of NKCC, was found to be affected by residues in the TM2–7 region, and also by residues in TM11 and TM12 [13]. In contrast, the conserved hydrophilic C-termini of hNKCC1 and sNKCC1 are not involved in the difference in affinities for the transported ions [14]. Therefore, the sequences within TM2 segments affect

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Fig. 1 Model of Naþ-Kþ-Cl– cotransporter protein based on its hydropathy profile. The NKCC protein is comprised 12 transmembrane (TM)-spanning domains, numbered TM1TM12. Glycosylation sites are predicted on an extracellular loop between putative transmembrane segments 7 and 8. Identified phosphorylated threonine residues are indicated by ‘‘*’’ with an adjacent ‘‘p’’ (produced by the kind permission of Curr. Opin. Nephrol. Hypertens from [1])

transport affinity for cations, but not for Cl–. The bumetanide binding probably requires conformational interaction between TM and extracellular domains.

4 Tissue and Cellular Distribution Tissue distribution analysis by Northern blot revealed presence of NKCC1 transcripts in all tissues [9]. Because of NKCC1’s location in basolateral membrane of epithelia, NKCC1 is important in transcellular ion transport in secretory epithelia. In the CNS, NKCC1 is expressed in neurons, glia, choroid plexus, and cerebral vascular endothelial cells [3,9]. In addition to salt transport, NKCC1 is also involved in maintenance and regulation of cell volume in both epithelial and nonepithelial cells. Moreover, NKCC1 functions in accumulation of intracellular Cl– and regulation of Cl– homeostasis in the CNS. Therefore, NKCC1 is important in GABAergic signaling and changes of neuronal excitability [15]. The role of NKCC1 in epileptic formation has been reviewed recently [16].

5 Biophysical and Electrophysiological Properties There are three fundamental characteristics of NKCC function [2]. First, ion translocation by NKCC requires that all three ions (Naþ, Kþ, Cl–) are simultaneously present on the same side of the membrane [2]. Second, ion transport mediated by NKCC is electroneutral in most cases, with a stoichiometry of 1

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Naþ:1Kþ:2 Cl– [2]. The driving force for ion influx is supplied by the combined chemical gradients for three transported ions [2], with a net electrochemical potential.

ð

Dm ¼ RT ln

½Naþ o ½Kþ o ½Cl 2o ½Naþ i ½Kþ i ½Cl 2i

Þ

:

Third, NKCCs are targets of the ‘‘loop’’ diuretics. Sulfamoybenzoic acid loop diuretics such as bumetanide bind to NKCCs and can inhibit ion translocation of three ions [17]. Bumetanide (IC50 0.3 mm) is the most widely used drug in NKCC1 inhibition and more potent than other sulfamoybenzoic acid loop diuretics such as furosemide (IC50 7 mm) [18].

6 Regulatory Mechanisms – Regulation of NKCC1 via Phosphorylation NKCC1 activity is regulated by multiple stimuli such as a decrease in [Cl–]i, hypertonic stress, an increase in [Ca2+]i, and a-adrenergic receptor stimulation [19–22]. Cl–-mediated regulation of NKCC1 activity has been extensively studied in epithelial cells. Epithelium secrets Cl– through Cl– channels located on the apical membrane. A decrease in [Cl–]i will increase the activity of basolateral membrane NKCC1, which will bring Cl– into the cell, thereby promoting continuous Cl– secretion across the apical membrane [2]. Lytle et al. have demonstrated that a decrease in [Cl]i stimulates both NKCC1 activity and NKCC1 protein phosphorylation in epithelial cells [23,24]. The degree of NKCC1 activation is closely related to its phosphorylation and this suggests that the Cl–-mediated regulation of NKCC1 activity is via phosphorylation [24]. Sequence analysis has revealed that Thr184, Thr189, Thr202, and Thr1114 of N- and C-termini are the conserved regulatory phosphoracceptor residues in NKCC1 [4,19]. Both kinases and phosphatases contribute to the precise regulation of NKCC1 activity through their opposite effects on NKCC1. Recently, several kinases have been discovered to mediate NKCC1 phosphorylation. OSR1 (oxidative stress response kinase) and SPAK (STE20/SPS-1 related, proline–alanine-rich kinase), along with its rat homolog PASK (proline–alanine-rich STE20-related kinase), were found to bind to the third conserved domain in the NKCC1 N-terminus and co-immunoprecipitated with NKCC1 protein [25,26]. Overexpression of PASK in HEK cells causes a small but significant increase in both hNKCC1 and sNKCC1 activity and the phosphorylation of PASK coincides with that of NKCC1 [25]. PASK and SPAK are preferentially expressed in the brain and in cells that are active in ion transport and rich in Naþ/Kþ-ATPase [26–28]. PASK is suggested as a stress-related kinase due to its putative caspase cleavage site and

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its ability to activate p38 MAPK pathway [28]. Neurons, glia, and epithelial choroid cells in the CNS have shown PASK expression [29]. On the other hand, protein phosphatase is responsible for dephosphorylation of NKCC1. The catalytic subunit of protein phosphatase 1 binds to the proteins that contain the consensus motif RVXFXD [30]. The N-terminus of NKCC1 contains a highly conserved RVNFVD sequence both in sNKCC1 (107–112) and hNKCC1 (140–145) [30]. The RVXFVD motif mutation produces an activation shift in NKCC1 regulation by Cl–. Dowd et al. [25] have shown that dominant negative PASK (DNPASK) decreases the activity of hNKCC1 by 80% and sNKCC1 by 60%. In addition, protein phosphatase type 1 inhibitor calyculin A nearly completely restores NKCC1 activity in the DNPASK transfected cells [25]. These findings strongly suggest that an alteration in phosphorylation/ dephosphorylation of NKCC1 regulates NKCC1 activity. Recent studies have demonstrated that a novel WNK family of serine–threonine kinases (kinases with no K = lysine, which is substituted with cysteine) plays an important role in the regulation of epithelial Naþ channels and ion transporters including NKCC1 [16]. In humans, there are four genes encoding WNK1, WNK2, WNK3, and WNK 4, respectively. WNK3 is most highly expressed in brain. Co-expression of WNK3 and NKCC1 results in an increase in NKCC1 activity and induces robust phosphorylation of threonine212 and threonine217, the known regulatory sites in NKCC1 [31]. Moreover, it has been suggested that WNK1/WNK4 can phosphorylate SPAK/OSR1, which in turn directly phosphorylates NKCC1 [16]. Therefore, NKCC1 function can be dynamically modulated by changing activities of the kinases and phosphatases.

7 Physiological Properties – NKCC1-Mediated Regulation of Kþ Uptake, Intracellular Cl–, and Cell Volume in the CNS NKCC1 is expressed in neurons, glia (astrocytes and oligodendrocytes), as well as choroid plexus epithelial cells and blood vessel endothelial cells [32–35]. Functional studies demonstrate that NKCC1 plays an important role in regulation of [Cl–]i, Kþ uptake, and cell volume of cells in the CNS [15,36]. g-Aminobutyric acid (GABA) and glycine are the main inhibitory neurotransmitters in the adult mammalian CNS. Activation of GABA receptors results in Cl– influx and hyperpolarization of the plasma membrane. However, during fetal and postnatal development, the equilibrium potential for Cl– is more positive than the resting potential and activation of GABA receptors triggers an excitatory response and a Cl– efflux [37–39]. The excitatory action of GABA in the immature CNS is important for the development of the nervous system [40]. Cells maintain Cl– homeostasis through the fine balance between inwardly directed Cl– flux and Cl–extrusion systems. NKCC1 functions as a Cl– influx system and contributes to the active accumulation of intracellular Cl– in

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immature neurons [15,39]. An emerging hypothesis asserts that the balance between the inwardly directed NKCC1 and the outwardly directed neuronal K-Cl cotransporter (KCC2) may be important in the determination of GABAA receptor-mediated depolarizing responses. This view is supported by the findings that expression of the inwardly directed NKCC1 in neurons precedes expression of the outwardly directed KCC2 [41]. Additionally, genetic ablation of NKCC1 leads to a decrease in resting [Cl–]i and abolishes the GABAmediated depolarization in mouse dorsal root ganglion cells [15]. Disruption of KCC2 results in frequent seizures, abolishment of respiration, and early lethality in mice due to anomalous excitatory actions of GABA and glycine [42]. Opening of the GABA-activated Cl– channel or depletion of intracellular Cl– significantly stimulates NKCC1 activity in neurons [21]. The GABA-mediated loss of intracellular Cl–, but not a subsequent membrane depolarization or shrinkage, leads to stimulation of NKCC1 [22]. This stimulation might be an important positive feedback mechanism to maintain intracellular Cl– level and GABA function in immature neurons. The depolarizing effect of GABA is of functional importance during neuronal maturation and differentiation. The depolarizing action of GABA subsequently activates NMDA and voltage-gated Ca2+ channels that leads to a rise in intracellular Ca2+ and activation of a wide range of intracellular signaling pathways. Importantly, synergistic excitatory actions of GABAA and glutamergic NMDA receptor have been found in the neonatal hippocampus [40]. The interplay between GABA and glutamate-mediated excitatory actions is required for the induction of synaptic plasticity during the development [40,43]. Interestingly, NKCC1 in immature cortical neurons is stimulated by group-I mGluR- and glutamate inotropic receptors NMDA- and AMPA-mediated signal transduction pathways [21,22]. The effects are dependent on a rise of intracellular Ca2+. A synergistic interaction between GABAA and NMDA receptors has been reported in formation of synchronous Caþ2 oscillations [40]. Stimulation of NKCC1 by glutamate could increase intracellular Cl– that would reinforce the depolarizing actions of GABA [21]. In addition to neurons, GABA also triggers a depolarizing efflux of Cl– in cultured oligodendrocytes [44]. Re-accumulation of intracellular Cl– after depletion is blocked by removal of extracellular Naþ or inhibition of NKCC1 activity [45]. Thus, NKCC1 may be important in maintaining high [Cl–]i in oligodendrocytes. While the trophic effect of GABA on neurons is now well established [46,47], the effect of GABA on oligodendrocyte survival and growth remains uncertain. NKCC1 is present and active in rat spinal cord oligodendrocytes [35]. Expression of NKCC1 protein in the rat spinal cord is increased during development. In cultured oligodendrocytes, 39% of the total Kþ (86Rbþ) influx represents NKCC1 activity. Activation of GABAA receptors with muscimol produces a reduction in intracellular Cl– content and a stimulation of NKCC1 activity. Muscimol also triggers an increase in [Ca2+]i; this increase depends on NKCC1 activity. Survival of oligodendrocytes following withdrawal of growth factors is enhanced by muscimol; this effect also requires

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NKCC1 activity [35]). These results imply that NKCC1 functions in rat oligodendrocytes to maintain [Cl–]i above electrochemical equilibrium, and that it is required for GABAergic trophic effects. Both astrocytes and the choroid plexus epithelia have been suggested to play an important role in Kþ uptake in the CNS. Co-localization of NKCC1 and glial fibrillary acidic protein (astrocyte marker protein) was detected in astrocytes of cortex, corpus callosum, hippocampus, and cerebellum of rat [48]. Expression of NKCC1 was also observed in the perivascular astrocytes of cortical cortex, cerebral white matter, and hippocampus [48]. One of the major functions of astrocytes in the CNS is to buffer the extracellular Kþ concentration by transporting the Kþ released by active neurons [49]. Astrocytes take up Kþ, which can then leave astrocytes at remote locations where extracellular Kþ concentrations have not risen, or Kþ can efflux at the endfeet of the astrocyte where it is subsequently transported by blood vessels [49]. NKCC1 in the processes of the perivascular astrocytes is co-localized with AQP4, the latter is abundantly expressed in the astrocyte endfeet [50]. Therefore, these data suggest that NKCC1 may play a role in transport of Kþ from brain to blood to restore the extracellular Kþ concentration after neuron firing cessation. Lastly, the epithelial cells in choroid plexus are important for cerebrospinal fluid production. NKCC1 is constitutively active in choroid plexus epithelial cells and functions to reabsorb Kþ from the CSF, therefore, buffering and regulating brain interstitial Kþ homeostasis [33]. In addition to Kþ uptake, NKCC1 may also function in the regulation of intracellular Cl- in astrocytes. In acutely isolated hippocampal astrocytes from mature Sprague-Dawley rat, intracellular Cl- level is higher than it is predicted by passive distribution and the Cl--equilibrium potential is far more positive than the membrane potential [51]. Application of GABA in the CA3 region of the neuronfree rat hippocampal slice induced depolarization of astrocytes [52]. A similar GABAA receptor-induced depolarization has also been observed in acutely isolated astrocytes from adult Sprague-Dawley rat or cultured mature hippocampal astrocytes from neonatal Wistar rat [51,53]. These findings suggest that NKCC1 may play a role in maintaining high intracellular Cl- levels in astrocytes. NKCC1 activity is important in maintaining cell volume. Inhibition of NKCC1 activity or genetic ablation of NKCC1 impairs regulatory volume increase in astrocytes and neurons following hypertonicity-induced cell shrinkage [22,52])

8 Pathophysiological Relevance in Stroke 8.1 NKCC1 Contributes to Neuronal Damage During Acute Neurotoxicity Cl movement has been shown to be a central component of the acute excitotoxic response in neurons [54,55]. The acute excitotoxicity is thought to be

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mediated by excessive depolarization of the postsynaptic membrane. Depolarization results in an osmotic imbalance, which is countered by an influx of Cl-, Naþ, and water and leads to cell lysis. Removal or reduction of Cl– from the extracellular medium during excitatory amino acid (EAA) exposure completely eliminates the acute excitotoxic response in hippocampal [54] and retinal neurons [55]. NKCC1 is important in maintenance of [Cl]i in neurons as described above [15,37,39,56]. Thus, NKCC1 may affect neuronal excitability through regulation of [Cl-]i [15,57]. Activation of the ionotropic glutamate NMDA receptor, the AMPA receptor, and the metabotropic glutamate receptor (group-I) significantly stimulates NKCC1 activity in SH-SY5Y neuroblastoma cells [56]. Interestingly, inhibition of NKCC1 activity with its potent inhibitor bumetanide abolishes glutamate-mediated neurotoxicity and significantly attenuates oxygen and glucose deprivation (OGD)-induced neuronal death (Fig. 2, [58]). Furthermore, NMDA-mediated increases in 36Cl content and intracellular Naþ overload are reduced in the presence of bumetanide [58]. Bumetanide also blocks NMDA-induced neuronal swelling [58]. Thus, NKCC1 may act as another mechanism that contributes to Naþ and Cl– overload during glutamate-mediated acute excitotoxicity. This view is further supported by the

Fig. 2 Inhibition of NKCC1 decreases glutamate-mediated neurotoxicity. Cell mortality in cultured neurons was assessed by propidium iodide and calcein staining after 24-h treatment with glutamate or NMDA. A. Data are means SE. * P < 0.05 vs. control, ** P < 0.05 vs. NMDA, # P < 0.05 vs. glutamate. B. Bumetanide was added 30 min prior to the glutamate treatment and cells were assessed at 24 h post-treatment. * P < 0.05 vs. control, # P < 0.05 vs. glutamate (adapted from [58])

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similar levels of neuroprotection exhibited in neurons from NKCC1-null (NKCC1–/–) mice [3]. In addition, in neurons with injured axons, Cl– accumulation is attributable to NKCC1-mediated Cl– influx and a decrease in Cl– extrusion by KCC2 [59]. Cl– accumulation causes a switch in GABA action from inhibitory to excitatory and leads to excitotoxicity [59]. Blockage of Naþ and Cl– entry by removal of extracellular Naþ and Cl– abolishes excitotoxic dendritic injury and NMDA-mediated neurodegeneration in retinal ganglion cells [60,61]. Downregulation of KCC2 and a subsequent decrease in GABAmediated inhibition may contribute to excitotoxicity in injured neurons [62]. These results imply that stimulation of NKCC1 activity and downregulation of KCC2 are important in ischemic neuronal damage.

8.2 NKCC1-mediated Ionic Dysregulation, Swelling, and Excitatory Amino Acid Release in Astrocytes Following Ischemia Glial cells outnumber neurons by 10 times and occupy up to 50% of the human brain volume [63]. Glial cells have a critical function in the glutamate–glutamine shuttle, lactic acid supply for neuronal energy metabolism, regulation of synaptic and perisynaptic glutamate levels, regulation of ion concentrations (Hþ, Kþ, and Ca2+) in the intracellular and extracellular spaces, and the coupling of neuronal activity and cerebral blood flow [64, 65]. Therefore, alterations of astrocyte function after ischemia could have a significant impact on cerebral brain damage. Early loss of astrocytes is a principal feature of hyperglycemic and complete ischemia [66]. Despite the importance of astrocytes in neurodegeneration, the mechanisms underlying ischemic astrocyte damage are poorly understood. The following research findings illustrate the role of NKCC1 in ischemia-induced astrocyte damage. 8.2.1 High [Kþ]o-mediated Astrocyte Swelling Astrocytes undergo rapid swelling in a number of acute pathological states, such as ischemia and traumatic brain injury [67]. Unresolved astrocyte swelling will have detrimental effects such as reduction of extracellular space, a decrease of normal cerebral blood flow, and an accumulation of EAAs such as glutamate [68]. In addition, swollen astrocytes would have a diminished capacity to perform their normal homeostatic functions. Considerable work has been focused on the role of cation-chloride cotransporters in maintenance and regulation of astrocyte volume. The expression of NKCC1 protein is found in astrocytes [48], which could function in clearing of excessive [Kþ]o as mentioned above [69]. During ischemia, one significant pathophysiological change in the CNS is an

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elevation of [Kþ]o. A few minutes of anoxia/ischemia raises [Kþ]o to 60 mM [8]. NKCC1 in astrocytes may play a role in Kþ uptake under high [Kþ]o conditions. In cultured astrocytes, 75 mM [Kþ]o causes NKCC1-mediated Kþ influx to be stimulated by 80% [70]. The high [Kþ]o-induced activation of NKCC1 is completely abolished by either removal of extracellular Ca2+ or by blocking the L-type voltage-dependent Ca2+ channels with nifedipine [71]. Additionally, intracellular 36Cl accumulation increases significantly in response to 75 mM [Kþ]o and is abolished by inhibition of NKCC1 activity either by bumetanide or genetic ablation [72,71]. These data suggest that the cotransporter activity is stimulated under high [Kþ]o via Ca2+-mediated signal transduction pathways. Seventy-five millimolar [Kþ]o also triggers cell swelling in NKCC1þ/þ astrocytes by 20% [70]. This high [Kþ]o-mediated swelling is abolished by inhibition of NKCC1 activity, either with bumetanide or genetic ablation [72,70]. In addition, high [Kþ]o-induced astrocyte swelling is also observed in the rat optic nerve model [73]. Furosemide and bumetanide reversibly suppress the high [Kþ]o-induced astrocyte swelling in enucleated nerves [73]. Additionally, astrocytes in optic nerves exhibited a Ca2+-independent swelling during ischemia that was blocked by bumetanide [24]. Taken together, these studies suggest that NKCC1 activation leads to astrocyte swelling. 8.2.2 High [Kþ]o-Mediated Excitatory Amino Acid Release from Astrocytes Glutamate is the principal excitatory neurotransmitter in the mammalian CNS. Elevation of glutamate in the extracellular space and excessive activation of NMDA receptors cause cell death [75]. Astrocytes maintain a large transmembrane glutamate gradient, with [Glu]i of 2–10 mM whereas [Glu]o is approximately 1 mM [76]. This gradient results from the high density of Naþ-dependent glutamate transporters in astrocytes, which regulate synaptic transmission by controlling glutamate diffusion and concentration in the extracellular space [77]. Therefore, reduced glutamate uptake or release of sequestered glutamate from astrocytes may contribute to excitotoxicity under pathophysiological conditions [67]. For example, one consequence of high [Kþ]o-induced astrocyte swelling will be the release of glutamate via activation of volume-sensitive organic anion channels (VSOACs) [71,78]. VSOACs are ubiquitously expressed chloride channels, which are permeable to a variety of small organic anions, including the amino acids taurine, glutamate, and aspartate [79]. In response to swelling, VSOAC-mediated efflux of amino acids can function as a regulatory volume decrease mechanism in astrocytes [82]. Release of [14C]-D-aspartate from astrocytes, which serves as a surrogate for glutamate, is negligible under control conditions [70,80]. However, high [Kþ]o triggers a significant release of [14C]-D-aspartate from astrocytes. High [Kþ]o-induced release of [14C]-Daspartate is abolished by the anion channel blocker DIDS, suggesting an involvement of VSOACs [53,69]. When NKCC1 is inhibited by bumetanide or genetic ablation, this high [Kþ]o-induced aspartate release is reduced by

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30–50% (Fig. 3) [72,70]. On the other hand, stimulation of NKCC1 under high [Kþ]o conditions leads to increases in intracellular Naþ, which could slow or even reverse Naþ-dependent glutamate transporters and increase glutamate release from astrocytes. Taken together, astrocyte NKCC1 stimulation will contribute to excitotoxic injury by increasing swellinginduced glutamate release or decreasing glutamate clearance from the extracellular space.

Fig. 3 Effect of bumetanide on the high-[Kþ]o-mediated release of preloaded D-[14C]aspartate from astrocytes. A. Cells loaded with D-[14C]aspartate were perfused with 75 mM [Kþ]o for 20 min. B. Astrocytes were exposed to bumetanide for 20 min and then exposed to 75 mM [Kþ]o + bumetanide. Cells were then washed with normal buffer and re-exposed to 75 mM [Kþ]o. (adapted from [72,71])

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8.2.3 Naþ and Cl– Dysregulation in Astrocytes Following Oxygen and Glucose Deprivation Loss of the trans-plasma membrane Naþ gradient is one of the key elements in promoting cellular damage in astrocytes during ischemia [81]. An increase in intracellular Naþ is found in rat spinal cord astrocytes [82], rat cortical astrocytes [81], and mouse cortical astrocytes [83] under ischemic conditions. While decreased activity of Naþ/Kþ-ATPase during ischemia will lead to reduced Naþ extrusion, the activation of several Naþ entry pathways, including NKCC1, also contributes to ischemia-induced loss of the Naþ homeostasis. OGD in astrocytes results in both an increase in NKCC1 phosphorylation and 86Rb influx [84]. This stimulation of NKCC1 activity is accompanied by an accumulation of 36Cl and 3.5-fold increase in [Naþ]i in astrocytes [85]. Either bumetanide or genetic ablation of NKCC1 significantly reduces the OGD/REOXmediated increase in [Naþ]i and [Cl–]i. Given that the extracellular ionic composition during hypoxia/ischemia is featured with a lower concentration of Naþ, Cl–, Ca2+ and a higher concentration of Kþ as well as Hþ [86], it is important to examine whether NKCC1 still functions in this altered extracellular ionic environment. Thus, an alternative in vitro hypoxic model was established by perfusing cells with hypoxic, acidic, ion shifted-Ringers (HAIR) that mimics the extracellular ionic and oxygen composition in vivo. Following 5 min hypoxia, [Naþ]i raises rapidly over the first 5 min REOX and plateaus to a value approximately four-fold of control and is sustained over the following 45 min REOX. Bumetanide or genetic ablation of NKCC1 reduces hypoxia-mediated rise in [Naþ]i by 65% [87]. The HAIRmediated increases in [Naþ]i are doubled when Naþ/Kþ-ATPase activity is blocked with ouabain [87]. This suggests that the NKCC1 mediates a Naþ overload that cannot be offset by Naþ/Kþ-ATPase activity. 8.2.4 Ca2+ Dysregulation in Astrocytes Following Oxygen and Glucose Deprivation NKCC1-mediated increases in [Naþ]i following in vitro ischemia have been linked to Ca2+ loading and subsequent mitochondrial damage. As a consequence of increases in [Naþ]i, Naþ/Ca2+ exchanger, (NCX)-mediated Ca2+ extrusion is reduced or NCX may function in a reverse mode (NCXrev), which leads to increases in [Caþ2]i (Fig. 4). Thus, NKCC1 and NCXrev may contribute to intracellular Naþ and Caþ2 overload in astrocytes following in vitro ischemia [85]. In addition, Naþ and Caþ2 overload in ischemic astrocytes leads to a dissipation of the mitochondrial membrane potential and release of mitochondrial cytochrome c, which can be attenuated by bumetanide or with genetic ablation of NKCC1 [87]. These findings suggest that the concerted activities of multiple ion transport proteins are important in the perturbations of Naþ and Ca2+ homeostasis and in astrocyte damage in response to hypoxia/ischemia.

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Fig. 4 Increase in [Ca2þ]i following HAIR/REOX depends on NKCC1 and NCX activity. A. [Ca2+]i was monitored in NKCC1þ/ + or NKCC1–/– astrocytes during 15 min HAIR and 45 min REOX. In studies involving KB-R7943 an inhibitor of NCXrev, the drug was present 30 min before HAIR and during REOX. B. Summary data of changes in [Ca2+]i at 45 min REOX. Data are means SE. * P < 0.05 vs. control, # P < 0.05 vs. NKCC1þ/+ HAIR/REOX. (adapted from [87])

8.3 NKCC1 in AMPA-Mediated Oligodendrocyte Excitotoxicity The primary role of oligodendrocytes in the CNS is the production of myelin, which forms an insulating sheath around the axons of neurons and allows for efficient electrical transmission [88]. Loss of oligodendrocyte function is involved in a number of neurodegeneration diseases. Likewise, early ischemic damage to white matter involves oligodendrocyte and axon damage [89]. NKCC1 plays a role in oligodendrocyte damage following activation of nonNMDA glutamate receptors. AMPA/kainate receptor-mediated excitotoxcity results in spinal cord white matter injury in a process that is linked to oligodendrocyte damage and axonal demyelination [90]. Indeed, the vulnerability of immature oligodendrocytes to ischemic injury is a major component of the brain injury associated with cerebral palsy [91]. Oligodendrocytes are thought to express primarily non-NMDA type ionotropic glutamate receptors, although recent studies suggest that NMDA-type glutamate receptors may predominate in processes, while AMPA/kainate receptors predominate on the soma [92]. Exposing cultured oligodendrocytes to AMPA plus the AMPA desensitization blocker cyclothiazide (CTZ) leads to a transient rise in [Ca2+]i, which is followed by a sustained intracellular [Ca2+]i overload, NKCC1 phosphorylation, and a NKCC1-mediated Naþ influx [93]. In the presence of a specific AMPA receptor inhibitor 6-cyano-7-nitroquinoxaline-2, 3-dione (CNQX), AMPA/CTZ fails to elicit any changes in [Ca2+]i. The AMPA/CTZ-induced sustained [Ca2+]i rise led to mitochondrial Caþ2 accumulation, release of cytochrome c from mitochondria, and cell death [93]. The AMPA/CTZ-elicited [Ca2+]i increase, mitochondrial damage, and cell death are significantly reduced by inhibiting NKCC1 or NCXrev. These data show that in cultured

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oligodendrocytes, activation of AMPA receptors leads to NKCC1 phosphorylation that enhances NKCC1-mediated Naþ influx. The latter triggers NCXrev-mediated [Ca2+]i overload and compromises mitochondrial function and cellular viability [93]. Such a link between Naþ overload and activation of NCXrev has been observed under other conditions [94]. NCXrev causes Ca2+ influx after NMDA and non-NMDA receptor activation in depolarized and glucose-deprived neurons [95,96]. Increase in intracellular Naþ and the subsequent induction of NCXrev have also been found in mechanical strain injury of astrocytes [97] and in axonal damage [94].

8.4 NKCC1 in Ischemic Hippocampal Damage Several studies have indicated that OGD in the brain slice induces an accumulation of [Cl–]i in area CA1 hippocampal neurons [98–101]. [Cl–]i started to increase during OGD and accumulated throughout REOX, reaching levels twice as high as those found immediately after OGD [98]. In another study, Cl– content in the cytoplasm, mitochondria, and nuclei was increased by 50% after 10 min OGD and by 300% at 30 min REOX following 10 min OGD [99]. The sustained rise in [Cl–]i following OGD can directly induce neuronal damage [100]. Given the apparent significance of the OGD-mediated rise in [Cl–]i, considerable effort has focused on the underlying mechanisms. Recent studies indicate that the rise in [Cl–]i during OGD is likely to be due a number of different Cl– transport mechanisms, including Cl– transporters and Cl– channels [101]. In contrast, the rise in [Cl–]i during REOX can be completely prevented by bumetanide [101]. Increased phosphorylation of NKCC1 has been demonstrated after OGD, consistent with the timing of changes of [Cl–]i [101]. NKCC1 activity may be primarily responsible for the changes in [Cl–]i during REOX.

8.5 NKCC1 in In Vivo Cerebral Ischemia While in vitro studies are useful for demonstrating proof of principle for a role of NKCC1 in cerebral ischemia, it is important to extend these studies by using in vivo animal models. The most widely used model of focal cerebral ischemia is the endovascular suture occlusion of the middle cerebral artery (MCAO) in rodents. Expression of NKCC1 detected with T4 antibody immunoreactive signals is observed in cell bodies and dendrites of pyramidal neurons in cerebral cortex of sham rats. Two hours of focal cerebral ischemia and 24 h reperfusion lead to increased NKCC1 immunostaining in neurons scattered in the ischemic cortex and striatum [34]. Immunoblotting revealed that expression of NKCC1 protein was increased following 2 h focal ischemia in cerebral cortex and striatum. A sustained upregulation of NKCC1 in cortex was detected at 4, 8,

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12, and 24 h of reperfusion [34]. An increase in the phosphorylated NKCC1 (NKCC1-p) was also found at 4 and 8 h of reperfusion in the ipsilateral cortex [34]. Because the change in phosphorylation state of NKCC1 is a major regulatory mechanism that stimulates NKCC1 activity, these findings suggest that elevated NKCC1 expression and activity may be involved in cerebral ischemic damage. Upregulation of NKCC1 is found in hippocampus in gerbils after 5 min global ischemia and 24 h reperfusion [102]. However, in the same model there is also a transient downregulation of NKCC1 protein expression at 12 h reperfusion [102], which may be a compensatory reaction to protect neurons from excitotoxicity. Two hours of MCAO and 24 h reperfusion result in an average infarction volume of 238.3 mm3 in spontaneous hypertensive rats [34]. However, when 100 mM bumetanide is continuously infused via microdialysis into the cortex during MCAO and reperfusion, the resulting infarct was reduced by 25%. Edema induced by MCAO was also reduced by 79% in the bumetanide-treated brains [34]. This indicates that NKCC1 contributes to cerebral ischemic cell damage and edema in vivo. In rats subjected to permanent MCAO, intravenous administration of bumetanide (7.6–30.4 mg/kg) immediately before occlusion attenuates edema formation as determined by magnetic resonance imaging [103]. This further suggests a role for NKCC1 in the edema formation during cerebral ischemia. In addition to pharmacological blockage of NKCC1, genetic ablation of NKCC1 can be used to further establish that NKCC1 contributes to ischemic damage. Two hours MCAO and 10 or 24 h reperfusion caused infarction (85 mm3) in NKCC1 wild-type (NKCC1þ/+) mice (Fig. 5). Infarction volume in NKCC1–/– mice was reduced by 30%–46% [3]. Heterozygous mutant (NKCC1þ/–) mice also showed 28% reduction in infarction [3]. Brain edema was significantly increased after 2 h MCAO and 24 h reperfusion in NKCC1þ/+ mice. In contrast, there was 50% less edema formation in either NKCC1þ/– or NKCC1–/– mice [3]. These results are consistent with the neuroprotective effects in rats mediated by pharmacological inhibition of NKCC1 with bumetanide. White matter ion homeostasis, especially Naþ and Kþ, is crucial to normal axonal functions and dissipation of ionic homeostasis will lead to irreversible injury. Amyloid precursor protein (APP) is transported by fast anterograde axon transport along microtubules and has recently been shown as a sensitive marker of axonal disruption in white matter during brain ischemia [104]. Disruption of this transport as a consequence of cytoskeleton derangement results in APP accumulation, indicating the dysfunction or discontinuity of the axons [105]. Two to 4 h focal ischemia causes cytoskeletal breakdown and disturbance of fast axonal transport in myelinated fiber tracts [105]. NKCC1þ/+ brains exhibit significant accumulation of APP in the ipsilateral internal capsule after 2 h focal ischemia or 10 h reperfusion. In contrast, APP accumulation is reduced by 55% in NKCC1–/– brains after 2 h focal ischemia and 10 h reperfusion [3]. White matter degeneration

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Fig. 5 Brain infarction is decreased in NKCC1–/– mice. A. Coronal sections of brains from NKCC1þ/+- or NKCC1–/– mice after 2 h MCAO and 10 or 24 h reperfusion were stained with TCC and infarct volume was calculated. B. Summarized infarct volume. Data are means SD, P < 0.05 vs. NKCC1þ/ + . Scale bar 5 mm. (adapted from [3])

is often assumed to be secondary to neuronal damage. However, no changes in myelin basic protein expression were observed after 2 h MCAO and 24 h reperfusion [3]. This implies that APP accumulation occurs early and is likely a result of primary axonal ischemic damage. Taken together, these results suggest that NKCC1 plays a role not only in gray matter but also in white matter ischemic damage. The in vitro studies in astrocytes or in oligodendrocytes show that NKCC1 activation leads to Naþ overload, which triggers NCXrev after ischemia and excitotoxicity [85,87,93]. Although these findings support that NKCC1 and NCXrev may collectively contribute to ischemic cell damage, it has not yet been tested in in vivo ischemic models until recently. Luo et al [106] examined whether reduction of NKCC1 and NCX1 could provide neuroprotection after in vitro or in vivo ischemia. Both neurons and astrocytes cultured from double heterozygous (NKCC1þ/–/NCX1þ/–) mice showed considerably reduced expression and function of NKCC1 and NCX1 [106]. In addition, as compared to NCX1þ/+ mice, infarct volume at either 24 or 72 h reperfusion following 30 min MCAO is reduced by 50% in NKCC1þ/–/NCX1þ/– mice [106]. Taken together, these studies demonstrate that NKCC1 plays a role in ischemic brain damage.

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9 Therapeutic Perspectives NKCC1 has a broad tissue distribution and serves to maintain cellular ion homeostasis. Over the past decades the function and regulation of NKCC1 have been extensively focused in epithelial cells. The study of the role of NKCC1 in the CNS is relatively new and exciting. NKCC1 protein is expressed in multiple cell types throughout the brain. Through the regulation of intracellular Cl–, NKCC1 activity is able to affect GABAergic signaling and neuronal excitability. Therefore, NKCC1 is being explored as a target for anti-convulsant therapy in neonates [107]. Preclinical in vitro and in vivo experimental studies also demonstrate that NKCC1 activation contributes to ischemic neuronal damage. Thus, bumetanide could be useful for reducing ischemic damage, edema, and seizures that are secondary to ischemic brain injury. Controls of bumetanide’s action in the periphery system to limit systemic side effects in kidney and electrolyte homeostasis should be considered in the therapy development.

10 Conclusion Loss of ion homeostasis plays a central role in pathogenesis of ischemic cell damage. Ischemia-induced perturbation of ion homeostasis leads to intracellular accumulation of Naþ and Ca2+ and subsequent activation of proteases, phospholipases, and formation of oxygen and nitrogen free radicals. This cascade of signal transduction events results in long-term functional and structural changes in membrane and cytoskeletal integrity and eventual cell death. Secondary active ion transport proteins including NKCC1 are important in maintaining steady-state intracellular ionic concentrations. Considerable research effort has been centered on roles of passive fluxes via cation and anion conductances in dissipation of the ion concentration gradients. Results from recent in vitro and in vivo experimental studies demonstrate the role of NKCC1 in ischemia-induced dissipation of ionic homeostasis. Therefore, the ion transport protein could be a target for the future development of therapeutic treatment of ischemic cerebral damage. Acknowledgments This work was supported in part by an NIH grant (R01NS38118). The author thanks Douglas B. Kintner for his assistance in preparation of the manuscript

Abbreviations APP CNS EAA GABA

Amyloid precursor protein Central nervous system Excitatory amino acid g-Aminobutyric acid

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[Cl–]i [Glu]i [Glu]o [Kþ]o hNKCC1 mNKCC1 MCAO NCX NCXrev NCC NKCC NKCC1 NKCC2 NKCC1þ/+ mice NKCC1þ/– mice NKCC1–/– mice OSR1 OGD PASK REOX sNKCC1 SPAK TM VSOACs WNK family of serine-threonine kinases

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Intracellular Cl– concentration Intracellular glutamate concentration Extracellular glutamate concentration Extracellular Kþ concentration Human NKCC1; KCC2, K-Cl cotransporter isoform 2 Mouse Na-K-Cl cotransporter isoform 1 Middle cerebral artery occlusion Naþ/Ca2+ exchanger Reverse mode operation of NCX Naþ-Cl– cotransporter Naþ-Kþ-Cl– cotransporter NKCC isoform 1 NKCC isoform 2 NKCC1 wild-type mice NKCC1 heterozygous mutant mice NKCC1 null mice Oxidative stress response kinase Oxygen and glucose deprivation Proline-alanine-rich STE20-related kinase Reoxygenation Shark rectal gland Na-K-Cl cotransporter STE20/SPS-1 related, Proline-alanine-rich kinase transmembrane domains Volume-sensitive organic anion channels kinases with no K = lysine, which is substituted with cysteine.

References 1. Haas M and Forbush B III. The Na-K-Cl cotransporter of secretory epithelia. Annu Rev Physiol 62: 515–534, 2000. 2. Russell JM. Sodium-potassium-chloride cotransport. Physiol Rev 80: 211–276, 2000. 3. Chen H, Luo J, Kintner DB, Shull GE, and Sun D. Na+-dependent chloride transporter (NKCC1)-null mice exhibit less gray and white matter damage after focal cerebral ischemia. J Cereb Blood Flow Metab 25: 54–66, 2005. 4. Xu JC, Lytle C, Zhu TT, Payne JA, Benz E Jr., and Forbush B III. Molecular cloning and functional expression of the bumetanide-sensitive Na-K-Cl cotransporter. Proc Natl Acad Sci USA 91: 2201–2205, 1994. 5. Delpire E and Mount DB. Human and murine phenotypes associated with defects in cation-chloride cotransport. Annu Rev Physiol 64: 803–843, 2002. 6. Flagella M, Clarke LL, Miller ML, Erway LC, Giannella RA, Andringa A, et al. Mice lacking the basolateral Na-K-2Cl cotransporter have impaired epithelial chloride secretion and are profoundly deaf. J Biol Chem 274: 26946–26955, 1999.

The ‘‘Loop’’ Diuretic Drug Bumetanide-Sensitive Naþ-Kþ-Cl– Cotransporter

107

7. Meyer JW, Flagella M, Sutliff RL, Lorenz JN, Nieman ML, Weber CS, et al. Decreased blood pressure and vascular smooth muscle tone in mice lacking basolateral Na+-K+2Cl– cotransporter. Am J Physiol Heart Circ Physiol 283: H1846–H1855, 2002. 8. Siesjo BK. Pathophysiology and treatment of focal cerebral ischemia. Part I: Pathophysiology. J Neurosurg 77: 169–184, 1992. 9. Gamba G. Molecular physiology and pathophysiology of electroneutral cation-chloride cotransporters. Physiol Rev 85: 423–493, 2005. 10. Payne JA, Xu JC, Haas M, Lytle CY, Ward D, and Forbush B III. Primary structure, functional expression, and chromosomal localization of the bumetanide-sensitive Na-KCl cotransporter in human colon. J Biol Chem 270: 17977–17985, 1995. 11. Gimenez I. Molecular mechanisms and regulation of furosemide-sensitive Na-K-Cl cotransporters. Curr Opin Nephrol Hypertens 15: 517–523, 2006. 12. Gerelsaikhan T and Turner RJ. Transmembrane topology of the secretory Na+-K+-2Cl– cotransporter NKCC1 studied by in vitro translation. J Biol Chem 275: 40471–40477, 2000. 13. Isenring P, Jacoby SC, Chang J, and Forbush B. Mutagenic mapping of the Na-K-Cl cotransporter for domains involved in ion transport and bumetanide binding. J Gen Physiol 112: 549–558, 1998. 14. Isenring P and Forbush B III. Ion and bumetanide binding by the Na-K-Cl cotransporter. Importance of transmembrane domains. J Biol Chem 272: 24556–24562, 1997. 15. Sung KW, Kirby M, McDonald MP, Lovinger DM, and Delpire E. Abnormal GABAA receptor-mediated currents in dorsal root ganglion neurons isolated from Na-K-2Cl cotransporter null mice. J Neuropsychiatry 20: 7531–7538, 2000. 16. Kahle KT and Staley K. Altered neuronal chloride homeostasis and excitatory GABAergic signaling in human temporal lobe epilepsy. Epilepsy Curr 8: 51–53, 2008. 17. Forbush B III and Palfrey HC. [3H]bumetanide binding to membranes isolated from dog kidney outer medulla. Relationship to the Na, K, Cl co-transport system. J Biol Chem 258: 11787–11792, 1983. 18. O’Grady SM, Palfrey HC, and Field M. Characteristics and functions of Na-K-Cl cotransport in epithelial tissues. Am J Physiol 253: C177–C192, 1987. 19. Darman RB and Forbush B. A regulatory locus of phosphorylation in the N terminus of the Na-K-Cl cotransporter, NKCC1. J Biol Chem 277: 37542–37550, 2002. 20. Kurihara K, Moore-Hoon ML, Saitoh M, and Turner RJ. Characterization of a phosphorylation event resulting in upregulation of the salivary Na(+)-K(+)-2Cl(–) cotransporter. Am J Physiol 277: C1184–C1193, 1999. 21. Schomberg SL, Su G, Haworth RA, and Sun D. Stimulation of Na-K-2Cl cotransporter in neurons by activation of Non-NMDA ionotropic receptor and group-I mGluRs. J Neurophysiol 85: 2563–2575, 2001. 22. Schomberg SL, Bauer J, Kintner DB, Su G, Flemmer A, Forbush B, et al. Cross talk between the GABAA receptor and the Na-K-Cl cotransporter is mediated by intracellular Cl–. J Neurophysiol 89: 159–167, 2003. 23. Lytle C and Forbush B III. Regulatory phosphorylation of the secretory Na-K-Cl cotransporter: modulation by cytoplasmic Cl. Am J Physiol 270: C437–C448, 1996. 24. Haas M, McBrayer D, and Lytle C. [Cl–]i-dependent phosphorylation of the Na-K-Cl cotransport protein of dog tracheal epithelial cells. J Biol Chem 270: 28955–28961, 1995. 25. Dowd BF and Forbush B. PASK (proline-alanine-rich STE20-related kinase), a regulatory kinase of the Na-K-Cl cotransporter (NKCC1). J Biol Chem 278: 27347–27353, 2003. 26. Piechotta K, Lu J, and Delpire E. Cation chloride cotransporters interact with the stressrelated kinases Ste20-related proline-alanine-rich kinase (SPAK) and oxidative stress response 1 (OSR1). J Biol Chem 277: 50812–50819, 2002. 27. Ushiro H, Tsutsumi T, Suzuki K, Kayahara T, and Nakano K. Molecular cloning and characterization of a novel Ste20-related protein kinase enriched in neurons and transporting epithelia. Arch Biochem Biophys 355: 233–240, 1998.

108

D. Sun

28. Johnston AM, Naselli G, Gonez LJ, Martin RM, Harrison LC, and DeAizpurua HJ. SPAK, a STE20/SPS1-related kinase that activates the p38 pathway. Oncogene 19: 4290–4297, 2000. 29. Piechotta K, Garbarini NJ, England R, and Delpire E. Characterization of the interaction of the stress kinase SPAK with the Na+-K+-2Cl– cotransporter in the nervous system: Evidence for a scaffolding role of the kinase. J Biol Chem 278: 52848–52856, 2003. 30. Darman RB, Flemmer A, and Forbush B. Modulation of ion transport by direct targeting of protein phosphatase type 1 to the Na-K-Cl cotransporter. J Biol Chem 276: 34359–34362, 2001. 31. Kahle KT, Rinehart J, de Los HP, Louvi A, Meade P, Vazquez N, et al. WNK3 modulates transport of Cl– in and out of cells: implications for control of cell volume and neuronal excitability. Proc Natl Acad Sci USA 102: 16783–16788, 2005. 32. Sun D, Lytle C, and O’Donnell ME. Astroglial cell-induced expression of Na-K-Cl cotransporter in brain microvascular endothelial cells. Am J Physiol 269: C1506–C1512, 1995. 33. Wu Q, Delpire E, Hebert SC, and Strange K. Functional demonstration of Na+-K+-2Cl– cotransporter activity in isolated, polarized choroid plexus cells. Am J Physiol 275: C1565–C1572, 1998. 34. Yan YP, Dempsey RJ, and Sun D. Na+-K+-Cl– cotransporter in rat focal cerebral ischemia. J Cereb Blood Flow Metab 21: 711–721, 2001. 35. Wang H, Yan Y, Kintner DB, Lytle C, and Sun D. GABA-mediated trophic effect on oligodendrocytes requires Na-K-2Cl cotransport activity. J Neurophysiol 90: 1257–1265, 2003. 36. Walz W. Role of Na/K/Cl cotransport in astrocytes. Can J Physiol Pharmacol 70 Suppl: S260–S262, 1992. 37. Misgeld U, Deisz RA, Dodt HU, and Lux HD. The role of chloride transport in postsynaptic inhibition of hippocampal neurons. Science 232: 1413–1415, 1986. 38. Cherubini E, Gaiarsa JL, and Ben Ari Y. GABA: an excitatory transmitter in early postnatal life. Trends Neurosci 14: 515–519, 1991. 39. Alvarez-Leefmans FJ. Intracellular chloride regulation. In: Sperelakis N (ed.) Cell Physiology Sourcebook: A Molecular Approach. San Diego: Academic Press, 301–318, 2001. 40. Ben Ari Y, Khazipov R, Leinekugel X, Caillard O, and Gaiarsa JL. GABAA, NMDA and AMPA receptors: a developmentally regulated ’menage a trois’. Trends Neurosci 20: 523–529, 1997. 41. Payne JA, Rivera C, Voipio J, and Kaila K. Cation-chloride co-transporters in neuronal communication, development and trauma. Trends Neurosci 26: 199–206, 2003. 42. Hubner CA, Stein V, Hermans-Borgmeyer I, Meyer T, Ballanyi K, and Jentsch TJ. Disruption of KCC2 reveals an essential role of K-Cl cotransport already in early synaptic inhibition. Neuron 30: 515–524, 2001. 43. Belhage B, Hansen GH, Elster L, and Schousboe A. Effects of gamma-aminobutyric acid (GABA) on synaptogenesis and synaptic function. Perspect Dev Neurobiol 5: 235–246, 1998. 44. Kettenmann H, Blankenfeld GV, and Trotter J. Physiological properties of oligodendrocytes during development. Ann N Y Acad Sci 633: 64–77, 1991. 45. Hoppe D and Kettenmann H. GABA triggers a Cl– efflux from cultured mouse oligodendrocytes. Neurosci Lett 97: 334–339, 1989. 46. Marty S, Berninger B, Carroll P, and Thoenen H. GABAergic stimulation regulates the phenotype of hippocampal interneurons through the regulation of brain-derived neurotrophic factor. Neuron 16: 565–570, 1996. 47. Shulga A, Thomas-Crusells J, Sigl T, Blaesse A, Mestres P, Meyer M, et al. Posttraumatic GABA(A)-mediated [Ca2+]i increase is essential for the induction of brain-derived neurotrophic factor-dependent survival of mature central neurons. J Neuropsychiatry 28: 6996–7005, 2008.

The ‘‘Loop’’ Diuretic Drug Bumetanide-Sensitive Naþ-Kþ-Cl– Cotransporter

109

48. Yan YP, Dempsey RJ, and Sun D. Expression of Na+-K+-Cl– cotransporter in rat brain during development and its localization in mature astrocytes. Brain Res 911: 43–55, 2001. 49. Paulson OB and Newman EA. Does the release of potassium from astrocyte endfeet regulate cerebral blood flow? Science 237: 896–898, 1987. 50. Nielsen S, Nagelhus EA, Amiry-Moghaddam M, Bourque C, Agre P, and Ottersen OP. Specialized membrane domains for water transport in glial cells: high-resolution immunogold cytochemistry of aquaporin-4 in rat brain. J Neuropsychiatry 17: 171–180, 1997. 51. Fraser DD, Duffy S, Angelides KJ, Perez-Velazquez JL, Kettenmann H, and MacVicar BA. GABAA/benzodiazepine receptors in acutely isolated hippocampal astrocytes. J Neuropsychiatry 15: 2720–2732, 1995. 52. MacVicar BA, Tse FW, Crichton SA, and Kettenmann H. GABA-activated Cl– channels in astrocytes of hippocampal slices. J Neuropsychiatry 9: 3577–3583, 1989. 53. Bekar LK and Walz W. Evidence for chloride ions as intracellular messenger substances in astrocytes. J Neurophysiol 82: 248–254, 1999. 54. Rothman SM. The neurotoxicity of excitatory amino acids is produced by passive chloride influx. J Neuropsychiatry 5: 1483–1489, 1985. 55. Olney JW, Price MT, Samson L, and Labruyere J. The role of specific ions in glutamate neurotoxicity. Neurosci Lett 65: 65–71, 1986. 56. Sun D and Murali SG. Stimulation of Na+-K+-2Cl– cotransporter in neuronal cells by excitatory neurotransmitter glutamate. Am J Physiol Cell Physiol 275: C772–C779, 1998. 57. Jang IS, Jeong HJ, and Akaike N. Contribution of the Na-K-Cl cotransporter on GABA(A) receptor-mediated presynaptic depolarization in excitatory nerve terminals. J Neuropsychiatry 21: 5962–5972, 2001. 58. Beck J, Lenart B, Kintner DB, and Sun D. Na-K-Cl cotransporter contributes to glutamate-mediated excitotoxicity. J Neuropsychiatry 23: 5061–5068, 2003. 59. Nabekura J, Ueno T, Okabe A, Furuta A, Iwaki T, Shimizu-Okabe C, et al. Reduction of KCC2 expression and GABAA receptor-mediated excitation after in vivo axonal injury. J Neuropsychiatry 22: 4412–4417, 2002. 60. Hasbani MJ, Hyrc KL, Faddis BT, Romano C, and Goldberg MP. Distinct roles for sodium, chloride, and calcium in excitotoxic dendritic injury and recovery. Exp Neurol 154: 241–258, 1998. 61. Chen Q, Olney JW, Lukasiewicz PD, Almli T, and Romano C. Ca2+-independent excitotoxic neurodegeneration in isolated retina, an intact neural net: a role for Cl– and inhibitory transmitters. Mol Pharmacol 53: 564–572, 1998. 62. Toyoda H, Ohno K, Yamada J, Ikeda M, Okabe A, Sato K, et al. Induction of NMDA and GABAA receptor-mediated Ca2+ oscillations with KCC2 mRNA downregulation in injured facial motoneurons. J Neurophysiol 89: 1353–1362, 2003. 63. Tower DB and Young OM. The activities of butyrylcholinesterase and carbonic anhydrase, the rate of anaerobic glycolysis, and the question of a constant density of glial cells in cerebral cortices of various mammalian species from mouse to whale. J Neurochem 20: 269–278, 1973. 64. Chen Y and Swanson RA. Astrocytes and brain injury. J Cereb Blood Flow Metab 23: 137–149, 2003. 65. Schurr A. Lactate: the ultimate cerebral oxidative energy substrate? J. Cereb Blood Flow Metab 26: 142–152, 2006. 66. Pulsinelli WA, Brierley JB, and Plum F. Temporal profile of neuronal damage in a model of transient forebrain ischemia. Ann Neurol 11: 491–498, 1982. 67. Kimelberg HK. Astrocytic swelling in cerebral ischemia as a possible cause of injury and target for therapy. Glia 50: 389–397, 2005. 68. Rutledge EM and Kimelberg HK. Release of [3H]-D-aspartate from primary astrocyte cultures in response to raised external potassium. J Neuropsychiatry 16: 7803–7811, 1996. 69. Walz W. Role of astrocytes in the clearance of excess extracellular potassium. Neurochem Int 36: 291–300, 2000.

110

D. Sun

70. Su G, Kintner DB, and Sun D. Contribution of Na(+)-K(+)-Cl(–) cotransporter to high-[K(+)](o)- induced swelling and EAA release in astrocytes. Am J Physiol Cell Physiol 282: C1136–C1146, 2002. 71. Su G, Haworth RA, Dempsey RJ, and Sun D. Regulation of Na+-K+-Cl– cotransporter in primary astrocytes by dibutyryl cAMP and high [K+]o. Am J Physiol Cell Physiol 279: C1710–C1721, 2000. 72. Su G, Kintner DB, Flagella M, Shull GE, and Sun D. Astrocytes from Na(+)-K(+)Cl(–) cotransporter-null mice exhibit absence of swelling and decrease in EAA release. Am J Physiol Cell Physiol 282: C1147–C1160, 2002. 73. MacVicar BA, Feighan D, Brown A, and Ransom B. Intrinsic optical signals in the rat optic nerve: role for K(+) uptake via NKCC1 and swelling of astrocytes. Glia 37: 114–123, 2002. 74. Thomas R, Salter MG, Wilke S, Husen A, Allcock N, Nivison M, et al. Acute ischemic injury of astrocytes is mediated by Na-K-Cl cotransport and not Ca2+ influx at a key point in white matter development. J Neuropathol Exp Neurol 63: 856–871, 2004. 75. Olney JW. Brain lesions, obesity, and other disturbances in mice treated with monosodium glutamate. Science 164: 719–721, 1969. 76. Erecinska M and Silver IA. Metabolism and role of glutamate in mammalian brain. Prog Neurobiol 35: 245–296, 1990. 77. Oliet SH, Piet R, and Poulain DA. Control of glutamate clearance and synaptic efficacy by glial coverage of neurons. Science 292: 923–926, 2001. 78. Abdullaev IF, Rudkouskaya A, Schools GP, Kimelberg HK, and Mongin AA. Pharmacological comparison of swelling-activated excitatory amino acid release and Cl-currents in cultured rat astrocytes. J Physiol 572: 677–689, 2006. 79. Strange K, Emma F, and Jackson PS. Cellular and molecular physiology of volumesensitive anion channels. Am J Physiol 270: C711–C730, 1996. 80. Rutledge EM, Aschner M, and Kimelberg HK. Pharmacological characterization of swellinginduced D-[3H]aspartate release from primary astrocyte cultures. Am J Physiol 274: 1–20, 1998. 81. Longuemare MC, Rose CR, Farrell K, Ransom BR, Waxman SG, and Swanson RA. K+-induced reversal of astrocyte glutamate uptake is limited by compensatory changes in intracellular Na+. Neuroscience 93: 285–292, 1999. 82. Rose CR, Waxman SG, and Ransom BR. Effects of glucose deprivation, chemical hypoxia, and simulated ischemia on Na+ homeostasis in rat spinal cord astrocytes. J Neuropsychiatry 18: 3554–3562, 1998. 83. Silver IA, Deas J, and Erecinska M. Ion homeostasis in brain cells: differences in intracellular ion responses to energy limitation between cultured neurons and glial cells. Neuroscience 78: 589–601, 1997. 84. Kintner DB, Su G, Lenart B, Ballard AJ, Meyer JW, Ng LL, et al. Increased tolerance to oxygen and glucose deprivation in astrocytes from Na+/H+ exchanger isoform 1 null mice. Am J Physiol Cell Physiol 287: C12–C21, 2004. 85. Lenart B, Kintner DB, Shull GE, and Sun D. Na-K-Cl cotransporter-mediated intracellular Na+ accumulation affects Ca2+ signaling in astrocytes in an in vitro ischemic model. J Neuropsychiatry 24: 9585–9597, 2004. 86. Bondarenko A and Chesler M. Rapid astrocyte death induced by transient hypoxia, acidosis, and extracellular ion shifts. Glia 34: 134–142, 2001. 87. Kintner DB, Luo J, Gerdts J, Ballard AJ, Shull GE, and Sun D. Role of Na+-K+-Cl– cotransport and Na+/Ca2+ exchange in mitochondrial dysfunction in astrocytes following in vitro ischemia. Am J Physiol Cell Physiol 292: C1113–C1122, 2007. 88. Waxman SG and Ritchie JM. Molecular dissection of the myelinated axon. Ann Neurol 33: 121–136, 1993. 89. Valeriani V, Dewar D, and McCulloch J. Quantitative assessment of ischemic pathology in axons, oligodendrocytes, and neurons: attenuation of damage after transient ischemia. J Cereb Blood Flow Metab 20: 765–771, 2000.

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90. Tekkok SB and Goldberg MP. Ampa/kainate receptor activation mediates hypoxic oligodendrocyte death and axonal injury in cerebral white matter. J Neuropsychiatry 21: 4237–4248, 2001. 91. Wilke S, Thomas R, Allcock N, and Fern R. Mechanism of acute ischemic injury of oligodendroglia in early myelinating white matter: the importance of astrocyte injury and glutamate release. J Neuropathol Exp Neurol 63: 872–881, 2004. 92. Salter MG and Fern R. NMDA receptors are expressed in developing oligodendrocyte processes and mediate injury. Nature 438: 1167–1171, 2005. 93. Chen H, Kintner DB, Jones M, Matsuda T, Baba A, Kiedrowski L, et al. AMPAmediated excitotoxicity in oligodendrocytes: role for Na(+)-K(+)-Cl(–) co-transport and reversal of Na(+)/Ca(2+) exchanger. J Neurochem, 2007, PM:17490438. 94. Stys PK. General mechanisms of axonal damage and its prevention. J Neurol Neurosurg Psychiatr 233: 3–13, 2005. 95. Czyz A and Kiedrowski L. In depolarized and glucose-deprived neurons, Na+ influx reverses plasmalemmal K+-dependent and K+-independent Na+/Ca2+ exchangers and contributes to NMDA excitotoxicity. J Neurochem 83: 1321–1328, 2002. 96. Hoyt KR, Arden SR, Aizenman E, and Reynolds IJ. Reverse Na+/Ca2+ exchange contributes to glutamate-induced intracellular Ca2+ concentration increases in cultured rat forebrain neurons. Mol Pharmacol 53: 742–749, 1998. 97. Floyd CL, Gorin FA, and Lyeth BG. Mechanical strain injury increases intracellular sodium and reverses Na+/Ca2+ exchange in cortical astrocytes. Glia 51: 35–46, 2005. 98. Taylor CP, Weber ML, Gaughan CL, Lehning EJ, and Lopachin RM. Oxygen/glucose deprivation in hippocampal slices: altered intraneuronal elemental composition predicts structural and functional damage. J Neuropsychiatry 19: 619–629, 1999. 99. Lopachin RM, Gaughan CL, Lehning EJ, Weber ML, and Taylor CP. Effects of ion channel blockade on the distribution of Na, K, Ca and other elements in oxygen-glucose deprived CA1 hippocampal neurons. Neuroscience 103: 971–983, 2001. 100. Galeffi F, Sah R, Pond BB, George A, and Schwartz-Bloom RD. Changes in intracellular chloride after oxygen-glucose deprivation of the adult hippocampal slice: effect of diazepam. J Neuropsychiatry 24: 4478–4488, 2004. 101. Pond BB, Berglund K, Kuner T, Feng G, Augustine GJ, and Schwartz-Bloom RD. The chloride transporter Na(+)-K(+)-Cl– cotransporter isoform-1 contributes to intracellular chloride increases after in vitro ischemia. J Neuropsychiatry 26: 1396–1406, 2006. 102. Kang TC, An SJ, Park SK, Hwang IK, Yoon DK, Shin HS, et al. Changes in Na(+)K(+)-Cl(–) cotransporter immunoreactivity in the gerbil hippocampus following transient ischemia. Neurosci Res 44: 249–254, 2002. 103. O’Donnell ME, Tran L, Lam TI, Liu XB, and Anderson SE. Bumetanide inhibition of the blood-brain barrier Na-K-Cl cotransporter reduces edema formation in the rat middle cerebral artery occlusion model of stroke. J Cereb Blood Flow Metab 24: 1046–1056, 2004. 104. Yam PS, Dunn LT, Graham DI, Dewar D, and McCulloch J. NMDA receptor blockade fails to alter axonal injury in focal cerebral ischemia. J Cereb Blood Flow Metab 20: 772–779, 2000. 105. Dewar D, Yam P, and McCulloch J. Drug development for stroke: importance of protecting cerebral white matter. Eur J Pharmacol 375: 41–50, 1999. 106. Luo J, Wang Y, Chen H, Kintner DB, Cramer SW, Gerdts JK, et al. A concerted role of Na(+)-K(+)-Cl(–) cotransporter and Na(+)/Ca(2+) exchanger in ischemic damage. J Cereb Blood Flow Metab 28: 737–746, 2008. 107. Dzhala VI, Brumback AC, and Staley KJ. Bumetanide enhances phenobarbital efficacy in a neonatal seizure model. Ann Neurol 63: 222–235, 2008.

The Naþ/Hþ Exchanger: A Target for Therapeutic Intervention in Cerebral Ischemia Jin Xue and Gabriel G. Haddad

1 Introduction The mammalian Naþ/Hþ exchanger (NHEs) family is a group of integral membrane transport proteins. It mediates an electroneutral 1:1 exchange of intracellular Hþ for extracellular Naþ and in doing so regulates intracellular pH (pHi) homeostasis and cell volume. NHEs contain two functional domains: the N-terminal transmembrane domain, which is necessary and sufficient to catalyze ion translocation, and the C-terminal cytoplasmic domain, which is crucial for modulating NHE1 activity. To date, nine isoforms have been cloned which have about 25–70% amino acid identity and are predicted to share a characteristic secondary structure. These isoforms differ in their tissue expression, subcellular distribution, kinetic properties, inhibitor sensitivity, and physiological functions. NHE1–5 are localized to the plasma membrane, though NHE3 and NHE5 can enter a recycling endosomal compartment. NHE1 is ubiquitously expressed and is considered to be a ‘‘housekeeping’’ isoform. NHE2–3 are highly expressed in the apical epithelia of kidney and intestine. NHE4 is mainly present in the stomach and basolateral epithelia of kidney. NHE5 predominantly resides in the brain. NHE6–9 are targeted to the membranes of intracellular organelles. Both NHE6 and NHE9 are present chiefly in the recycling endosomes, while NHE7 is largely in the trans-Golgi network and NHE8 is in the mid- to transGolgi pools [36, 39, 27, 47]. NHE1 is by far the most extensively studied isoform and is associated with many pathophysiological conditions. Numerous studies have shown that the activation of NHE1 during cardiac ischemia/reperfusion is deleterious. At present, there is literature on the role of NHE1 in brain ischemia and hypoxia. In this review, we focus on NHE1 and summarize recent progress in our G.G. Haddad (*) Department of Pediatrics and Neuroscience, University of California San Diego, La Jolla, CA 92093-0735; Rady Children’s Hospital, San Diego, CA 92123, USA e-mail: [email protected]

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understanding of the physiological and pathological roles of NHE1 in the brain, particularly under ischemic conditions.

2 Gene Structure The human NHE1 gene was first cloned by Sardet et al. [44]. The gene is located on human chromosome 1p36.1–p35, spans 70 kb and consists of 12 exons and 11 introns [35]. The NHE1 promoter is regulated by several transcription factors, including AP-1, AP-2, C/EBP, chicken ovalbumin upstream promoter transcription factor (COUP-TF) type I and II, as well as thyroid hormone receptor (TRa1). Moreover, a conserved poly (dA:dT) region of the NHE1 promoter is important in regulating its own expression [8]. It is worth mentioning that the NHE1 promoter is responsive to reactive oxygen species (ROS) [1]. This finding is of clinical importance because NHE1 expression/activity is elevated following ischemic disease, which may be related to the effects of ROS on the NHE1 promoter.

3 Structural Features NHE1 protein contains 815 amino acids with a calculated molecular weight of 85 kDa. But NHE1 has an apparent size of 110 kDa due to its N- and O-linked glycosylation on the extracellular loop 1 (EL1). Although NHE1 forms a stable homodimer in intact cells, individual subunits do function independently. The topological analysis has revealed that NHE1 consists of two domains: an amphipathic N-terminal membrane domain of 500 amino acids and a hydrophilic C-terminal cytoplasmic domain of 315 amino acids. The highly conserved N-terminal domain has 12 transmembrane spanning regions (TM I-XII) and 3 membrane-associated loops (intracellular loops 2 and 4 and extracellular loop 5), and is responsible for cation translocation and drug recognition. The less conserved C-terminal domain includes an ‘‘Hþ sensor’’ and serves to mediate regulation by other molecules (Fig. 1) [47, 8]. The crystal structure of Naþ/Hþ antiporter from Escherichia coli (EcNhaA) was recently elucidated [14]. Using it as a template, a 3D structure of human NHE1 is predicted. The model structure features a cluster of titratable residues that are evolutionarily conserved and are involved in the cation binding and translocation. Furthermore, an alternating-access mechanism is proposed for the transport of cations across the membrane. Low pH signal (Hþ) is probably attracted by the acidic Glu262 at the entry to the cytoplasmic funnel, then protonates Asp267. The protonation elicits a conformational change and leads to the transfer Hþ from cytoplasmic funnel to extracellular funnel. Driven by chemical gradients of Naþ and Hþ across the membrane, Hþ is exchanged for Naþ perhaps via Ser351 in the extracellular space. Finally, the movement to the alternative conformation allows the replacement of Naþ by Hþ at the cytoplasmic side. The continuance of the cycle is controlled by pHi [25].

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Fig. 1 Model of NHE1 structure. Upper panel: topology of the membrane domain which functions to transport cations. This illustration is based on the findings of [50]. Pink circles, residues implicated in both ion transport and inhibitor binding; orange circles, residues implicated in ion binding and transport; yellow circles, residues implicated in inhibitor binding; green circles, residues implicated in NHE1 folding and targeting to the plasma membrane. Lower panel: representation of the cytoplasmic domain which functions to regulate the membrane domain through interactions with signaling molecules [47] (with Journal permission)

4 Molecular Biology As illustrated in Fig. 1, the transmembrane N-terminal domain mediates cation transport and inhibitor recognition, while the cytoplasmic C-terminal domain regulates NHE1 activity and is associated with a number of functionally distinct signaling molecules. Table 1 summarizes (1) the N-terminal: the amino acids that are critical for exchanger activity, Naþ affinity, inhibitor binding/sensitivity, pHi sensing, and membrane targeting of NHE1 and (2) the C-terminal: regulatory proteins and their binding sites [36, 47, 33, 54].

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Table 1 Molecular biology of NHE1 Transmembrane (TM) N-terminal domain Function Exchanger activity Naþ affinity/inhibitor binding Inhibitor sensitivity pHi sensing Membrane targeting Cytoplasmic C-terminal domain Regulatory proteins Phosphatidylinositol 4,5-bisphosphate (PIP2) Calcineurin B homologous protein (CHP) Ezrin, radixin, moesin (ERM) family Nck-interacting kinase (NIK) Ca2þ/calmodulin Adaptor protein 14-3-3b Tescalcin Carbonic anhydrase II (CAII) Heat shock protein (HSP)70 6-phosphofructokinase-1 (PFK-1) Src homology 2 domain-containing protein tyrosine phosphatase (SHP-2)

Critical amino acids Asp159, Pro167, Pro168, Phe161 in TM IV Glu262, Asp267 in TM VII Phe161, Phe162, Leu163, Gly174 in TM IV Leu163 and Gly174 in TM IV His349 in TM IX Gly455 and Gly456 in TM XI Arg440 in IL5 Tyr454 and Arg458 in TM XI Binding sites Amino acids 513–520 and 556–564 Amino acids 515–530 Amino acids 553–564 Amino acids 538–638 Amino acids 636–656 for high-affinity site, Amino acids 657–684 for low-affinity site Amino acids 700–705 Amino acids 633–815 Amino acids 790–802 Un-determined Un-determined Un-determined

5 Tissue and Cellular Distribution NHE1 is ubiquitously expressed on the plasma membrane of virtually all mammalian cell types. In several cell types, NHE1 is not uniformly distributed along the plasma membrane but preferentially accumulates at discrete microdomains, for example, at the basolateral membrane of polarized epithelial cells, at intercalated disks and transverse tubules in cardiomyocytes, and at membrane protrusions or lamellipodia in fibroblasts [39]. We and others have demonstrated that both mRNA and protein of NHE1 are present at high levels throughout the CNS [37, 30, 7].

6 Biophysical and Electrophysiological Properties Electroneutral ion flux via NHE1 is driven by an inwardly directed gradient for Naþ and an outward gradient for Hþ, requiring no direct energy input. NHE1 exhibits a simple Michaelis–Menten dependence on extracellular Naþ, with a Km of 5–50 mM. Extracellular Liþ or Hþ competes with Naþ for the

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Naþ-binding site and extracellular Hþ suppresses NHE1 activity. In contrast, intracellular acidification allosterically stimulates NHE1 activity with a Hill coefficient of 3, which is attributed to the presence of an ‘‘Hþ-modifier site’’. During ischemia, both extracellular and intracellular pH drop, the net effect on NHE1 activity relies on the extent of extracellular inhibition versus intracellular activation. Growth factors and hormones as well as osmotic stress can activate NHE1 at normal pHi by causing alkaline shift in the set point pHi (i.e., a reduced Km for intracellular Hþ), which is believed to be at least partially dependent on protein phosphorylation/dephosphorylation [36, 39].

7 Regulatory Mechanisms: Receptorial, Transcriptional, and Transductional NHE1 is activated by a wide variety of stimuli including intracellular acidification, growth factors, hormones, cytokines, osmotic, and mechanical stresses. This activation takes place through distinct cell surface receptors including receptor tyrosine kinase, G-protein-coupled receptors, integrin receptors, and intracellular signaling networks. Modulation of NHE1 activity occurs at the COOH terminus by at least three mechanisms: (1) phosphorylation, a number of serine and threonine residues in the distal C-tail of NHE1 (amino acids 700–815) can be phosphorylated by protein kinases in response to sustained acidosis, hormones or growth factors stimulation. Such kinases include extracellular-signal-regulated kinase 1/2 (ERK 1/2), p90 ribosomal S6 kinase (p90rsk), Rho-activated kinase p160ROCK, Nck-interacting kinase (NIK), Ca2þ/calmodulin-dependent kinase II (CaMKII), and p38 mitogen-activated protein kinase (p38 MAPK). The phosphorylation results in increased NHE1 activity except for p38 MAPK, which appears to inhibit exchanger activity in response to angiotensin II. Protein kinases C and D also affect NHE1 activity but are thought not due to direct phosphorylation; (2) interaction with regulatory proteins, e.g., association with phosphatidylinositol 4,5-bisphosphate (PIP2), calcineurin B homologous protein (CHP), Ca2þ/calmodulin (CaM), adaptor protein 14-3-3b or carbonic anhydrase II (CAII) enhances NHE1 activity while with tescalcin inhibits NHE1 activity. The interaction with ezrin/radixin/moesin (ERM) is important in restrictive localization of NHE1 and plays role in structure integrity, focal adhesion, and cell migration. Src homology 2 domain-containing protein tyrosine phosphatase (SHP2)-NHE1 interaction influences NHE1 functions in pHi regulation, cell proliferation, and cell death under hypoxia; and (3) allosteric modulation via altering affinity for Hþ at ‘‘Hþ-modifier site’’ (see above Section 6) [47, 8, 33, 54]. In addition, NHE1 can be regulated at the transcriptional level (see above Section 2). Vice versa, NHE1 deficiency alters gene expression in mouse brain [57] and NHE1 activity regulates the expression of genes associated with growth factor signaling, oncogenesis, and cell cycle progression in fibroblasts [43]. In the CNS,

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ischemia/reperfusion [28, 56, 45] and epilepsy [15] elevate NHE1 expression/ activity. In contrast, chronic intermittent hypoxia downregulates the expression of NHE1 [6].

8 Physiological Properties The physiological roles of NHE1 include: (1) pHi regulation, NHE1 is activated by intracellular acidification and mediates Hþ efflux which efficiently restores pHi; (2) cellular volume regulation, NHE1 is stimulated by cell shrinkage and mediates Naþ influx, along with Cl- influx via anion exchange, which normalizes cell volume; (3) cell proliferation, NHE1-induced increase in pHi promotes G2/M entry and transition, thus facilitates cell cycle progression; (4) cell growth and differentiation; (5) an anchor for actin filament and a scaffold for the assembly of signaling complexes, roles in cytoskeleton remodeling, cell shape, focal adhesion, cell migration as well as in signal relay; (6) apoptosis, role of NHE1 differs depending on stimulation and cell type; and (7) a possible mediator of immunity [36, 39, 33, 5].

9 Pathophysiological Relevance in Stroke Cerebral ischemia is a complex pathological process, which results in energy failure, cell swelling, membrane depolarization, abnormal release of excitatory neurotransmitters and Kþ ions, spreading depression, injurious elevation in [Naþ]i and [Ca2þ]i, generation of free radicals, blood–brain barrier (BBB) disruption and edema, as well as an inflammatory response. Ionic disturbance, particularly for Hþ, Naþ, Caþ2, and Kþ, is an early event in the cascade leading to hypoxic injury in the mammalian brain. NHE1, a major regulator of pHi and ionic homeostasis, has been implicated in ischemia/reperfusion injury. NHE1 is widely distributed in neurons, glial cells, BBB endothelial cells, and choroid plexus [39]. Because of its high metabolic rate, the CNS produces a large amount of metabolic acid, particularly under stressful conditions, such as hypoxia/ischemia. This feature renders neurons/glial cells more vulnerable to injury from acidosis. Therefore, the efficient acid-extrusion mechanism is crucial for proper brain function. The growing evidence indicates that NHE1 is a key player in pHi regulation in both neurons and glial cells under steady-state conditions and after intracellular acidification [40]. Consistent with this view, acutely dissociated CA1 neurons from NHE–/– mice or from NHEþ/+ mice treated with NHE1 inhibitor HOE694 exhibit reduced steady-state pHi and reduced recovery rate from acid load [55]. Similar results have been reported in primary cortical neurons and astrocytes either with inhibitor or from genetic ablation [29, 20]. Both beneficial and detrimental effects of acidic pH (‘‘pH paradox’’) have been described in the brain [39]. Mild acidosis is protective in

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cerebral ischemia by attenuating Ca2þ influx, by reducing energy demand and ATP depletion [48], and by inhibition of NHE1 via acidic pHo [49]. However, if pHi is too low, cell death ensues due to: (1) depolarization, over-excitability and oxygen consumption; (2) potentiation of free radical formation; (3) inhibition of mitochondrial metabolism and glycolysis, thus accelerating ATP depletion; and (4) enhancement of the neurotoxicity mediated by -amino-3-hydroxy-5methyl-4-isoxazole propionic acid receptors [52]. Ischemia/hypoxia stimulates NHE1 by a reduction in pHi or via signaling pathways (e.g., ERK-p90RSK or PKA or PKC) [28, 56, 45, 18]. In acutely isolated mouse CA1 neurons, 5 min of chemical anoxia elicits pHi elevation in HEPES buffer, which can be reduced in NHE1 null mice, or by NHE1 antagonist 3-methylsulfonyl-4-piperidinobenzoyl-guanidine methansulfonate (HOE694), or inhibitors of PKA or PKC, indicating that anoxia activates NHE1 acutely, probably via PKA or PKC (Fig. 2) [56]. Similarly, in rat hippocampal CA1 neurons, NHE1 is stimulated immediately after anoxia via cAMP–PKA signaling, yet is apparently inhibited during the anoxia phase,

Fig. 2 NHE1 contributes to anoxia-induced pHi changes in mouse CA1 neurons. (1) Effect of HOE-694 on the anoxia-induced pHi change in HEPES buffer. A: With 100 mM HOE-694 in HEPES buffer, 5 min of anoxia induced no change in pHi in contrast to an increase in pHi in controls. B: Mean changes of pHi in control and the HOE-694 group. Means are significantly different from each other. (2). Anoxia-induced pHi increase was smaller in the NHE1 null mutant neurons. A: Two neurons (one was a wild-type (WT) and the other a NHE1 mutant) with similar initial pHi. Both neurons were perfused with HEPES buffer and subjected to 5 min of anoxia. Although both neurons had a similar pattern in pHi change in response to anoxia, a smaller increase was seen in the mutant neuron. B: Mean change of pHi induced by 5 min of anoxia in both NHE1 mutant and wild-type neurons. Means are significantly different from each other [56] (with Journal permission)

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Fig. 2 (continued)

possibly due to a fall in cellular ATP level [46]. However, OGD/REOX (oxygen and glucose deprivation/re-oxygenation)-induced increase in NHE1 activity is independent of prolonged acidosis but depends in part on the activation of ERK1/2 in cultured cortical neurons or astrocytes [28, 18]. The role of NHE1 in cerebral ischemia has been examined in two types of in vitro ischemic models, the hypoxic, acidic, ion-shifted Ringer’s solution (HAIR) or OGD/REOX. HAIR solution simulates interstitial ionic changes of cerebral ischemia. After a 20–40-min exposure to HAIR, most astrocytes die within 30 min upon return to normal saline (i.e., ‘‘reperfusion’’). Astrocyte death requires external Ca2þ and is blocked by 2-[2-[4-(4-nitrobenzyloxy)phenyl]ethyl] isothiourea methanesulfonate (KB-R7943), an inhibitor of reverse Naþ–Ca2þ exchange (NCX), suggesting that injury is triggered by a rise in [Ca2þ]i. Moreover, the rise of [Ca2þ]i and injury are prevented by NHE inhibitors ethyl isopropyl amiloride and HOE-694. Acidic reperfusion media, which inhibit NHE1 activity, is also protective. These data indicate that Naþ loading via NHE1 fosters reversal of NCX and subsequent cytotoxic elevation of [Ca2þ]i, which eventually causes ischemic damage of astrocytes [19, 3]. In agreement with this idea, OGD/REOX evokes a 7-fold and 1.5fold increase in [Naþ]i and [Ca2þ]i, respectively in cortical neurons, accompanied by mitochondrial Ca2þ accumulation, cytochrome c release and neuronal death. These deleterious changes can be attenuated by NHE1 inhibitor (4-Isopropyl-3methylsulfonyl-benzoyl)-guanidine methanesulfonate (HOE642) or genetic ablation of NHE1 activity or 2-[4-[(2,5-difluorophenyl) methoxy]phenoxy]-5-ethoxyaniline (SEA0 400, a potent blocker of reversal mode of NCX), confirming again the importance of the above-mentioned injury mechanism (Fig. 3) [29].

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Fig. 3 Inhibition or ablation of NHE1 activity abolishes OGD-mediated cell death. Cell mortality was assessed in 12–15 DIV cortical neurons after 3 h of OGD and 21 h of REOX. For HOE 642 treatment, NHE1þ/+ neurons were incubated in the presence of 1 mM HOE 642 at 378C for 3 h of OGD and 21 h of REOX. Sister NHE1þ/+ cultures were incubated for 24 h in normoxic control buffers (Con). Similar assays were also performed in NHE1þ/– and NHE1–/– cultures. At the end of the experiment, cells were stained with PI and calcein-AM, and cell images were acquired. A, C, E, Calcein-AM. B, D, F, PI. A, B, Con. C–F, OGD/ REOX. E, F, OGD/REOX plus HOE. Scale bar, 256 mm. G, summary data. Data are means SD (n = 5–8 cultures). *P < 0.05 versus Con; #P < 0.05 versus NHE1þ/+ OGD/REOX. Error bars represent SD [29] (with Journal permission)

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Cerebroprotection by NHE1 blockade has been validated in an in vivo ischemic model (middle cerebral artery occlusion (MCAO)). A marked decrease of infarct volume is found in NHE1þ/– mice or HOE642-treated NHEþ/+ mice following 1-h focal ischemia (Fig. 4) [29]. Other NHE1 inhibitors, N-(aminoiminomethyl)-1-methyl-1 H-indole-2-carboxamide methanesulfonate (SM-20220), 3-(2,5-dichlorothiophen-3-yl)-N-[2-(dimethylamino)ethyl]-5-guanidinocarbonylbenzamide dihydrochloride (FR183998), and N-carbamimidoyl-4-[4-(1Hpyrrol-2-ylcarbonyl)piperazin-1-yl]-3-(trifluoromethyl) benzamide (sabipor-

Fig. 4 Inhibition or knockdown of NHE1 activity reduces infarction after focal ischemia. (1) Inhibition of NHE1 activity. A, B, After 2 h of MCAO and 24 h of reperfusion, saline-treated control and HOE 642-treated NHE1þ/+ mice were killed. Infarction volume (outlined area) was determined by TTC staining. Data are means SD (n = 4–6). *P < 0.05 versus NHE1þ/+. Scale bar, 5 mm. C, focal ischemia induced a significant increase in water content. *P < 0.01 versus contralateral. Error bars represent SD. (2) Knockdown of NHE1 activity. A, crude cortex membrane proteins of three genotypes were separated electrophoretically. Expression of NHE1 and tubulin III in NHE1þ/+, NHE1þ/–, and NHE1–/– mouse cortex was shown on the same blot. Densitometric analysis of immunoblots was presented as a ratio of NHE1/-tubulin III band intensity. Data are means SD (n = 3–5). B, C, After 2 h MCAO and 24 h reperfusion, infarction volume in NHE1þ/+ or NHE1þ/– mice was determined by TTC staining. *P < 0.05 versus NHE1þ/+. Data are means SD (n = 4). Scale bar, 5 mm. D, water content in NHE1þ/ + and NHE1þ/– mice after 2 h of MCAO and 24 h of reperfusion. Data are means SD (n = 4–6). *P < 0.01 versus contralateral. Error bars represent SD [29] (with Journal permission)

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ide), have also been proven to provide similar beneficial effects in MCAO model [22, 11, 21, 38]. Most importantly, SM-20220 [24] and sabiporide [38] show effectiveness even with post-ischemia administration, emphasizing their therapeutic potential for stroke. Brain edema is another characteristic of cerebral ischemia. It is formed during early hours of ischemia and by a process involving increased secretion of Naþ and water across an intact BBB with a concomitant increased uptake of Naþ, Cl–, and water into astrocytes which cause cytotoxic edema. NHE1 along with Naþ-Kþ-2Cl– cotransporter isoform 1 (NKCC1) contributes to both BBB secretion and astrocyte-uptake processes [17]. Supporting this notion is that NHE1 inhibition significantly alleviates cellular swelling induced either by OGD/REOX [29, 20] or glutamate [32] in cultured cortical neurons or astrocytes. Moreover, reduction on the extent of cerebral edema by NHE1 blockade is observed in an in vivo ischemic model (MCAO) [22, 21, 38]. Cerebroprotective benefits of NHE1 inhibition is manifested further by the facts that NHE1 inhibition: (1) attenuates leukocyte adhesion [10] and improves endothelial dysfunction [12] induced by ischemia-reperfusion; (2) facilitates consciousness recovery and preserves neurological function after transient cerebral ischemia [23]; (3) reduces the magnitude of the ischemia-induced increase in locomotor neuronal activity, which represents the degree of damage of hippocampal CA1 neurons [41]; (4) protects against excitotoxicity [38, 32]; and (5) prevents ischemia/reperfusion-elicited free fatty acid release [42]. Spontaneous [4] and targeted [2] NHE1–/– mice suffer from epileptic-like seizure, ataxia, growth retardation, and selective neuronal death in the cerebellum and brainstem, suggesting that NHE1 is important in neuronal function, growth, and survival. Mice lacking NHE1 have been shown to (1) upregulate Naþ channel expression in hippocampal and cortical regions, leading to an increase in Naþ current density and neuronal excitability [9, 51]; (2) change the expression of other membrane transporters at both protein and mRNA levels [53]; and (3) alter the expression of genes implicated in neurodegeneration [57]. These findings may help us understand the molecular basis of the phenotypical changes in NHE1 null mutant and potentially why they seize. Systemic administration of NHE1 inhibitors is associated with a higher rate of cerebrovascular events [34] and shows neurotoxic effects [13] with the latter being reversible by 7 days after treatment. Clearly, future investigation is needed to delineate the precise mechanism of the pertinent adverse effects in order to dissociate such unwanted side effects from the cerebroprotective efficacy of NHE1 inhibitors.

10 Pharmacological Modulation Pharmacological inhibitors of NHE1 are categorized into three groups: (1) amiloride and its analogues; (2) benzoylguanidine and its derivatives; and (3) bicyclic guanidines. Amiloride, a Kþ-sparing diuretic, was the first drug

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described as a NHE1 inhibitor, which also inhibits a conductive Naþ channel, Naþ/Ca2þ exchanger, and Naþ/Kþ ATPase. Double substitution of the nitrogen of the 5-amino group of amiloride gives dimethylamiloride (DMA), 5-(Nethyl-N-isopropyl)-amiloride (EIPA), 5-(N-methyl-N-isobutyl)-amiloride (MIBA), and 5-(N, N-hexamethylene) amiloride (HMA). They are much more effective than amiloride but weak selective toward NHE1. The simultaneous replacement of the pyrazine ring by a phenyl and of the 6-chloro by a sulfomethyl leads to drugs such as HOE-694, HOE642, 2-methyl-5-methylsulfonyl-4-(1-pyrrolyl)-benzoylguanidine methanesulfate (eniporide), and benzamide-N-(aminoiminomethyl)-4-[4-(2-furanylcarbonyl)-1-piperazinyl]-3(methylsulfonyl)methanesulfonate (BIIB-513), which have high potency/selectivity for NHE1, excellent solubility, resorption, and bioavailability. In the last decade several molecules based on the bicyclic template have been designed, including [1-(quinolin-5-yl)-5-cyclopropyl-1H-pyrazole-4-carbonyl] guanidine (zoniporide), SM-20220, N-(aminoiminomethyl)-1,4-dimethyl-1H-indole-2carboxamide methanesulfonic acid (SM-20550), (1R,3R)-N-(Diaminomethylene)-3-(2,3-dihydro-1-benzofuran-4-yl)-2,2-dimethylcyclopropanecarboxamide (BMS-284640), (5E,7S)-[[7-(5-fluoro-2-methylphenyl)-4-methyl-7,8-dihydro-5(6H)-quinolinylidene]amino]guanidine dimethanesulfonate (T-162559), 6,7,8, 9-tetrahydro-2-methyl-5H-cyclohepta[b]pyridine-3-carbonylguanidine maleate (TY-12533), or 3-[(cyclopropylcarbonyl)amino]-N-[2-(dimethylamino)ethyl]-4-[4-(5-methyl-1H-imidazol-4-yl)piperidin-1-yl]benzamide (SL 59.1227). The IC50 for the human NHE1 are as follows: Amiloride = 10.7 mM, Cariporide = 0.08 mM, T-165229 = 13 nM. Both chemical structure and ionization of the guanidine determine the potency of NHE inhibitors. During ischemia/ reperfusion, the pH can fall down to 6.2 or lower, drugs like cariporide (pKa = 6.28), TY-12533 (pKa = 6.93) or zoniporide (pKa = 7.2), which are positively charged, become more efficient. Indeed, cariporide and TY-12533 are more active at pH 6.2 than 6.7 (cariporide: IC50 = 22 nM/120 nM; TY-12533: IC50=17 nM/32 nM). In an aqueous medium, Naþ ion is surrounded by three molecules of H2O and guanidinium ion of inhibitors is similar in structure, thus can block NHE1 by competitive binding to the extracellular Naþ-binding site [26, 31].

11 Preliminary Clinical Trials Extensive preclinical studies have demonstrated that NHE1 inhibition affords excellent and consistent protection from myocardial ischemia/reperfusion injury in a wide variety of experimental models and animal species by different investigators [16]. However, in human, clinical trials have resulted in only modest success [34], i.e., a significant reduction in the combined endpoint of death and myocardial infarction in high-risk patients undergoing coronary artery bypass graft surgery. Unfortunately, this beneficial effect is associated with a higher

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rate of cerebrovascular events. The underlying mechanisms for increased cerebrovascular risk are unknown. Further investigation is needed to determine whether it reflects NHE1 inhibition in general or whether this is a specific effect of a particular inhibitor. Our current knowledge regarding the role of NHE in cerebral ischemia is limited, though emerging evidence points to cerebroprotective effects of NHE inhibition. More evaluation is warranted in animal models of cerebral ischemia before devising clinical trial.

12 Therapeutic Perspectives Stroke is the second-most prevalent cause of death worldwide. About 80% of all strokes are ischemic. Despite years of research efforts, the effective treatment and prevention of ischemic brain injury remains a major medical challenge. NHE1 inhibition with either pharmacological agents or genetic ablation has been reported to reduce brain damage after ischemia/reperfusion insult, both in vitro culture and in vivo animal models. These encouraging findings hold promise for a new drug that could significantly reduce injury associated with cerebral ischemia.

13 Conclusions In this review, we have documented the research progress in understanding the physiopathological roles of NHE1 in the CNS. Although NHE1 is pivotal for maintaining pHi and ionic homeostasis under physiological stresses, its excessive activity, such as following cerebral ischemia, may be a major cause of cell damage and death. The possible injury mechanisms by ischemia-induced NHE1 activation include (1) [Naþ]i accumulation leading to NCX reversal, elevated [Ca2þ]i, and consequent cell death that may involve mitochondrial death pathway; (2) changes in pHi; (3) brain edema and cell swelling; (4) activation of MAPKs; and (5) release of excitatory amino acids or ROS [39]. Cerebroprotective roles of NHE1 inhibition have been demonstrated in in vitro and in vivo ischemic models, making it an attractive therapeutic target for cerebral ischemia. However, adverse effects by NHE1 inhibitors have also been noted. Thus, better understanding of the underlying mechanisms both for protection and for side effects should be future research focus.

References 1. Akram S, Teong HF, Fliegel L, Pervaiz S, and Clement MV. Reactive oxygen speciesmediated regulation of the Na+-H+ exchanger 1 gene expression connects intracellular redox status with cells’ sensitivity to death triggers. Cell Death Differ 13: 628–641, 2006. 2. Bell SM, Schreiner CM, Schultheis PJ, Miller ML, Evans RL, Vorhees CV, Shull GE, and Scott WJ. Targeted disruption of the murine Nhe1 locus induces ataxia, growth retardation, and seizures. Am J Physiol 276: C788–C795, 1999.

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3. Bondarenko A, Svichar N, and Chesler M. Role of Na+-H+ and Na+-Ca2+ exchange in hypoxia-related acute astrocyte death. Glia 49: 143–152, 2005. 4. Cox GA, Lutz CM, Yang CL, Biemesderfer D, Bronson RT, Fu A, Aronson PS, Noebels JL, and Frankel WN. Sodium/hydrogen exchanger gene defect in slow-wave epilepsy mutant mice. Cell 91: 139–148, 1997. 5. De Vito P. The sodium/hydrogen exchanger: a possible mediator of immunity. Cell Immunol 240: 69–85, 2006. 6. Douglas RM, Xue J, Chen JY, Haddad CG, Alper SL, and Haddad GG. Chronic intermittent hypoxia decreases the expression of Na/H exchangers and HCO3-dependent transporters in mouse CNS. J Appl Physiol 95: 292–299, 2003. 7. Douglas RM, Schmitt BM, Xia Y, Bevensee MO, Biemesderfer D, Boron WF, and Haddad GG. Sodium-hydrogen exchangers and sodium-bicarbonate co-transporters: ontogeny of protein expression in the rat brain. Neuroscience 102: 217–228, 2001. 8. Fliegel L. Molecular biology of the myocardial Na+/H+ exchanger. J Mol Cell Cardiol 44: 228–237, 2008. 9. Gu XQ, Yao H, and Haddad GG. Increased neuronal excitability and seizures in the Na(+)/ H(+) exchanger null mutant mouse. Am J Physiol Cell Physiol 281: C496–C503, 2001. 10. Horikawa N, Kuribayashi Y, Matsui K, Kawamura N, and Ohashi N. Na+/H+ exchange inhibitor SM-20220 attenuates leukocyte adhesion induced by ischemia-reperfusion. J Cardiovasc Pharmacol 37: 668–677, 2001. 11. Horikawa N, Nishioka M, Itoh N, Kuribayashi Y, Matsui K, and Ohashi N. The Na(+)/ H(+) exchanger SM-20220 attenuates ischemic injury in in vitro and in vivo models. Pharmacology 63: 76–81, 2001a. 12. Horikawa N, Kuribayashi Y, Itoh N, Nishioka M, Matsui K, Kawamura N, and Ohashi N. Na+/H+ exchange inhibitor SM-20220 improves endothelial dysfunction induced by ischemia-reperfusion. Jpn J Pharmacol 85: 271–277, 2001. 13. Hubler N, Gottschling B, Jacobs M, von Landenberg F, and Hewicker-Trautwein M. Functional observational battery and motor activity in rats after single administration of two NHE 1 inhibitors. Toxicol Appl Pharmacol 208: 266–276, 2005. 14. Hunte C, Screpanti E, Venturi M, Rimon A, Padan E, and Michel H. Structure of a Na+/ H+ antiporter and insights into mechanism of action and regulation by pH. Nature 435: 1197–1202, 2005. 15. Kang TC, An SJ, Park SK, Hwang IK, Suh JG, Oh YS, Bae JC, and Won MH. Alterations in Na+/H+ exchanger and Na+/HCO3 – cotransporter immunoreactivities within the gerbil hippocampus following seizure. Brain Res Mol Brain Res 109: 226–232, 2002. 16. Karmazyn M, Sawyer M, and Fliegel L. The Na(+)/H(+) exchanger: a target for cardiac therapeutic intervention. Curr Drug Targets Cardiovasc Haematol Disord 5: 323–335, 2005. 17. Kimelberg HK. Cell swelling in cerebral ischemia. In: Walz W and Totawa HJ (ed) Cerebral Ischemia: Molecular and Cellular Pathophysiology. Clifton: Humana, 1999. 18. Kintner DB, Look A, Shull GE, and Sun D. Stimulation of astrocyte Na+/H+ exchange activity in response to in vitro ischemia depends in part on activation of ERK1/2. Am J Physiol Cell Physiol 289: C934–C945, 2005. 19. Kintner DB, Luo J, Gerdts J, Ballard AJ, Shull GE, Sun D. Role of Na+-K+-Cl– cotransport and Na+/Ca2+ exchange in mitochondrial dysfunction in astrocytes following in vitro ischemia. Am J Physiol Cell Physiol 292: C1113–C1122, 2007. 20. Kintner DB, Su G, Lenart B, Ballard AJ, Meyer JW, Ng LL, Shull GE, and Sun D.. Increased tolerance to oxygen and glucose deprivation in astrocytes from Na(+)/H(+) exchanger isoform 1 null mice. Am J Physiol Cell Physiol 287: C12–C21, 2004. 21. Kitayama J, Kitazono T, Yao H, Ooboshi H, Takaba H, Ago T, Fujishima M, Ibayashi S. Inhibition of Na+/H+ exchanger reduces infarct volume of focal cerebral ischemia in rats. Brain Res 922: 223–228, 2001.

A Target for Therapeutic Intervention in Cerebral Ischemia

127

22. Kuribayashi Y, Itoh N, Kitano M, and Ohashi N. Cerebroprotective properties of SM20220, a potent Na(+)/H(+) exchange inhibitor, in transient cerebral ischemia in rats. Eur J Pharmacol 383: 163–168, 1999. 23. Kuribayashi Y, Itoh N, Horikawa N, and Ohashi N. SM-20220, a potent Na+/H+ exchange inhibitor, improves consciousness recovery and neurological outcome following transient cerebral ischaemia in gerbils. J Pharm Pharmacol 52: 441–444, 2000. 24. Kuribayashi Y, Horikawa N, Itoh N, Kitano M, and Ohashi N. Delayed treatment of Na+/H+ exchange inhibitor SM-20220 reduces infarct size in both transient and permanent middle cerebral artery occlusion in rats. Int J Tissue React 21: 29–33, 1999b. 25. Landau M, Herz K, Padan E, and Ben-Tal N. Model structure of the Na+/H+ exchanger 1 (NHE1): functional and clinical implications. J Biol Chem 282: 37854–37863, 2007. 26. Lang HJ. Chemistry of NHE inhibitors. In: Karmazyn M, Avkiran M, Fliegel L (ed) The Sodium–Hydrogen Exchanger: From Molecule to Its Role in Disease. Boston/Dordrecht/London: Kluwer Academic Publishers, 2003. 27. Luo J and Sun D. Physiology and pathophysiology of Na(+)/H(+) exchange isoform 1 in the central nervous system. Curr Neurovasc Res 4: 205–215, 2007. 28. Luo J, Kintner DB, Shull GE, and Sun D. ERK1/2-p90RSK-mediated phosphorylation of Na+/H+ exchanger isoform 1. A role in ischemic neuronal death. J Biol Chem 282: 28274–28284, 2007. 29. Luo J, Chen H, Kintner DB, Shull GE, and Sun D. Decreased neuronal death in Na+/H+ exchanger isoform 1-null mice after in vitro and in vivo ischemia. J Neurosci 25: 11256–11268, 2005. 30. Ma E and Haddad GG. Expression and localization of Na+/H+ exchangers in rat central nervous system. Neuroscience 79: 591–603, 1997. 31. Masereel B, Pochet L, and Laeckmann D. An overview of inhibitors of Na(+)/H(+) exchanger. Eur J Med Chem 38: 547–554, 2003. 32. Matsumoto Y, Yamamoto S, Suzuki Y, Tsuboi T, Terakawa S, Ohashi N, and Umemura K. Na+/H+ exchanger inhibitor, SM-20220, is protective against excitotoxicity in cultured cortical neurons. Stroke 35: 185–190, 2004. 33. Meima ME, Mackley JR, and Barber DL. Beyond ion translocation: structural functions of the sodium-hydrogen exchanger isoform-1. Curr Opin Nephrol Hypertens 16: 365–372, 2007. 34. Mentzer RM Jr, Bartels C, Bolli R, Boyce S, Buckberg GD, Chaitman B, Haverich A, Knight J, Menasche P, Myers ML, Nicolau J, Simoons M, Thulin L, and Weisel RD. Sodium-hydrogen exchange inhibition by cariporide to reduce the risk of ischemic cardiac events in patients undergoing coronary artery bypass grafting: results of the EXPEDITION study. Ann Thorac Surg 85: 1261–1270, 2008. 35. Miller RT, Counillon L, Pages G, Lifton RP, Sardet C, and Pouyssegur J. Structure of the 5’-flanking regulatory region and gene for the human growth factor-activatable Na/H exchanger NHE-1. J Biol Chem 266: 10813–10819, 1991. 36. Orlowski J and Grinstein S. Diversity of the mammalian sodium/proton exchanger SLC9 gene family. Pflugers Arch 447: 549–565, 2004. 37. Orlowski J, Kandasamy RA, and Shull GE. Molecular cloning of putative members of the Na/H exchanger gene family. cDNA cloning, deduced amino acid sequence, and mRNA tissue expression of the rat Na/H exchanger NHE-1 and two structurally related proteins. J Biol Chem 267: 9331–9339, 1992. 38. Park HS, Lee BK, Park S, Kim SU, Lee SH, Baik EJ, Lee S, Yi KY, Yoo SE, Moon CH, and Jung YS. Effects of sabiporide, a specific Na+/H+ exchanger inhibitor, on neuronal cell death and brain ischemia. Brain Res 1061: 67–71, 2005. 39. Pedersen SF, O’Donnell ME, Anderson SE, and Cala PM. Physiology and pathophysiology of Na+/H+ exchange and Na+-K+-2Cl– cotransport in the heart, brain, and blood. Am J Physiol Regul Integr Comp Physiol 291: R1–R25, 2006.

128

J. Xue and G.G. Haddad

40. Pedersen SF, Jorgensen NK, Damgaard I, Schousboe A, and Hoffmann EK. Mechanisms of pHi regulation studied in individual neurons cultured from mouse cerebral cortex. J Neurosci Res 51: 431–441, 1998. 41. Phillis JW, Estevez AY, Guyot LL, and O’Regan MH. 5-(N-Ethyl-N-isopropyl)-amiloride, an Na(+)-H(+) exchange inhibitor, protects gerbil hippocampal neurons from ischemic injury. Brain Res 839: 199–202, 1999. 42. Pilitsis JG, Diaz FG, O’Regan MH, and Phillis JW. Inhibition of Na(+)/H(+) exchange by SM-20220 attenuates free fatty acid efflux in rat cerebral cortex during ischemiareperfusion injury. Brain Res 913: 156–158, 2001. 43. Putney LK and Barber DL. Expression profile of genes regulated by activity of the Na-H exchanger NHE1. BMC Genomics 5: 46, 2004. 44. Sardet C, Franchi A, and Pouyssegur J. Molecular cloning, primary structure, and expression of the human growth factor-activatable Na+/H+ antiporter. Cell 56: 271–280, 1989. 45. Sheldon C and Church J. Intracellular pH response to anoxia in acutely dissociated adult rat hippocampal CA1 neurons. J Neurophysiol 87: 2209–2224, 2002. 46. Sheldon C and Church J. Reduced contribution from Na+/H+ exchange to acid extrusion during anoxia in adult rat hippocampal CA1 neurons. J Neurochem 88: 594–603, 2004. 47. Slepkov ER, Rainey JK, Sykes BD, and Fliegel L. Structural and functional analysis of the Na+/H+ exchanger. Biochem J 401: 623–633, 2007. 48. Trafton J, Tombaugh G, Yang S, and Sapolsky R. Salutary and deleterious effects of acidity on an indirect measure of metabolic rate and ATP concentrations in CNS cultures. Brain Res 731: 122–131, 1996. 49. Vornov JJ, Thomas AG, and Jo D. Protective effects of extracellular acidosis and blockade of sodium/hydrogen ion exchange during recovery from metabolic inhibition in neuronal tissue culture. J Neurochem 67: 2379–2389, 1996. 50. Wakabayashi S, Pang T, Su X, and Shigekawa M. A novel topology model of the human Na(+)/H(+) exchanger isoform 1. J Biol Chem 275: 7942–7949, 2000. 51. Xia Y, Zhao P, Xue J, Gu XQ, Sun X, Yao H, and Haddad GG. Na+ channel expression and neuronal function in the Na+/H+ exchanger 1 null mutant mouse. J Neurophysiol 89: 229–236, 2003. 52. Xue J, Zhou D, Yao H, and Haddad GG. Role of transporters and ion channels in neuronal injury under hypoxia. Am J Physiol Regul Integr Comp Physiol 294: R451–R457, 2008. 53. Xue J, Douglas RM, Zhou D, Lim JY, Boron WF, and Haddad GG. Expression of Na+/ H+ and HCO3–-dependent transporters in Na+/H+ exchanger isoform 1 null mutant mouse brain. Neuroscience 122: 37–46, 2003. 54. Xue J, Zhou D, Yao H, Gavrialov O, McConnell MJ, Gelb BD, and Haddad GG. Novel functional interaction between Na+/H+ exchanger 1 and tyrosine phosphatase SHP-2. Am J Physiol Regul Integr Comp Physiol 292: R2406–R2416, 2007. 55. Yao H, Ma E, Gu XQ, and Haddad GG. Intracellular pH regulation of CA1 neurons in Na(+)/H(+) isoform 1 mutant mice. J Clin Invest 104: 637–645, 1999. 56. Yao H, Gu XQ, Douglas RM, and Haddad GG. Role of Na(+)/H(+) exchanger during O(2) deprivation in mouse CA1 neurons. Am J Physiol Cell Physiol 281: C1205–C1210, 2001. 57. Zhou D, Xue J, Gavrialov O, and Haddad GG. Na+/H+ exchanger 1 deficiency alters gene expression in mouse brain. Physiol Genomics 18: 331–339, 2004.

The Naþ/Kþ-ATPase as a Drug Target for Ischemic Stroke Melissa A. Gottron and Donald C. Lo

The Naþ/Kþ-ATPase (also known as the ‘‘sodium–potassium pump’’ or just ‘‘sodium pump’’) is a ubiquitous and critically important protein complex in the human body. In the brain, between 40 and 70% of all energy is spent maintaining its ionic transport activity [1, 2, 3]. This energy is used for the active exchange of cytosolic sodium for extracellular potassium in a 3:2 ratio, a process required for the maintenance of transmembrane ionic gradients for all mammalian cells, which in turn is essential for setting the cellular resting potential, regulating osmolarity, and powering the secondary transport of other important solutes like calcium (via the sodium–calcium exchanger) and protons (via the sodium–hydrogen exchanger). While these functions are especially critical for neuronal signaling (notably action potentials and synaptic transmission) and blood filtration in the kidney, all cells in the body ultimately depend on the Naþ/Kþ-ATPase to support their fundamental membrane processes. As the brain’s primary consumer of ATP, the sodium–potassium pump is particularly vulnerable to ATP depletion common in ischemic stroke. This vulnerability suggests that pharmacological inhibition of the Naþ/Kþ-ATPase would further compromise ATP-depleted neurons, yet there is accumulating evidence that inhibiting the sodium–potassium pump can actually provide neuroprotection in the context of ischemia as well as other traumatic and neurodegenerative conditions. The present chapter will thus review several aspects of the basic properties of the Naþ/Kþ-ATPase, mechanisms responsible for regulating Naþ/Kþ-ATPase function, and recent studies implicating the Naþ/Kþ-ATPase in ischemic stroke.

D.C. Lo (*) Center for Drug Discovery and Department of Neurobiology, Duke University Medical Center, Durham, NC, USA e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_8, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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1 Basic Properties 1.1 Gene Structure The Naþ/Kþ-ATPase was the first ionic transporter to be discovered and remains the focus of much important research more than 50 years later [4]. Like the hydrogen–potassium and calcium pumps, it is a member of the P-type ATPase protein family [5]. Members of this family earn their name from the gamma phosphate group that controls their activation; they become autophosphorylated by the ATP they hydrolyze, and this phosphorylation catalyzes conformational changes essential for pump function. Although all P-type ATPases share a common structure containing a heterodimeric protein core, significant functional differences arise from variations in the specific composition of this general structure. The Naþ/Kþ-ATPase consists of a heterodimeric core of a and b subunits that may be accompanied by a third g subunit [reviewed in 6, 7, 8]. These subunits are assembled in the endoplasmic reticulum before transport to the plasma membrane, and vectorial Golgi sorting as well as stabilization mechanisms play important roles in the pump’s targeted delivery [9, 10]. Both the a and b subunits are obligatory for the pump’s proper assembly and function [reviewed in 11]. The different subunits of the Naþ/Kþ-ATPase have distinct properties with respect to its overall functions (see Fig. 1). The a subunit of the Naþ/KþATPase is a 112 kDa protein with 10 membrane-spanning units whose N and C termini are intracellular. Because this subunit contains sites important for

Fig. 1 General structure of the sodium–potassium pump. The pump is a transmembrane protein composed of two major subunits that transport sodium and potassium between the cytoplasm and the extracellular milieu in a 3:2 ratio. While the a subunit is crucial for ATP binding and phosphorylation, ion occlusion, and catalytic activity, the b subunit plays a key role in pump maturation, stabilization, and transport. Both subunits are influenced by pharmacological modulation of the pump.

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ATP binding and phosphorylation as well as ion occlusion, it forms the catalytic core of the pump. The a subunit is also the primary binding site for many pharmacological agents that affect pump activity. Although their unique functional roles are still poorly understood, four isoforms of the a subunit are known to exist. The a1, a2, a3, and a4 isoforms differ in their kinetics, affinity for different ligands, and tissue and cellular distribution. Non-redundant functions for the different subunit isoforms are suggested by the findings that (1) although the a1 isoform is alone sufficient for the resting ion-transfer state, it cannot compensate for a lack of a2 during conditions of greater pump activity [12], (2) mutants lacking a2 and a3 isoforms show deficits in learning and motor activity as well as symptoms of anxiety [13], and (3) mutations of the gene encoding the a2 isoform have been associated with migraine disorders in humans [14]. The b subunit is a 35-kDa protein with a single, type II transmembranous region. This glycosylated subunit plays a key role in the maturation, transport, and stabilization of the pump as it migrates from the endoplasmic reticulum to the plasma membrane, and it also contains key residues necessary for potassium occlusion and ouabain binding [15, 16]. As for the a subunit, multiple b subunit isoforms (b1, b2, and b3) differentially influence Naþ/Kþ-ATPase affinity and stability. The g subunit, a member of the FXYD family, is a 6.5-kDa type I membrane protein associated with the Naþ/Kþ-ATPase in a tissue-specific manner. While not essential for basic pump expression or activity, the g subunit is typically present in a 1:1 molar ratio with the a and b subunits and interacts with the C-terminus of the a subunit in certain tissues [17, 18, 19]. This subunit modulates both the function and distribution of the pump by increasing its affinity for ATP [20, 21, 22]. Other members of the FXYD protein family that show a high degree of homology to the g subunit, like phospholemman [23], channelinducing factor [24], and mammary tumor-associated 8-kDa protein [25], have similarly been suggested to interact with the Naþ/Kþ-ATPase in various nonneural tissues [26].

1.2 Tissue/Cellular Distribution The importance of the Naþ/Kþ-ATPase to cellular function is highlighted by the fact that it is present in every cell of the body. Although a single cell may express more than one a or b isoform [27], the expression of different a and b isoforms is regulated in both a developmental and tissue-specific manner [28, 29, 30]. Tissue Distribution. The a1, a2, and a3 (but not a4) isoforms of the Naþ/KþATPase are expressed in the brain [31, 32]. While the a1 isoform is ubiquitously expressed in the central nervous system, the a2 and a3 isoforms show more restricted neural expression patterns [27, 28, 30, 33, 34]. The a2 isoform is widely

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expressed in neurons during development [28, 34] but becomes confined primarily to glia and a few types of neuron in the adult brain [27, 30]; in contrast, the a3 isoform shows primarily neuronal expression in the adult [27, 30]. Similar to the a subunit, the three b subunit isoforms are expressed to varying degrees in the brain. Pumps with the most common isoform, b1, are expressed ubiquitously throughout the body [33, 35]; those with b2 are largely found in excitable tissues like the heart and brain [36]; and those with b3 show only minimal neural expression [37]. The b isoforms show cell-specific mRNA [30] and protein expression [33, 38, 39], with the b1 and b2 isoforms expressed primarily in neurons and glia, respectively. Finally, the isoform composition of sodium–potassium pumps in different parts of the brain appears to be both complex and idiosyncratic [33, 38]. Neither preferential association of different a and b isoforms nor neurotransmitter-specific pairings has been observed. Due to their differing affinities for ligands, however, different isoform pairings confer unique functional properties to the cell types in which they are expressed. Subcellular Distribution. The a and b isoforms of the sodium–potassium pump can be targeted to specific subcellular locations within neurons and glia [39, 40, 41]. Although the distribution of a1 isoform is relatively uniform throughout the plasma membrane [42, 43], knockout studies have suggested that interactions between the a1 isoform and the IP3 receptor are functionally important [44]. In contrast, interactions with the cytoskeletal proteins ankyrin, spectrin, actin, and adducin preferentially target the a2 and a3 isoforms to specialized lipid rafts in the plasma membrane overlying the endoplasmic reticulum [42, 43, 45]. Because these locations are also enriched with the sodium–calcium exchanger, this places the a2 and a3 isoforms in an ideal situation to regulate the local concentrations of sodium and calcium in the cytosolic space between the plasma membrane and endoplasmic reticulum [46]. The subcellular localization of the pump thus permits specific functional control over cellular processes.

1.3 Biophysical Properties A well-accepted general description of the pump’s electrogenic activity is given by the Albers-Post scheme [47, 48, 49]. In the Albers-Post model, the pump alternates between two states (E1 and E2) that differ in both conformation and ion-binding affinity. Transitions between the E1 and E2 states are dependent on both voltage as well as local concentrations of sodium, potassium, and ATP, and these transitions are accompanied by the successive transport of sodium ions out of and potassium ions into the cell. In the Albers-Post model, the resting pump exists in the E1 conformation (Fig. 2). This conformation opens the Naþ/Kþ-ATPase’s cationbinding pocket to the intracellular space, which allows it to bind three

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intracellular sodium ions and one molecule of ATP. Hydrolysis of the bound ATP permits phosphorylation of pump and leads to the occlusion of the three sodium ions. A spontaneous conformational change then causes the pump to assume the E2 conformation, which opens toward the extracellular space. Because this conformation has a low affinity for sodium, the Naþ/Kþ-ATPase releases the occluded sodium ions into the extracellular space to complete the outward transport of sodium ions. The complementary inward transport of potassium begins with the exchange of sodium with potassium in the pump’s phosphorylated E2 conformation (Fig. 2). The extracellular binding of two potassium ions is followed by the spontaneous dephosphorylation of the E2 conformation, which results in the occlusion of the two potassium ions. Upon relaxation of the pump to the E1

Fig. 2 Albers-Post model of sequential ion transport. The Albers-Post model describes a potential mechanism for the exchange of sodium and potassium. The resting pump exists in the E1 state facing the cytoplasm. ATP as well as three sodium ions bind to the pump, and hydrolysis of the ATP permits phosphorylation of the pump and occlusion of the bound sodium ions. A spontaneous change in conformation allows the pump to assume the E2 conformation, which opens toward the extracellular space and releases the bound sodium into the extracellular space. The transport of potassium begins with the binding of two potassium ions to the pump’s phosphorylated E2 state. The spontaneous dephosphorylation of the E2 conformation occludes the two bound ions. When the pump relaxes to the E1 conformation, the potassium ions dissociate from their binding pocket and are released into the cytoplasm.

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conformation, the binding pocket containing the potassium ions opens into the cell. The binding of ATP inside the cell stimulates the release of the potassium ions and thus begins a new catalytic activity cycle. The Albers-Post scheme implies that a single a/b heterodimer is sufficient for activity of the Naþ/Kþ-ATPase, because ions are successively transported within a shared binding pocket. However, it should be noted that other schemes in which the Naþ/Kþ-ATPase functions as a higher-order oligomer have also been proposed [50, 51, 52]. Due to its role in the transport of sodium and potassium in a 3:2 ratio, the Naþ/Kþ-ATPase is electrogenic and works to hyperpolarize the cell. This active process requires both the consumption of O2 and hydrolysis of ATP [7, 53]. When either of these substrates is not available (as in ischemic stroke), pump failure produces a defined set of intracellular changes that includes the decrease of intracellular potassium, increase in intracellular sodium, and depolarization of the membrane [54]. These changes affect numerous intracellular processes, promulgating the damage caused by ATP scarcity.

1.4 Structural Features Although a great deal of structural similarity exists among the P-type ATPases, the Naþ/Kþ-ATPase is unique in that it is the only cation transporter with highaffinity binding sites for both potassium and ATP. While the molecular composition of the ion-binding pockets of the pump has spurred debate, numerous molecular biology studies were recently confirmed by the solution of a crystal structure describing the ion binding pockets of the Naþ/Kþ-ATPase [55]. The crystal structure identified a potassium-binding region between the fourth, fifth, and sixth transmembrane regions of the a subunit that simultaneously holds both potassium ions. Interestingly, two presumed sodium-binding sites overlap significantly with this potassium-binding pocket. Because such a shared binding pocket suggests sequential ion transport, this finding supports the Albers-Post sequential transport model described above. The location of the third sodiumbinding site remains unclear, but a distinct binding site may exist in the C-terminus of the a subunit. Other interactions of the Naþ/Kþ-ATPase’s functional domains have also been identified. The cytoplasmic loop between transmembrane segments four and five of the a subunit is essential for ATP hydrolysis, and regions near the seventh and ninth transmembrane segments are critical for interactions with the b and g subunits, respectively. These latter interactions differ sterically in that the b subunit is thought to completely cover the cytoplasmic loops between transmembrane segments 5–6 and 7–8 in the a subunit like a lid, whereas the g subunit is thought to intercalate between the a and b subunits. These findings are remarkably consistent with those implied by earlier molecular biology and modeling studies [16, 56, 57, 58].

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Finally, the pharmacological agents discussed in detail later in the chapter affect the pump through actions at specific locations. Cardiac steroids prevent pump activation by binding within a groove between the extracellular loops of the a subunit that permits its interaction with the b subunit, and mutagenesis studies have identified a number of amino acid residues critical for this interaction [59, 60, 61]. It is interesting and clinically relevant that the cellular ionic environment can influence the efficacy of pump blockade: high potassium levels lower the affinity of the Naþ/Kþ-ATPase for pharmacological blockers whereas sodium inhibits this effect [7]. Finally, although the binding sites for other drugs affecting pump activity are known to be distinct from those of the cardiac steroids, their precise locations remain elusive [62].

2 Regulation 2.1 Specific Pharmacological Inhibition Because of its pivotal role in cellular function, it is not surprising that the Naþ/ Kþ-ATPase is the target of a host of natural toxins derived from both plants and animals. These toxins typically stop the pump from functioning either by blocking its ion transport activities or by holding it ‘‘open’’ such that it resembles an open ionic channel. Both of these changes critically impair the maintenance of proper sodium and potassium gradients within the cell. Toxins that block the pump’s ion transport activities include ouabain and the cardenolide cardiac glycosides. As noted above, these toxins reversibly bind to the extracellular face of the Naþ/Kþ-ATPase in the phosphorylated E2 conformation. Cardiac glycosides consist of two main structural moieties: the aglycone moiety contains an R-group that defines the cardiac glycoside’s class and is important for its function, whereas the sugar moiety does not greatly influence biological activity but likely plays a role in the targeting and modulation of the drug. These drugs are toxic at high doses due to their blockade of pump activity, which leads to rundown of the sodium and potassium gradients, membrane depolarization, and secondary dysregulation of other ionic gradients. However, in lower doses the cardiac glycosides have been clinically effective and have been in therapeutic use since the eighteenth century in treating congestive heart failure [63](see Section 3.1). Palytoxin and sanguinarine, derived from the marine coral Palythoa toxica [64] and the plume poppies Macleaya cordata [65], respectively, are the primary members of a second class of Naþ/Kþ-ATPase toxins. Members of this class bind extracellularly to the phosphorylated E1 conformation of the Naþ/KþATPase to lock the pump in an open conformation [66]. This essentially converts the pump into a low-conductance ion channel, permitting a 10 pS current carried by potassium ions to flow out of the cell [64, 67]. The resulting rundown

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of sodium and potassium gradients across the cell membrane is extremely toxic and, indeed, palytoxin is perhaps the most toxic substance known – just 4 mg constitutes a lethal dose for humans.

2.2 Intrinsic Regulatory Mechanisms Because of its expression in every cell of the body, it is hardly surprising that multiple mechanisms exist to regulate the Naþ/Kþ-ATPase. These regulatory mechanisms are particularly important in cells, like neurons or those in nephrons, that undergo large fluctuations in ionic content due to neural activity or changes in diet and exercise. To ensure proper functioning, these cells must be able to titrate Naþ/Kþ-ATPase activity quickly and precisely in response to a variety of environmental challenges. That the Naþ/Kþ-ATPase operates at just 50% of its maximum capacity at rest implies that it can respond to a wide range of physiological stimuli by changing its activity level. The global activity of the Naþ/Kþ-ATPase within a cell can be changed either by (1) altering the number of pumps in the membrane or (2) altering the activity of individual pumps already existing within the membrane. Signaling pathways leading to both of these types of changes via modulation of Naþ/KþATPase transcription, transport, and functional kinetics have been described downstream of interactions with a variety of membrane proteins and hormones [reviewed in 68]. Regulation of pump number. Changes in the complement of Naþ/Kþ-ATPases in the membrane can arise through transcriptional, transport, and membrane recycling mechanisms. Transcriptional control of the Naþ/Kþ-ATPase was initially suggested by the observation that Naþ/Kþ-ATPase subunit transcription increases when the pump is blocked pharmacologically [69]; later work indicated that the 50 -flanking sequence of the gene encoding the a subunit contains both positive and negative transcriptional regulatory sites [70]. Cascades involving protein kinase A (PKA), extracellular signal-regulated kinase 1/2 (ERK1/2), and histone deacetylase have been implicated as critical regulators of a subunit transcription, whereas protein kinase C (PKC), c-Jun-N-terminal kinase (JNK), and p38 mitogen-activated protein kinase (MAPK) have been implicated as critical regulators of b subunit transcription [71]. Other brain-derived substances, such as dopamine and nitric oxide, also influence the transcription of the Naþ/Kþ-ATPase via activation of different signaling cascades [72, 73]. Finally, manipulations altering extracellular calcium concentrations affect the transcription and degradation rates of both the a and b subunits [69]. Changes in the transport of the Naþ/Kþ-ATPase from the ER also influence the complement of Naþ/Kþ-ATPase that accumulates in the plasma membrane. Many proteins are essential for proper targeting of the Naþ/Kþ-ATPase to specific domains within the plasma membrane, and cascades leading to the activation or inhibition of these proteins thereby influence Naþ/Kþ-ATPase activity. For example, arrestin and spinophilin compete to regulate the

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trafficking of the Naþ/Kþ-ATPase via interaction with its cytoplasmic loop [74]; activity promoting the dominance of one or another of these proteins therefore influences the complement of pumps at the membrane surface. Similarly, cascades that alter the presence or activity of the cytoskeletal proteins important for correct targeting of the pump to plasma membrane subdomains (e.g., ankyrin and spectrin) also influence overall Naþ/Kþ-ATPase levels and activity at the membrane. Finally, alterations in the rates at which the Naþ/Kþ-ATPase is either endoor exocytosed from the membrane rapidly affect the net complement of functional pumps available to transport sodium and potassium. Dopamine, RhoA, and the AMP kinases have all been shown to increase the rate of Naþ/KþATPase endocytosis from the membrane, thereby reducing its activity [68, 75, 76]. Other mechanisms of Naþ/Kþ-ATPase regulation, including that by IL-1, work by destabilizing the actin cytoskeleton and redistributing the pump via membrane recycling [77]. Regulation of pump activity. The second general class of Naþ/Kþ-ATPase regulation involves altering how effectively pumps already present in the membrane transport ions. This can be accomplished indirectly via restriction of the rate-limiting factors required for the pump’s activity or by direct interference with the pump via phosphorylation or protein binding. The most straightforward mechanisms of pump regulation involve varying the concentrations of its three major substrates: sodium, potassium, and ATP. The Naþ/Kþ-ATPase is normally most sensitive to changes in the concentration of intracellular sodium, because its half-maximal activation occurs at steady-state sodium levels [78]. This means that variations due to the activation of other sodium transporters greatly influence Naþ/Kþ-ATPase activity. In contrast, potassium is typically in excess under resting conditions and affects Naþ/Kþ-ATPase activity only to the extent that it competes with sodium for access to the pump [79]. Finally, while most cells contain adequate stores of ATP to power the Naþ/Kþ-ATPase, the binding of ATP is critical regulator of pump activity in cells with low ATP levels due to ischemia [80]. In addition to these changes in catalytic activity, Naþ/Kþ-ATPase stability is also heavily modulated by the cellular ionic environment [81]. The pump’s activity is modulated by direct interference with the pump itself as well. In fact, endogenous cardiotonic steroids have been discovered that are highly regulated under physiological and pathophysiological contexts and that inhibit the activity of the Naþ/Kþ-ATPase [reviewed in 82, 83]. The structure of these endogenous inhibitors is almost identical to that of the plant-derived Naþ/ Kþ-ATPase inhibitor ouabain. They are released from the adrenal gland and midbrain in response to a variety of environmental stimuli, and appear to interact with a specific hydrophobic binding partner that may influence their tissue distribution and inhibitory effects on ion transport [82]. Finally, modulation of Naþ/Kþ-ATPase activity can occur via phosphorylation/dephosphorylation by such proteins as PKA, PKC, PP2B, PKG, and others [reviewed in 68]. The idiosyncratic effects of phosphorylation on pump

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activity are both tissue- and species-specific, so that the activation of a particular signaling cascade may inhibit the pump in certain circumstances while increasing its activity in others. Environmental stimuli exert their effects through such signaling pathways in regulating the function of the Naþ/Kþ-ATPase. It is relevant for the current discussion of stroke that both hypoxia and hyperoxia can decrease the activity of the Naþ/Kþ-ATPase [84]. Although some such inhibition is likely downstream of ATP depletion due to improper oxygenation of mitochondria, proximal causes of decreased Naþ/Kþ-ATPase activity also include phosphorylation [85], free radical-induced damage [86], altered rates of endocytosis [85, 87], and changes in the complement of a and b subunit isoforms [88]. The physiological implications of such Naþ/Kþ-ATPase downregulation for stroke are discussed further in the following section.

3 Physiological Relevance 3.1 General Physiological Roles Ion transport-dependent function. The clinical relevance of the Naþ/Kþ-ATPase is highlighted by the use of cardiac glycoside drugs to increase heart contractility in the treatment of congestive heart failure. Although it is initially counterintuitive that blocking the Naþ/Kþ-ATPase would provide beneficial effects in heart failure, digoxin and other cardiac glycosides are thought to provide this benefit via the mechanism shown in the left panel of Fig. 3. This model centers on the role of the sodium–potassium pump in maintaining ionic gradients within the cell, and provides a general description of the ionic disturbances caused by Naþ/Kþ-ATPase inhibition. In the heart, inhibition of the Naþ/Kþ-ATPase leads to the accumulation of intracellular sodium, which decreases the steepness of the sodium gradient across the membranes of cardiac muscle cells. This reduced sodium gradient in turn limits the activity of the sodium–calcium exchanger, which normally taps the sodium gradient for energy in the extrusion of calcium. Each cardiac action potential is thus followed by elevated levels of residual intracellular calcium, the net effect of which is to strengthen successive heart contractions [89]. In this way, inhibition of the Naþ/Kþ-ATPase by cardiac glycoside drugs, notably digitoxin (Crystodigin) and digoxin (Lanoxin), produces beneficial effects in patients with congestive heart failure. Ion transport-independent signaling. There is increasing evidence that Naþ/ þ K -ATPase can also influence cellular processes by acting as part of a stable signalosome that regulates intracellular signaling pathways [90, 91]. A significant proportion of the sodium–potassium pumps expressed on the surface of certain cell types may not actively pump ions; rather, they may act as receptors for molecules that bind to the pump [92]. Such a signaling function would be thus independent of ionic transport activity and would work through the

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Fig. 3 Effects of pump blockade on ionic homeostasis and intracellular signaling cascades. Top panel: Under normal conditions, each cycle of the sodium–potassium pump transports three sodium ions out of and two potassium ions into the cell. This active, ATP-dependent process creates and maintains the normal sodium gradient across the cell plasma membrane, and secondly powers the extrusion of intracellular calcium via the sodium–calcium exchanger, which couples the inward flow of sodium down its concentration gradient with the outward transport of calcium. Left panel: Inhibition of the sodium–potassium pump degrades the sodium and potassium gradients, allowing a buildup of extracellular potassium and intracellular sodium. The increase in intracellular sodium reduces the sodium gradient that is the driving force for the sodium–calcium exchanger, thus limiting its capacity to extrude intracellular calcium. Right panel: Blockade of the sodium–potassium pump with low doses of ouabain also activates ion transport-independent signaling pathways. Src directly interacts with the pump to activate multiple signaling cascades. The downstream effects of these cascades include activation of the NFkB and AP1 transcription factors, and such signaling through the Naþ/Kþ-ATPase has been implicated in growth, proliferation, apoptosis, and ischemic preconditioning.

activation of second messenger cascades [93]. Low concentrations of pumpblocking compounds like ouabain activate these receptors in an isoformspecific manner because the binding affinities of the a2 and a3 isoforms for ouabain are much higher than that of the a1 and low doses of ouabain specifically inhibit just a subset of the a isoforms [94]. A simplified schematic of signaling downstream of the sodium–potassium pump is given in the right panel of Fig. 3. The binding of pump-blocking molecules like endogenous cardiosteroids or low doses of ouabain activates Src. Src directly interacts with the Naþ/Kþ-ATPase, and the activation of this molecule initiates a cascade involving a number of other signaling

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intermediates, culminating in the downstream activation of the MAPK cascade, PLC, and PKC. While the entire complement of cellular changes resulting from Naþ/Kþ-ATPase signaling remains unknown, signaling downstream of the Naþ/Kþ-ATPase initiates a host of cellular events including alteration of a subunit localization in the plasma membrane in an isoform-specific manner [95]. In turn, the combination of dysregulated intracellular calcium and Naþ/KþATPase signalosome activity induced by pump blockade influences cellular transcription via activation of transcription factors like NFkB and AP-1. Such ion transport-independent Naþ/Kþ-ATPase signaling has been shown to influence physiologically relevant processes such as hypertrophic growth in cardiac myocytes [96, 97], proliferation in smooth muscle [98], and apoptosis in malignant cells [99, 100]. Finally, it is relevant for the present discussion of stroke that MAP kinase activation downstream of Naþ/Kþ-ATPase signaling mediates a form of neuronal pre-ischemic conditioning that protects cells later exposed to hypoxic conditions [101].

3.2 Pathological Relevance to Stroke Ischemic stroke, which is defined by the loss of blood and oxygen flow to a region in the brain, profoundly affects the activity of the sodium–potassium pump. In rat models, ischemia causes ATP levels to drop precipitously within minutes due to the inhibition of mitochondrial respiration [102]. Because the Naþ/Kþ-ATPase uses such a large proportion of the cell’s energy store, its activity becomes especially vulnerable under these conditions. Both Naþ/Kþ-ATPase levels and activity are downregulated in model systems of focal ischemia and traumatic brain injury [103, 104, 105, 106], and the reduction of Naþ/Kþ-ATPase activity is partly responsible for the increased sodium influx observed in the hippocampus during and after anoxia [107]. Naþ/Kþ-ATPase activity is also downregulated in a variety of other neuropathologic and apoptotic contexts clearly associated with compromised cellular health [108, 109, 110]. Surprisingly, however, evidence also suggests that inhibition of the Naþ/KþATPase can be neuroprotective. Low concentrations of ouabain that preferentially inhibit the a2 and a3, but not a1, isoforms are protective against hypoxiainduced cell death in cortical [111], striatal [112], and cerebellar [113] neurons. Such inhibition of the Naþ/Kþ-ATPase promotes neuron survival as well as the activation of factors like PI3K and pAkt that are important for somal health and neurite extension [114, 115, 116, 117, 118]. Independently, in a blinded screen of several thousand known drug-like compounds (including all FDA-approved drugs), we identified a cardiac glycoside as one of the two most efficacious compounds in providing neuroprotection in a cortical brain-slice explant model for focal ischemia (Fig. 4) [119]. This compound, neriifolin, is structurally closely related to the FDA-approved cardiac glycosides digoxin and digitoxin, and indeed these drugs were

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Fig. 4 The cardiac glycoside neriifolin is neuroprotective in models of focal ischemia. a A highthroughput biology screen was developed for ischemic stroke. In this screen, biolistically transfected cortical brain-slice explants were subjected to transient oxygen-glucose deprivation (OGD), causing the degeneration of YFP-transfected neurons over the course of 1–3 days (compare control brain slice (left) to OGD brain slice (right) at 3 days post-OGD). b This brain-slice model for ischemic stroke resulted in the identification of the cardiac glycoside neriifolin as a potent neuroprotective agent. c Summary data from a portion of this screen are shown; the solid line at a value of 1 on the y-axis shows the OGD negative control level whereas the solid line at a value of 28 shows the positive control level for parallel brain slices co-transfected with the antiapoptotic gene Bcl-xL. The dotted line depicts 5 SD above the negative OGD control. While several compounds rescued against OGD-induced neuronal death above this threshold level, neriifolin afforded the greatest neuroprotection in this run (red arrow). d Neriifolin continued to provide neuroprotection even when administered 6 or more hours after OGD. e Neriifolin provides a dose-dependent reduction of infarct volume in the adult rat middle cerebral artery occlusion (MCAO) model of focal ischemia. *P < 0.07, **P < 0.01. Data adapted from [119].

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subsequently also shown to provide neuroprotection in the brain-slice stroke model but with lower potency. Importantly, we found that neriifolin could provide quantitative neuroprotection even when administered 6 or more hours after ischemic injury, corresponding to the therapeutic time window in which stroke victims are first likely to reach the emergency room [120]. Finally, the neuroprotective action of neriifolin was confirmed in two independent in vivo stroke models. It is possible to reconcile these apparently conflicting sets of data by considering the hypothesis that ATP conservation during ischemic conditions can serve to ameliorate cellular damage and/death consequent to mitochondrial collapse. Reducing demand for ATP by inhibiting the brain’s greatest consumer of ATP may thus be able to promote neuronal survival even under adverse conditions such as ischemia. Such a reduction of metabolic demand under conditions of stress is not a novel concept and is consistent with the ‘‘channel lock’’ hypothesis proposed by Hochachka in the 1980s [121]. In fact, reduction in brain energy consumption forms the underpinnings of several adaptive neuroprotective strategies found in the animal kingdom: gall flies survive the a-energetic conditions of the winter hibernation period by reducing Naþ/Kþ-ATPase activity [122]; snails entering a hypometabolic state termed estivation reduce Naþ/ Kþ-ATPase ATP consumption through phosphorylation of the a subunit [123]; sea turtles reduce energy consumption by the Naþ/Kþ-ATPase to survive prolonged underwater hypoxia [124]; ground squirrels show phosphorylation-induced reductions in Naþ/Kþ-ATPase activity during winter hibernation [125]. In addition, that hypoxia in rats stimulates a large and rapid increase in the production of an endogenous ouabain analog to reduce Naþ/Kþ-ATPase activity is another example of adaptive ATP conservation under hypoxic conditions [126]. Finally, a principal mechanism underlying therapeutic hypothermia used in the clinic is the conservation of ATP, much of which would otherwise be used to power the Naþ/Kþ-ATPase [127]. In addition to ATP conservation, blockade of the sodium–potassium pump may also provide neuroprotection in ischemia through modulating intracellular calcium levels just as the cardiac glycosides do in the heart contraction cycle. Although high levels of calcium are clearly harmful for neuronal viability [100], abnormally low levels of calcium can also lead to neuronal death [128, 129]. The ‘‘calcium set-point’’ hypothesis for neuronal trophic dependence thus suggests that elevation of cytosolic calcium is critical for neuronal survival when normal trophic support is not available – as in the case of ischemic stroke. Consistent with this hypothesis, hippocampal neurons exposed to transient ischemia experience lower than normal levels of intracellular calcium and NMDA receptors for 1–3 days subsequent to the ischemic insult [130], and the administration of NMDA receptor agonists to neurons during this period paradoxically attenuates neurologic deficits and restores cognitive performance [131]. Calcium starvation and its consequent apoptosis may even be the predominant cause of cell death in the ischemic penumbra, particularly at later time intervals

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after stroke [132]. Therefore, elevating calcium levels in neurons exposed to ischemic events via suppression of sodium–calcium exchange as a secondary consequence of Naþ/Kþ-ATPase inhibition may be an important mechanism for providing protection against the delayed neuronal death following stroke. In further support of Naþ/Kþ-ATPase inhibition providing neuroprotection to mammalian neurons under stress, low doses of ouabain have been found to increase the survival of both cultured retinal ganglion cells and mesencephalic dopaminergic neurons [133, 134], and a non-cardiac glycoside compound with Naþ/Kþ-ATPase inhibitory properties has recently been found also to provide neuroprotection in an in vivo stroke model [135]. Finally, a screen of the NINDS Custom Collection for new drug targets and candidates for the treatment of spinobulbar muscular atrophy showed that cardiac glycosides inhibit the polyglutamine-dependent activation of caspase-3 and thus provided neuroprotection against polyglutamine-mediated cytotoxicity [136].

3.3 Therapeutic Perspectives Inhibition of the Naþ/Kþ-ATPase thus offers novel neuroprotective mechanisms for providing therapeutic benefit after stroke. This exciting possibility, together with the availability of FDA-approved Naþ/Kþ-ATPase inhibiting drugs, however, is tempered by some points of potential concern. First, the cardiac glycoside drug digoxin, which is in current clinical use, has limited access to the CNS under normal conditions. Although numerous examples of CNS-related side effects of digitoxin treatment argues for some degree of CNS penetration [reviewed in 137], penetration of digoxin into the CNS is limited by the efflux transporter P-glycoprotein in the blood–brain barrier (BBB) endothelium [138]. Targeted deletion of the mdr1a gene in mice lacking this transporter results in substantially higher levels of digoxin penetration into the brain [139, 140], and inhibitors of the transporter (e.g., quinidine and Pluronic p85) also increase digoxin penetration across the BBB [141, 142, 143, 144]. The development of tractable drugs inhibiting the P-glycoprotein transporter therefore may offer a pathway for increasing the penetration of drugs blocking the sodium–potassium pump into the CNS [145]. A second concern over the therapeutic use of Naþ/Kþ-ATPase inhibitors in stroke centers on a problem common to all drugs intended as stroke treatments. Because the average stroke patient does not arrive at the hospital until 6 or more hours after stroke onset [120], any drugs developed for the treatment of stroke must retain their therapeutic effects even after a significant delay of administration. In fact, the beneficial effects of the ‘‘clotbuster’’ tissue plasminogen activator (tPA), the only drug treatment currently FDA-approved for stroke, decline rapidly in the hours after stroke onset and tPA is not indicated for use after 3-h post-stroke onset. This hurdle has produced significant problems for the development of effective therapeutic agents for this disease, and one

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promising aspect of our studies of neriifolin described above is that it retains neuroprotective effects even 6 h or more after ischemic injury in the context of brain tissue explants. Finally, the narrow therapeutic window with respect to dose for cardiac glycosides and other Naþ/Kþ-ATPase inhibitors presents a challenge for clinical management. The beneficial effects of energy conservation due to low levels of Naþ/Kþ-ATPase inhibition could quickly become detrimental in terms of excessive rundown of cellular ionic gradients, particularly in off-target organs such as the heart. In this context, it will be critical, as described above, to develop strategies for increasing the penetration of Naþ/Kþ-ATPase inhibitors into the CNS and ideally to develop such inhibitors with preferential action on neural tissues.

4 Summary As the brain’s main consumer of ATP, the Naþ/Kþ-ATPase is a critical determinant of normal neuronal function in the brain. In addition to its direct role in ionic transport, the sodium–potassium pump is crucial for maintaining key cellular properties (e.g., osmolarity, pH, and membrane potential), modulating important signaling cascades, and controlling intracellular calcium levels. Each of these functions is threatened when Naþ/Kþ-ATPase activity slows in ischemia, but recent evidence from a range of studies suggests that pharmacological inhibition of the pump’s energy requirements may in fact promote neuron survival under ischemic conditions. Based on this evidence, the Naþ/KþATPase clearly merits additional study as a potential drug target for ischemic stroke.

References 1. Clausen T, Van Hardeveld C, and Everts ME. Significance of cation transport in control of energy metabolism and thermogenesis. Physiol Rev 71: 733–774, 1991. 2. Friberg H and Wieloch T. Mitochondrial permeability transition in acute neurodegeneration. Biochimie 84: 241–250, 2002. 3. Astrup J. Energy-requiring cell functions in the ischemic brain. Their critical supply and possible inhibition in protective therapy. J Neurosurg 56: 482–497, 1982. 4. Skou JC. The influence of some cations on an adenosine triphosphatase from peripheral nerves. Biochim Biophys Acta 23: 394–401, 1957. 5. Skou JC. The influence of some cations on an adenosine triphosphatase from peripheral nerves. 1957. Biochim Biophys Acta 1000: 439–446, 1989. 6. Kaplan JH. Biochemistry of Na,K-ATPase. Annu Rev Biochem 71: 511–535, 2002. 7. Scheiner-Bobis G. The sodium pump. Its molecular properties and mechanics of ion transport. Eur J Biochem 269: 2424–2433, 2002. 8. Lingrel JB, Young RM, and Shull MM. Multiple forms of the Na,K-ATPase: their genes and tissue specific expression. Prog Clin Biol Res 268B: 105–112, 1988.

The Naþ/Kþ-ATPase as a Drug Target for Ischemic Stroke

145

9. Zurzolo C and Rodriguez-Boulan E. Delivery of Na+,K(+)-ATPase in polarized epithelial cells. Science 260: 550–552; author reply 554–556, 1993. 10. Gottardi CJ and Caplan MJ. Delivery of Na+,K(+)-ATPase in polarized epithelial cells. Science 260: 552–554; author reply 554–6, 1993. 11. Fambrough DM, Lemas MV, Hamrick M, Emerick M, Renaud KJ, Inman EM, Hwang B, and Takeyasu K. Analysis of subunit assembly of the Na-K-ATPase. Am J Physiol 266: C579–C589, 1994. 12. Radzyukevich TL, Moseley AE, Shelly DA, Redden GA, Behbehani MM, Lingrel JB, Paul RJ, and Heiny JA. The Na(+)-K(+)-ATPase alpha2-subunit isoform modulates contractility in the perinatal mouse diaphragm. Am J Physiol Cell Physiol 287: C1300–C1310, 2004. 13. Moseley AE, Williams MT, Schaefer TL, Bohanan CS, Neumann JC, Behbehani MM, Vorhees CV, and Lingrel JB. Deficiency in Na,K-ATPase alpha isoform genes alters spatial learning, motor activity, and anxiety in mice. J Neurosci 27: 616–626, 2007. 14. Castro MJ, Nunes B, de Vries B, Lemos C, Vanmolkot KR, van den Heuvel JJ, Temudo T, Barros J, Sequeiros J, Frants RR, Koenderink JB, Pereira-Monteiro JM, and van den Maagdenberg AM. Two novel functional mutations in the Na+,K+-ATPase alpha2-subunit ATP1A2 gene in patients with familial hemiplegic migraine and associated neurological phenotypes. Clin Genet 73: 37–43, 2008. 15. Geering K. The functional role of beta subunits in oligomeric P-type ATPases. J Bioenerg Biomembr 33: 425–438, 2001. 16. Lutsenko S and Kaplan JH. An essential role for the extracellular domain of the Na,KATPase beta-subunit in cation occlusion. Biochemistry 32: 6737–6743, 1993. 17. Hardwicke PM and Freytag JW. A proteolipid associated with Na,K-ATPase is not essential for ATPase activity. Biochem Biophys Res Commun 102: 250–257, 1981. 18. Beguin P, Wang X, Firsov D, Puoti A, Claeys D, Horisberger JD, and Geering K. The gamma subunit is a specific component of the Na,K-ATPase and modulates its transport function. Embo J 16: 4250–4260, 1997. 19. Scheiner-Bobis G and Farley RA. Subunit requirements for expression of functional sodium pumps in yeast cells. Biochim Biophys Acta 1193: 226–234, 1994. 20. Sweadner KJ, Arystarkhova E, Donnet C, and Wetzel RK. FXYD proteins as regulators of the Na,K-ATPase in the kidney. Ann N Y Acad Sci 986: 382–387, 2003. 21. Geering K, Beguin P, Garty H, Karlish S, Fuzesi M, Horisberger JD, and Crambert G. FXYD proteins: new tissue- and isoform-specific regulators of Na,K-ATPase. Ann N Y Acad Sci 986: 388–394, 2003. 22. Therien AG, Karlish SJ, and Blostein R. Expression and functional role of the gamma subunit of the Na, K-ATPase in mammalian cells. J Biol Chem 274: 12252–12256, 1999. 23. Palmer CJ, Scott BT, and Jones LR. Purification and complete sequence determination of the major plasma membrane substrate for cAMP-dependent protein kinase and protein kinase C in myocardium. J Biol Chem 266: 11126–11130, 1991. 24. Attali B, Latter H, Rachamim N, and Garty H. A corticosteroid-induced gene expressing an ‘‘IsK-like’’ K+ channel activity in Xenopus oocytes. Proc Natl Acad Sci USA 92: 6092–6096, 1995. 25. Morrison BW and Leder P. Neu and ras initiate murine mammary tumors that share genetic markers generally absent in c-myc and int-2-initiated tumors. Oncogene 9: 3417–3426, 1994. 26. Mercer RW, Biemesderfer D, Bliss DP Jr., Collins JH, Forbush B 3rd. Molecular cloning and immunological characterization of the gamma polypeptide, a small protein associated with the Na,K-ATPase. J Cell Biol 121: 579–586, 1993. 27. McGrail KM, Phillips JM, and Sweadner KJ. Immunofluorescent localization of three Na,K-ATPase isozymes in the rat central nervous system: both neurons and glia can express more than one Na,K-ATPase. J Neurosci 11: 381–391, 1991.

146

M.A. Gottron and D.C. Lo

28. Herrera VL, Cova T, Sassoon D, and Ruiz-Opazo N. Developmental cell-specific regulation of Na(+)-K(+)-ATPase alpha 1-, alpha 2-, and alpha 3-isoform gene expression. Am J Physiol 266: C1301–C1312, 1994. 29. Lu XP and Leffert HL. Induction of sodium pump beta 1-subunit mRNA expression during hepatocellular growth transitions in vitro and in vivo. J Biol Chem 266: 9276–9284, 1991. 30. Watts AG, Sanchez-Watts G, Emanuel JR, and Levenson R. Cell-specific expression of mRNAs encoding Na+,K(+)-ATPase alpha- and beta-subunit isoforms within the rat central nervous system. Proc Natl Acad Sci USA 88: 7425–7429, 1991. 31. Shull GE, Greeb J, and Lingrel JB. Molecular cloning of three distinct forms of the Na+,K+-ATPase alpha-subunit from rat brain. Biochemistry 25: 8125–8132, 1986. 32. Shamraj OI and Lingrel JB. A putative fourth Na+,K(+)-ATPase alpha-subunit gene is expressed in testis. Proc Natl Acad Sci USA 91: 12952–12956, 1994. 33. Peng L, Martin-Vasallo P, and Sweadner KJ. Isoforms of Na,K-ATPase alpha and beta subunits in the rat cerebellum and in granule cell cultures. J Neurosci 17: 3488–3502, 1997. 34. Moseley AE, Lieske SP, Wetzel RK, James PF, He S, Shelly DA, Paul RJ, Boivin GP, Witte DP, Ramirez JM, Sweadner KJ, and Lingrel JB. The Na,K-ATPase alpha 2 isoform is expressed in neurons, and its absence disrupts neuronal activity in newborn mice. J Biol Chem 278: 5317–5324, 2003. 35. Emanuel JR, Garetz S, Stone L, and Levenson R. Differential expression of Na+,K+ATPase alpha- and beta-subunit mRNAs in rat tissues and cell lines. Proc Natl Acad Sci USA 84: 9030–9034, 1987. 36. Shyjan AW, Gottardi C, and Levenson R. The Na,K-ATPase beta 2 subunit is expressed in rat brain and copurifies with Na,K-ATPase activity. J Biol Chem 265: 5166–5169, 1990. 37. Arystarkhova E and Sweadner KJ. Tissue-specific expression of the Na,K-ATPase beta3 subunit. The presence of beta3 in lung and liver addresses the problem of the missing subunit. J Biol Chem 272: 22405–22408, 1997. 38. Cameron R, Klein L, Shyjan AW, Rakic P, and Levenson R. Neurons and astroglia express distinct subsets of Na,K-ATPase alpha and beta subunits. Brain Res Mol Brain Res 21: 333–343, 1994. 39. Lecuona E, Luquin S, Avila J, Garcia-Segura LM, and Martin-Vasallo P. Expression of the beta 1 and beta 2(AMOG) subunits of the Na,K-ATPase in neural tissues: cellular and developmental distribution patterns. Brain Res Bull 40: 167–174, 1996. 40. Brines ML and Robbins RJ. Cell-type specific expression of Na+, K(+)-ATPase catalytic subunits in cultured neurons and glia: evidence for polarized distribution in neurons. Brain Res 631: 1–11, 1993. 41. Wetzel RK, Arystarkhova E, and Sweadner KJ. Cellular and subcellular specification of Na,K-ATPase alpha and beta isoforms in the postnatal development of mouse retina. J Neurosci 19: 9878–9889, 1999. 42. Juhaszova M and Blaustein MP. Distinct distribution of different Na+ pump alpha subunit isoforms in plasmalemma. Physiological implications. Ann N Y Acad Sci 834: 524–536, 1997. 43. Juhaszova M and Blaustein MP. Na+ pump low and high ouabain affinity alpha subunit isoforms are differently distributed in cells. Proc Natl Acad Sci USA 94: 1800–1805, 1997. 44. Chen Y, Cai T, Yang C, Turner DA, Giovannucci DR, and Xie Z. Regulation of inositol 1,4,5-trisphosphate receptor-mediated calcium release by the Na/K-ATPase in cultured renal epithelial cells. J Biol Chem 283: 1128–1136, 2008. 45. Dunbar LA and Caplan MJ. Ion pumps in polarized cells: sorting and regulation of the Na+, K+- and H+, K+-ATPases. J Biol Chem 276: 29617–29620, 2001. 46. Golovina V, Song H, James P, Lingrel J, and Blaustein M. Regulation of Ca2+ signaling by Na+ pump alpha-2 subunit expression. Ann N Y Acad Sci 986: 509–513, 2003. 47. Charnock JS and Post RL. Evidence of the mechanism of ouabain inhibition of cationactivated adenosine triphosphate. Nature 199: 910–911, 1963.

The Naþ/Kþ-ATPase as a Drug Target for Ischemic Stroke

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48. Albers RW, Fahn S, and Koval GJ. The role of sodium ions in the activation of electrophorus electric organ adenosine triphosphatase. Proc Natl Acad Sci USA 50: 474–481, 1963. 49. Glynn IM. Annual review prize lecture. ‘All hands to the sodium pump’. J Physiol 462: 1–30, 1993. 50. Clarke RJ and Kane DJ. Two gears of pumping by the sodium pump. Biophys J 93: 4187–4196, 2007. 51. Laughery M, Todd M, and Kaplan JH. Oligomerization of the Na,K-ATPase in cell membranes. J Biol Chem 279: 36339–36348, 2004. 52. Geering K, Beggah A, Good P, Girardet S, Roy S, Schaer D, and Jaunin P. Oligomerization and maturation of Na,K-ATPase: functional interaction of the cytoplasmic NH2 terminus of the beta subunit with the alpha subunit. J Cell Biol 133: 1193–1204, 1996. 53. Jorgensen PL, Hakansson KO, and Karlish SJ. Structure and mechanism of Na,KATPase: functional sites and their interactions. Annu Rev Physiol 65: 817–849, 2003. 54. Jones SW. On the resting potential of isolated frog sympathetic neurons. Neuron 3: 153–161, 1989. 55. Morth JP, Pedersen BP, Toustrup-Jensen MS, Sorensen TL, Petersen J, Andersen JP, Vilsen B, and Nissen P. Crystal structure of the sodium-potassium pump. Nature 450: 1043–1049, 2007. 56. Li C, Grosdidier A, Crambert G, Horisberger JD, Michielin O, and Geering K. Structural and functional interaction sites between Na,K-ATPase and FXYD proteins. J Biol Chem 279: 38895–38902, 2004. 57. Ogawa H and Toyoshima C. Homology modeling of the cation binding sites of Na+K+ATPase. Proc Natl Acad Sci USA 99: 15977–15982, 2002. 58. Vilsen B, Ramlov D, and Andersen JP. Functional consequences of mutations in the transmembrane core region for cation translocation and energy transduction in the Na+,K(+)-ATPase and the SR Ca(2+)-ATPase. Ann N Y Acad Sci 834: 297–309, 1997. 59. Keenan SM, DeLisle RK, Welsh WJ, Paula S, Ball WJ Jr. Elucidation of the Na+, K+ATPase digitalis binding site. J Mol Graph Model 23: 465–475, 2005. 60. Lowndes JM, Hokin-Neaverson M, and Ruoho AE. Photoaffinity labeling of (Na+K+)ATPase with [125I]iodoazidocymarin. J Biol Chem 259: 10533–10538, 1984. 61. Rogers TB and Lazdunski M. Photoaffinity labelling of a small protein component of a purified (Na+-K+)ATPase. FEBS Lett 98: 373–376, 1979. 62. Vale-Gonzalez C, Pazos MJ, Alfonso A, Vieytes MR, and Botana LM. Study of the neuronal effects of ouabain and palytoxin and their binding to Na,K-ATPases using an optical biosensor. Toxicon 50: 541–552, 2007. 63. Withering W. An Account of the Floxglove and Some of Its Medical Uses with Practical Remarks on Dropsy and Other Diseases. Birmingham, UK: M. Swinney, 1785. 64. Hirsh JK and Wu CH. Palytoxin-induced single-channel currents from the sodium pump synthesized by in vitro expression. Toxicon 35: 169–176, 1997. 65. Scheiner-Bobis G. Sanguinarine induces K+ outflow from yeast cells expressing mammalian sodium pumps. Naunyn Schmiedebergs Arch Pharmacol 363: 203–208, 2001. 66. Habermann E. Palytoxin acts through Na+,K+-ATPase. Toxicon 27: 1171–1187, 1989. 67. Takeuchi A, Reyes N, Artigas P, Gadsby DC. The ion pathway through the opened Na(+),K(+)-ATPase pump. Nature 456: 413–416, 2008. 68. Therien AG and Blostein R. Mechanisms of sodium pump regulation. Am J Physiol Cell Physiol 279: C541–C566, 2000. 69. Rayson BM. [Ca2+]i regulates transcription rate of the Na+/K(+)-ATPase alpha 1 subunit. J Biol Chem 266: 21335–21338, 1991. 70. Pathak BG, Neumann JC, Croyle ML, and Lingrel JB. The presence of both negative and positive elements in the 50 -flanking sequence of the rat Na,K-ATPase alpha 3 subunit gene are required for brain expression in transgenic mice. Nucleic Acids Res 22: 4748–4755, 1994.

148

M.A. Gottron and D.C. Lo

71. Wang G, Kawakami K, and Gick G. Divergent signaling pathways mediate induction of Na,K-ATPase alpha1 and beta1 subunit gene transcription by low potassium. Mol Cell Biochem 294: 73–85, 2007. 72. Guerrero C, Lecuona E, Pesce L, Ridge KM, and Sznajder JI. Dopamine regulates Na-KATPase in alveolar epithelial cells via MAPK-ERK-dependent mechanisms. Am J Physiol Lung Cell Mol Physiol 281: L79–L85, 2001. 73. Kone BC and Higham S. Nitric oxide inhibits transcription of the Na+-K+-ATPase alpha1-subunit gene in an MTAL cell line. Am J Physiol 276: F614–F621, 1999. 74. Kimura T, Allen PB, Nairn AC, and Caplan MJ. Arrestins and spinophilin competitively regulate Na+,K+-ATPase trafficking through association with a large cytoplasmic loop of the Na+,K+-ATPase. Mol Biol Cell 18: 4508–4518, 2007. 75. Dada LA, Novoa E, Lecuona E, Sun H, and Sznajder JI. Role of the small GTPase RhoA in the hypoxia-induced decrease of plasma membrane Na,K-ATPase in A549 cells. J Cell Sci 120: 2214–2222, 2007. 76. Vadasz I, Dada LA, Briva A, Trejo HE, Welch LC, Chen J, Toth PT, Lecuona E, Witters LA, Schumacker PT, Chandel NS, Seeger W, and Sznajder JI. AMP-activated protein kinase regulates CO2-induced alveolar epithelial dysfunction in rats and human cells by promoting Na,K-ATPase endocytosis. J Clin Invest 118: 752–762, 2008. 77. Namekata K, Harada C, Kohyama K, Matsumoto Y, and Harada T. Interleukin-1 stimulates glutamate uptake in glial cells by accelerating membrane trafficking of Na+/K+-ATPase via actin depolymerization. Mol Cell Biol 28: 3273–3280, 2008. 78. Soltoff SP and Mandel LJ. Active ion transport in the renal proximal tubule. II. Ionic dependence of the Na pump. J Gen Physiol 84: 623–642, 1984. 79. Garay RP and Garrahan PJ. The interaction of sodium and potassium with the sodium pump in red cells. J Physiol 231: 297–325, 1973. 80. Soltoff SP and Mandel LJ. Active ion transport in the renal proximal tubule. III. The ATP dependence of the Na pump. J Gen Physiol 84: 643–662, 1984. 81. Fodor E, Fedosova NU, Ferencz C, Marsh D, Pali T, and Esmann M. Stabilization of Na,K-ATPase by ionic interactions. Biochim Biophys Acta 1778: 835–843, 2008. 82. Schoner W, Bauer N, Muller-Ehmsen J, Kramer U, Hambarchian N, Schwinger R, Moeller H, Kost H, Weitkamp C, Schweitzer T, Kirch U, Neu H, and Grunbaum EG. Ouabain as a mammalian hormone. Ann N Y Acad Sci 986: 678–684, 2003. 83. Schoner W. Endogenous cardiac glycosides, a new class of steroid hormones. Eur J Biochem 269: 2440–2448, 2002. 84. Petrushanko IY, Bogdanov NB, Lapina N, Boldyrev AA, Gassmann M, and Bogdanova AY. Oxygen-induced regulation of Na/K ATPase in cerebellar granule cells. J Gen Physiol 130: 389–398, 2007. 85. Dada LA, Welch LC, Zhou G, Ben-Saadon R, Ciechanover A, and Sznajder JI. Phosphorylation and ubiquitination are necessary for Na,K-ATPase endocytosis during hypoxia. Cell Signal 19: 1893–1898, 2007. 86. Comellas AP, Dada LA, Lecuona E, Pesce LM, Chandel NS, Quesada N, Budinger GR, Strous GJ, Ciechanover A, Sznajder JI. Hypoxia-mediated degradation of Na,K-ATPase via mitochondrial reactive oxygen species and the ubiquitin-conjugating system. Circ Res 98: 1314–1322, 2006. 87. Dada LA, Chandel NS, Ridge KM, Pedemonte C, Bertorello AM, and Sznajder JI. Hypoxia-induced endocytosis of Na,K-ATPase in alveolar epithelial cells is mediated by mitochondrial reactive oxygen species and PKC-zeta. J Clin Invest 111: 1057–1064, 2003. 88. Ostadal P, Elmoselhi AB, Zdobnicka I, Lukas A, Chapman D, and Dhalla NS. Ischemiareperfusion alters gene expression of Na+-K+ ATPase isoforms in rat heart. Biochem Biophys Res Commun 306: 457–462, 2003. 89. Hardman JG and Limbird LL. Goodman & Gilman’s The Pharmacological Basis of Therapeutics, 9th Ed. New York: McGraw-Hill, 1996.

The Naþ/Kþ-ATPase as a Drug Target for Ischemic Stroke

149

90. Xie Z and Askari A. Na(+)/K(+)-ATPase as a signal transducer. Eur J Biochem 269: 2434–2439, 2002. 91. Xie Z and Cai T. Na+-K+-ATPase-mediated signal transduction: from protein interaction to cellular function. Mol Interv 3: 157–168, 2003. 92. Liang M, Tian J, Liu L, Pierre S, Liu J, Shapiro J, and Xie ZJ. Identification of a pool of non-pumping Na/K-ATPase. J Biol Chem 282: 10585–10593, 2007. 93. Liu J, Tian J, Haas M, Shapiro JI, Askari A, Xie Z. Ouabain interaction with cardiac Na+/K+-ATPase initiates signal cascades independent of changes in intracellular Na+ and Ca2+ concentrations. J Biol Chem 275: 27838–27844, 2000. 94. Monteith GR and Blaustein MP. Different effects of low and high dose cardiotonic steroids on cytosolic calcium in spontaneously active hippocampal neurons and in co-cultured glia. Brain Res 795: 325–340, 1998. 95. Taguchi K, Kumanogoh H, Nakamura S, and Maekawa S. Ouabain-induced isoformspecific localization change of the Na+, K+-ATPase alpha subunit in the synaptic plasma membrane of rat brain. Neurosci Lett 413: 42–45, 2007. 96. Kometiani P, Li J, Gnudi L, Kahn BB, Askari A, and Xie Z. Multiple signal transduction pathways link Na+/K+-ATPase to growth-related genes in cardiac myocytes. The roles of Ras and mitogen-activated protein kinases. J Biol Chem 273: 15249–15256, 1998. 97. Xie Z, Kometiani P, Liu J, Li J, Shapiro JI, and Askari A. Intracellular reactive oxygen species mediate the linkage of Na+/K+-ATPase to hypertrophy and its marker genes in cardiac myocytes. J Biol Chem 274: 19323–19328, 1999. 98. Aydemir-Koksoy A, Abramowitz J, and Allen JC. Ouabain-induced signaling and vascular smooth muscle cell proliferation. J Biol Chem 276: 46605–46611, 2001. 99. McConkey DJ, Lin Y, Nutt LK, Ozel HZ, and Newman RA. Cardiac glycosides stimulate Ca2+ increases and apoptosis in androgen-independent, metastatic human prostate adenocarcinoma cells. Cancer Res 60: 3807–3812, 2000. 100. Xiao AY, Wei L, Xia S, Rothman S, and Yu SP. Ionic mechanism of ouabain-induced concurrent apoptosis and necrosis in individual cultured cortical neurons. J Neurosci 22: 1350–1362, 2002. 101. Gonzalez-Zulueta M, Feldman AB, Klesse LJ, Kalb RG, Dillman JF, Parada LF, Dawson TM, and Dawson VL. Requirement for nitric oxide activation of p21(ras)/ extracellular regulated kinase in neuronal ischemic preconditioning. Proc Natl Acad Sci USA 97: 436–441, 2000. 102. Nicholls DG and Budd SL. Mitochondria and neuronal survival. Physiol Rev 80: 315–360, 2000. 103. Mintorovitch J, Yang GY, Shimizu H, Kucharczyk J, Chan PH, and Weinstein PR. Diffusion-weighted magnetic resonance imaging of acute focal cerebral ischemia: comparison of signal intensity with changes in brain water and Na+,K(+)-ATPase activity. J Cereb Blood Flow Metab 14: 332–336, 1994. 104. Jamme I, Petit E, Gerbi A, Maixent JM, MacKenzie ET, and Nouvelot A. Changes in ouabain affinity of Na+, K+-ATPase during focal cerebral ischaemia in the mouse. Brain Res 774: 123–130, 1997. 105. Tavalin SJ, Ellis EF, and Satin LS. Inhibition of the electrogenic Na pump underlies delayed depolarization of cortical neurons after mechanical injury or glutamate. J Neurophysiol 77: 632–638, 1997. 106. Jung YW, Choi IJ, and Kwon TH. Altered expression of sodium transporters in ischemic penumbra after focal cerebral ischemia in rats. Neurosci Res 59: 152–159, 2007. 107. Sheldon C, Diarra A, Cheng YM, and Church J. Sodium influx pathways during and after anoxia in rat hippocampal neurons. J Neurosci 24: 11057–11069, 2004. 108. Lees GJ. Inhibition of sodium-potassium-ATPase: a potentially ubiquitous mechanism contributing to central nervous system neuropathology. Brain Res Brain Res Rev 16: 283–300, 1991.

150

M.A. Gottron and D.C. Lo

109. Yu SP. Regulation and critical role of potassium homeostasis in apoptosis. Prog Neurobiol 70: 363–386, 2003. 110. Yu SP. Na(+), K(+)-ATPase: the new face of an old player in pathogenesis and apoptotic/hybrid cell death. Biochem Pharmacol 66: 1601–1609, 2003. 111. Bruer U, Weih MK, Isaev NK, Meisel A, Ruscher K, Bergk A, Trendelenburg G, Wiegand F, Victorov IV, and Dirnagl U. Induction of tolerance in rat cortical neurons: hypoxic preconditioning. FEBS Lett 414: 117–121, 1997. 112. Golden WC and Martin LJ. Low-dose ouabain protects against excitotoxic apoptosis and up-regulates nuclear Bcl-2 in vivo. Neuroscience 137: 133–144, 2006. 113. Isaev NK, Stelmashook EV, Halle A, Harms C, Lautenschlager M, Weih M, Dirnagl U, Victorov IV, and Zorov DB. Inhibition of Na(+),K(+)-ATPase activity in cultured rat cerebellar granule cells prevents the onset of apoptosis induced by low potassium. Neurosci Lett 283: 41–44, 2000. 114. Zhou X, Jiang G, Zhao A, Bondeva T, Hirszel P, and Balla T. Inhibition of Na,KATPase activates PI3 kinase and inhibits apoptosis in LLC-PK1 cells. Biochem Biophys Res Commun 285: 46–51, 2001. 115. Anderson CN and Tolkovsky AM. A role for MAPK/ERK in sympathetic neuron survival: protection against a p53-dependent, JNK-independent induction of apoptosis by cytosine arabinoside. J Neurosci 19: 664–673, 1999. 116. Bonni A, Brunet A, West AE, Datta SR, Takasu MA, and Greenberg ME. Cell survival promoted by the Ras-MAPK signaling pathway by transcription-dependent and -independent mechanisms. Science 286: 1358–1362, 1999. 117. Atwal JK, Massie B, Miller FD, and Kaplan DR. The TrkB-Shc site signals neuronal survival and local axon growth via MEK and P13-kinase. Neuron 27: 265–277, 2000. 118. Perron JC and Bixby JL. Distinct neurite outgrowth signaling pathways converge on ERK activation. Mol Cell Neurosci 13: 362–378, 1999. 119. Wang JK, Portbury S, Thomas MB, Barney S, Ricca DJ, Morris DL, Warner DS, and Lo DC. Cardiac glycosides provide neuroprotection against ischemic stroke: discovery by a brain slice-based compound screening platform. Proc Natl Acad Sci USA 103: 10461–10466, 2006. 120. Grotta J. Neuroprotection is unlikely to be effective in humans using current trial designs. Stroke 33: 306–307, 2002. 121. Hochachka PW. Defense strategies against hypoxia and hypothermia. Science 231: 234–241, 1986. 122. McMullen DC and Storey KB. Suppression of Na(+)K(+)-ATPase activity by reversible phosphorylation over the winter in a freeze-tolerant insect. J Insect Physiol 54: 1023–1027, 2008. 123. Ramnanan CJ and Storey KB. Suppression of Na+/K+-ATPase activity during estivation in the land snail Otala lactea. J Exp Biol 209: 677–688, 2006. 124. Perez-Pinzon MA, Rosenthal M, Sick TJ, Lutz PL, Pablo J, and Mash D. Downregulation of sodium channels during anoxia: a putative survival strategy of turtle brain. Am J Physiol 262: R712–R715, 1992. 125. MacDonald JA and Storey KB. Regulation of ground squirrel Na+K+-ATPase activity by reversible phosphorylation during hibernation. Biochem Biophys Res Commun 254: 424–429, 1999. 126. De Angelis C and Haupert GT Jr.. Hypoxia triggers release of an endogenous inhibitor of Na(+)-K(+)-ATPase from midbrain and adrenal. Am J Physiol 274: F182–F188, 1998. 127. Liu L and Yenari MA. Therapeutic hypothermia: neuroprotective mechanisms. Front Biosci 12: 816–825, 2007. 128. Johnson EM Jr. and Deckwerth TL. Molecular mechanisms of developmental neuronal death. Annu Rev Neurosci 16: 31–46, 1993.

The Naþ/Kþ-ATPase as a Drug Target for Ischemic Stroke

151

129. Johnson EM Jr., Koike T, and Franklin J. A ‘‘calcium set-point hypothesis’’ of neuronal dependence on neurotrophic factor. Exp Neurol 115: 163–166, 1992. 130. Connor JA, Razani-Boroujerdi S, Greenwood AC, Cormier RJ, Petrozzino JJ, and Lin RC. Reduced voltage-dependent Ca2+ signaling in CA1 neurons after brief ischemia in gerbils. J Neurophysiol 81: 299–306, 1999. 131. Biegon A, Fry PA, Paden CM, Alexandrovich A, Tsenter J, and Shohami E. Dynamic changes in N-methyl-D-aspartate receptors after closed head injury in mice: implications for treatment of neurological and cognitive deficits. Proc Natl Acad Sci USA 101: 5117–5122, 2004. 132. Lee JM, Grabb MC, Zipfel GJ, and Choi DW. Brain tissue responses to ischemia. J Clin Invest 106: 723–731, 2000. 133. de Rezende Correa G, Araujo dos Santos A, Frederico Leite Fontes C, Giestal de Araujo E. Ouabain induces an increase of retinal ganglion cell survival in vitro: the involvement of protein kinase C. Brain Res 1049: 89–94, 2005. 134. Salthun-Lassalle B, Hirsch EC, Wolfart J, Ruberg M, and Michel PP. Rescue of mesencephalic dopaminergic neurons in culture by low-level stimulation of voltagegated sodium channels. J Neurosci 24: 5922–5930, 2004. 135. Tzen JT, Jinn TR, Chen YC, Li FY, Cheng FC, Shi LS, She H, Chen BC, Hsieh V, and Tu ML. Magnesium lithospermate B possesses inhibitory activity on Na+,K+-ATPase and neuroprotective effects against ischemic stroke. Acta Pharmacol Sin 28: 609–615, 2007. 136. Piccioni F, Roman BR, Fischbeck KH, and Taylor JP. A screen for drugs that protect against the cytotoxicity of polyglutamine-expanded androgen receptor. Hum Mol Genet 13: 437–446, 2004. 137. von Moltke LL and Greenblatt DJ. Drug transporters revisited. J Clin Psychopharmacol 21: 1–3, 2001. 138. Beringer PM and Slaughter RL. Transporters and their impact on drug disposition. Ann Pharmacother 39: 1097–1108, 2005. 139. Schinkel AH, Wagenaar E, van Deemter L, Mol CA, and Borst P. Absence of the mdr1a P-Glycoprotein in mice affects tissue distribution and pharmacokinetics of dexamethasone, digoxin, and cyclosporin A. J Clin Invest 96: 1698–1705, 1995. 140. Kawahara M, Sakata A, Miyashita T, Tamai I, and Tsuji A. Physiologically based pharmacokinetics of digoxin in mdr1a knockout mice. J Pharm Sci 88: 1281–1287, 1999. 141. Sadeque AJ, Wandel C, He H, Shah S, and Wood AJ. Increased drug delivery to the brain by P-glycoprotein inhibition. Clin Pharmacol Ther 68: 231–237, 2000. 142. Mayer U, Wagenaar E, Dorobek B, Beijnen JH, Borst P, and Schinkel AH. Full blockade of intestinal P-glycoprotein and extensive inhibition of blood-brain barrier P-glycoprotein by oral treatment of mice with PSC833. J Clin Invest 100: 2430–2436, 1997. 143. Fromm MF, Kim RB, Stein CM, Wilkinson GR, and Roden DM. Inhibition of P-glycoprotein-mediated drug transport: A unifying mechanism to explain the interaction between digoxin and quinidine [seecomments]. Circulation 99: 552–557, 1999. 144. Batrakova EV, Miller DW, Li S, Alakhov VY, Kabanov AV, and Elmquist W. Pluronic P85 enhances the delivery of digoxin to the brain: in vitro and in vivo studies. J Pharmacol Exp Ther 296: 551–557, 2001. 145. Fricker G and Miller DS. Modulation of drug transporters at the blood-brain barrier. Pharmacology 70: 169–176, 2004.

Acid-Sensing Ion Channels (ASICs): New Targets in Stroke Treatment Giuseppe Pignataro

1 Introduction In the last decades enormous effort has been made in an attempt to attenuate stroke injury. Despite the attractiveness of the excitotoxicity hypothesis, all clinical trials using glutamate antagonists as protective agents against ischemia have failed. This may be partially due to other injury pathways and non-excitotoxic processes involved in the complex mechanisms triggered by the lack of blood supply in brain. In this regard, the role played by acid-sensing ion channels (ASICs) in cerebral ischemia has been recently underlined [15, 57, 83]. A stable pH is critical for normal cellular function [24]. In physiological conditions, extracellular pH (pHo) and intracellular pH (pHi) are maintained at 7.3 and 7.0 through different Hþ-transporting mechanisms [24]. In pathological conditions such as tissue inflammation, neurotrauma, epileptic seizure, and especially ischemic stroke, accumulation of lactic acid due to enhanced anaerobic glucose metabolism and release of Hþ from ATP hydrolysis result in marked reduction of tissue pH, a condition termed acidosis. In particular, during severe ischemia brain pH can drop to as low as 6.0 [25, 57, 62]. It has been known for several decades that acidosis occurring after ischemia is associated with neuronal injury [67]. Recent studies have reported that activation of the Ca2þ-permeable ASIC1a plays a pivotal role in the development of acidosis-mediated ischemic damage [57, 83, 86]. These findings dramatically changed the view of acid signaling and offered new pharmacological targets for those

G. Pignataro (*) Department of Neuroscience, School of Medicine, Federico II University of Naples, Via Sergio Pansini 5, 80131 Naples, Italy e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_9, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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neurological diseases in which acidosis is a relevant pathophysiological event [15, 57, 79, 83, 86].

2 Biochemical and Molecular Biology of the Plasma Membrane Protein Since the first subunit was cloned in 1997 [75], six ASICs, encoded by four genes, have been identified: ASIC1a, ASIC1b, ASIC2a, ASIC2b, ASIC3, and ASIC4. All ASICs belong to the degenerin/epithelial Naþ channel (DEG/ ENaC) superfamily, which are Naþ-selective cation channels sensitive to the Kþ-sparing diuretic amiloride [3, 75]. ASICs are found throughout the mammalian central and peripheral nervous systems with ASIC1a, ASIC2a, and ASIC2b present in the brain, especially in cerebral cortex, cerebellum, hippocampus, amygdala, and olfactory bulb, and ASIC1a, ASIC1b, ASIC2a, ASIC2b, and ASIC3, abundant in the sensory neurons of the peripheral nervous system [4, 75, 77]. Based on the biochemical analysis of ENaC [63, 75] and the glycosylation studies of ASIC2a subunit [65], the proposed membrane topology of each ASIC subunit consists of two transmembrane domains (TM1 and TM2), linked by a large extracellular cystein-rich loop, and intracellular N- and C- termini. Functional ASICs are believed to be tetrameric assemblies of homomeric or heteromeric subunits [50]. However, based on the stoichiometric studies of other DEG/ENaC, the possibility that ASICs are assembled with three to nine subunits cannot be excluded [35, 38]. Recently, with the aid of crystallographic studies it has been possible to state that ASIC1 subunit stoichiometry is trimeric, and thus the obligate heteromeric ENaCs are almost certainly trimers, defined by an A1B1C1 stoichiometry [47]. Furthermore, the large extracellular portion of ASIC1 polypeptide is composed of multiple domains that together form a complex structure rich in cavities and protrusions and perhaps prominent ‘‘handholds’’ for additional interacting proteins. Multiple close acidic residue pairs have been identified as proton-binding sites, thus demonstrating that the pH-sensing sites are distributed in primary structure and 3D space, far from the channel region. To explain how proton binding at these distant sites is translated into channel gating, it has been suggested that the disulphide-rich thumb domain undergoes movement on proton binding/unbinding and that these movements are transmitted to the ion channel transmembrane domain by way of primarily non-covalent interactions, mediated by residues at the base of the thumb. Trp 288 (thumb) may act like a ball sitting in a socket-like joint of the transmembrane domain and perhaps when the thumb domain moves, its movement is coupled to the transmembrane domain by way of this thumb–transmembrane domain ball-and-socket joint (Fig. 1) [47].

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Fig. 1 Putative structure of acid-sensing ion channel 1(ASIC1)

3 Tissue Distribution In the CNS, ASIC1 is the most abundantly expressed channel of the ASIC family [41, 75]. With the aid of in situ hybridization, ASIC1 mRNA has been found in the olfactory bulb, cerebral cortex, hippocampus, basolateral amydgaloid nuclei, subthalamic nuclei, and cerebellum [41, 75]. It should be here underlined that effective activation of ASIC1 requires rapid and large decreases in pHo of 1 pH unit [4, 10, 14, 75]. Protons also induce desensitization, thus, re-activation of channels requires the pHo to return to values >7.3. Recovery from desensitization is slow, with <50% recovery after 4 s of exposure to neutral pHo [14]. When all of these properties are considered, it is possible to conclude that effective activation of ASIC1 in the CNS may occur in locations where the pHo can change rapidly, profoundly, and reversibly. Those conditions may be met in a few microenvironments such as the lumen of intracellular vesicles or the synaptic cleft. This is an attractive possibility because at this level, synaptic vesicles repeatedly release acidic content, pH 5.6 [54], in a small and delimited space. In response to repetitive high-frequency stimulation, the release of the acidic content of synaptic vesicles could temporarily overwhelm the mechanisms for proton buffering, diffusion, and extrusion in the synaptic cleft, and the pHo could decrease sufficiently to gate ASIC1. It should be noticed, however, that the kinetics and extent of pHo changes in the synaptic cleft have not been experimentally determined. Within neurons, ASIC1 is expressed predominantly on the plasma membrane of the soma and, to a lesser degree, over the dendrites and axon. ASIC1 is detected at E12, and the levels of expression remain constant until adulthood [4]. There is no increase in the expression of ASIC1 during the periods of most active synaptogenesis. ASIC1 appears early and persists at almost constant levels throughout the development of the mouse brain [4].

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ASIC1 is expressed in almost all areas of the rat CNS, including the forebrain, midbrain, brainstem, pons, cerebellum, hippocampus, and spinal cord.

4 Biophysical and Electrophysiological Aspects Individual ASIC properties have been analyzed largely in heterologous expression systems by means of electrophysiology technique. Homomeric ASIC1a channels respond to low pHo by mediating a fast and transient inward current with a threshold pH of 7.0, and the pH for half maximal activation (pH0.5) ranges from 6.2 [75] to 6.8 [14]. In addition to conducting Naþ ions, homomeric ASIC1a channels are permeable to Ca2þ ions [14]. ASIC1b is a splice variant of ASIC1a with restricted expression in sensory neurons [21]. Homomeric ASIC1b channels respond to pHo drop with a similar transient current and a pH0.5 of 5.9 [12, 21]. Unlike ASIC1a, homomeric ASIC1b channels do not show Ca2þ permeability. ASIC2a is expressed broadly in peripheral sensory and CNS neurons. Homomeric ASIC2a channels have a low sensitivity to protons with a pH0.5 of 4.4 [53, 76]. ASIC2b is a splice variant of ASIC2a [53]. Though widely expressed in peripheral sensory and central neurons, ASIC2b subunits do not form functional homomeric channels. However, they may associate with other subunits to form heteromeric ASICs with distinct properties [53]. ASIC3 is predominantly expressed in dorsal root ganglia [74]. Homomeric ASIC3 channels respond to pHo drops by a biphasic response with a fast desensitizing current followed by a sustained component [70, 74]. These ASIC3 channels have a high sensitivity to protons and a pH0.5 of 6.7 has been reported [70]. ASIC4 subunits show high level of expression in pituitary gland. Similar to ASIC2b, they do not form functional homomeric channels [1, 43]. Although the exact subunit combination and stoichiometry of ASICs in native neurons remain to be determined, the relative contributions by ASIC1a, ASIC2a, or ASIC3 subunit to acid-evoked currents in peripheral sensory and CNS neurons have been examined [7, 14, 78, 83]. In mediumsized DRG neurons, for example, acid-activated currents match those recorded from heterologous cells expressing a mix of ASIC1, ASIC2, and ASIC3 subunits [14]. Deletion of any one subunit did not abolish acid-activated currents, but altered currents in a manner consistent with heteromultimerization of the two remaining subunits, indicating that combinations of two or more ASIC subunits co-assemble as heteromultimeric channels in mouse DRG neurons [14]. In cortical and hippocampal neurons, however, knock out of ASIC1 gene alone almost completely eliminated the acid-activated current [7, 78, 83], suggesting that ASIC1a is a predominant functional ASIC subunit in CNS neurons. Further studies suggest that the acid-activated currents in CNS neurons are largely mediated by a combination of ASIC1a homomeric channels and ASIC1a/ASIC2a heteromeric channels [7, 10]. ASIC1a is key in establishing the

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current amplitude. ASIC2a, on the other hand, has little effect on the amplitude but influences desensitization, recovery from desensitization, and pH sensitivity of the channels [10].

5 Regulatory Mechanisms Although proton is the only known agonist for the activation of ASICs, a variety of extracellular and intracellular signaling molecules can modulate the activities of ASICs and have profound influence on the functions of these channels in both physiological and pathological processes. First, not all ASICs interact with the same partners. For example, protein interacting with C kinase (PICK)1 interacts directly with ASIC1 and ASIC2, but not ASIC3 [84]. Second, some interactions occur via the C-terminal PDZ domains, at least one (with annexin) occurs via the N-terminus [84], and for others the interacting sites have not yet been identified. Third, at least two interactions, with PICK1 and CaMKII, facilitate channel phosphorylation [84]. Fourth, some interactions affect channel-gating properties whereas others might increase or decrease channel density on the cell surface. One of the interactions more deeply studied has been the one between ASICs and PICK1. PICK1 is a synaptic protein with a PDZ domain; it interacts with the C-termini of both ASIC1a and ASIC2a, thus mediating the regulation of these channels by protein kinase C (PKC) [9, 32, 46]. Similar to the involvement of MEC-2 in mechanosensation in Caenorhabditis elegans, stomatin, the mammalian homolog of MEC-2, has been shown to interact with and modulate the activity of ASIC1a, ASIC2a, and ASIC3 [60]. These interacting proteins have been shown to influence the function of ASICs. While PICK1 binding to ASIC1a is associated with phosphorylation and changes in the cellular localization of the channel [51], the interaction between stomatin and ASICs has been suggested to modulate channel gating [60]. Recently, a kinase-anchoring protein 150 (AKAP150) and the Ca2þ/calmodulin-dependent protein phosphatase 2B, also called calcineurin, have been proposed as proteins interacting with ASIC1a and ASIC2a. AKAP150, the neuron-specific rat ortholog of human AKAP79, is present in the postsynaptic density (PSD) in association with cAMP-dependent protein kinase (PKA), PKC, and calcineurin [26, 27, 48]. These enzymes are associated with AKAP79/150 in inactive states. Each of these interactions might have important consequences, but much remains to be learned. A detailed knowledge of these interactions is crucial to understand how ASICs function in complex systems and the interaction with the anchoring protein has different consequences for the activity of these enzymes. In the case of PKA, only the regulatory subunit, RII, is anchored to AKAP and the catalytic subunit, a serine/threonine kinase, interacts with RII forming active enzyme [42, 48]. PKC and calcineurin, on the other hand, are inactive when associated with AKAP79/150, until they are released from the

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anchoring protein [26, 37]. Disruption of the RII binding to AKAP150 by synthetic peptide Ht-31 decreased acid-evoked currents in mouse cortical neurons and Chinese hamster ovarian cancer (CHO) cells expressing homomeric ASIC1a or ASIC2a function, implying involvement of AKAP-anchored PKA in the function of ASIC1a and ASIC2a. Furthermore, inhibition of calcineurin by cyclosporin A significantly increased acid-evoked current in neuronal cells as well as in CHO cells. Consequently, it is possible to state that both AKAP150 and calcineurin are involved in regulation of ASIC1a and ASIC2a, possibly by changing the phosphorylation status of these channels [18]. Among the other known interactions, peptides such as FMRFamide can enhance ASIC1 and ASIC3 activity [82], and arachidonic acid can potentiate ASIC activation [2, 69]. Interestingly, it has recently been shown that the physiological pro-inflammatory mediator nitric oxide enhances the activation of ASICs by protons [16]. The action is at the external membrane surface and is likely to involve the formation of new cysteine–cysteine bonds. The dependence of channel activation on pH is steepened, consistent with an unmasking of additional residues whose protonation leads to channel activation [16]. The potentiation of ASICs activity by nitric oxide is likely to be of physiological relevance in pathologies in which both tissue acidosis and a high level of NO and/or other free radicals may occur together. For example, in stroke the occurrence of all these conditions is easily achievable during the reperfusion period. An enhancement of ASIC function by NO may therefore play an important role in stroke.

6 Pharmacological Modulation 6.1 Amiloride Amiloride has been in clinical use as a diuretic since 1967. Its classic therapeutic action consists in the inhibition of the epithelial sodium channel (ENaC) expressed in the renal tubules. In addition, amiloride is also able to inhibit the Naþ/Hþ and Naþ/Ca2þ antiporters [17, 49], and ASICs, in a non-specific manner. It reversibly inhibits the ASIC currents with an IC50 of 10–50 mM [1, 2, 21, 74, 75, 83]. Similar to its effect on the ASIC currents, amiloride inhibits acid-induced increase of [Ca2þ]i and membrane depolarization [73, 81, 83, 86]. The sustained component of the ASIC3 current, on the other hand, is much less or completely insensitive to amiloride [13, 74]. In fact, a recent study has shown that in cardiac sensory neurons, the small-sustained ASIC3-like current activated at pH 7.0 is in fact increased by amiloride [85]. Based on the studies of ENaC [66], it is believed that amiloride inhibits the ASIC current by a direct blockade of the channel, and that the pre-TM2 region of the channel is critical for amiloride effect. Mutation of Gly-430 in this region dramatically increases the sensitivity of ASIC2a to amiloride [19].

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6.2 A-317567 A-317567, a small molecule unrelated to amiloride (Fig. 2), is a new non-selective ASIC blocker [68]. It inhibits the ASIC1a-like, ASIC2a-like, and ASIC3-like currents in rat DRG neurons with IC50 of 2–30 mM. Unlike amiloride, A-317567 blocks both the fast and the sustained components of the ASIC3-like currents. In rat thermal hyperalgesia model, A-317567 is fully efficacious at a dose 10-fold lower than amiloride. It is also effective in skin incision model of postoperative pain. A-317567 does not show diuresis or natriuresis activity [68], suggesting that it might be more specific for ASICs than amiloride.

Fig. 2 Chemical structure of amiloride and A-317567

6.3 PcTX1 Psalmotoxin 1 (PcTX1) is a peptide toxin, isolated from the venom of South American tarantula Psalmopoeus cambridge, that specifically inhibits the ASIC1a current [34]. The toxin contains 40 amino acids cross-linked by three disulfide bridges (Fig. 3). In heterologous expression systems, PcTX1 potently and specifically inhibits the acid-activated current mediated by homomeric ASIC1a subunits in a nanomolar concentration range (IC50 < 1 nM), without affecting the currents mediated by other configurations of ASICs [34]. At concentrations that effectively inhibit the ASIC1a currents, it has no effect on voltage-gated Naþ, Kþ, Ca2þ channels, as well as several ligand-gated ion channels [83]. Thus, PcTX1 is so far the best-known specific blocker for ASICs and an indispensable pharmacological tool for the studies of ASIC1amediated processes [28, 34, 36, 83]. Unlike amiloride which directly blocks the channel, PcTX1 acts as a gating modifier. In fact, PcTX1 shifts the channel from its resting toward the inactivated state through an increase of its apparent affinity for protons [23]. Interestingly, this PcTX1-induced shift of the pH-dependent inactivation of ASIC1a

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Fig. 3 Aminoacid sequence and disulfide linkage for Pc TX1 and APETX2

is Ca2þ dependent, as an increase in extracellular Ca2þ results in a decrease of the PcTX1 inhibition [23]. This finding implies that, in neurological conditions (e.g., brain ischemia) where significant drop of the extracellular Ca2þ occurs [33, 44, 57], the potency for PcTX1 inhibition of ASIC1a channels would increase. The binding site for PcTX1 has recently been analyzed using radio-labeled tools [64]. It binds principally on cysteine-rich domains I and II (CRDI and CRDII) of ASIC1 extracellular loop. Although the post-transmembrane domain I (M1) and pre-transmembrane domain II (M2) regions are not involved in the binding, they are crucial for the ability of PcTX1 to inhibit the ASIC1a current [64].

6.4 APETX2 APETX2 is a 42-amino-acid peptide toxin isolated from sea anemone Anthopleura elegantissima. It is a potent and selective inhibitor for homomeric ASIC3- and ASIC3- containing channels [29]. It reduces transient peak acidevoked current mediated by homomeric ASIC3 channels in heterologous expression systems and in primary cultures of sensory neurons [29]. In contrast to the peak ASIC3 current, the sustained component of the ASIC3 current is insensitive to APETX2. In addition to homomeric ASIC3 channels (IC50 ¼ 63 nM for rat and 175 nM for human), APETX2 inhibits heteromeric ASIC3/1a (IC50 ¼ 2 mM), ASIC3/1b (IC50 ¼ 900 nM), and ASIC3/2b (IC50 ¼ 117 nM). Homomeric ASIC1a, ASIC1b, ASIC2a, and heteromeric ASIC3/2a channels, on the other hand, are not sensitive to APETX2. Similar to PcTX1, APETX2 is cross-linked by three disulfide bonds (Fig. 3) [29]. However, it does not show any sequence homologies with PcTX1. At present, the mode of action for APETX2 is still unknown.

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6.5 Non-steroid Anti-inflammatory Drugs Non-steroid anti-inflammatory drugs (NSAIDs) are the most commonly used anti-inflammatory and analgesic agents. The well-accepted mechanism for the effect of NSAIDs is the inhibition of the synthesis of prostaglandins (PGs). However, exceptions to the correlation of PG synthetase inhibition with anti-inflammatory activity have been noted, suggesting additional mechanism(s) may be involved. A recent study demonstrated that various NSAIDs also inhibit the activity of ASICs at their therapeutic doses for analgesic effects [72]. Ibuprofen and flurbiprofen, for example, inhibit ASIC1a-containing channels with an IC50 of 350 mM. Aspirin and salicylate inhibit ASIC3-containing channels with an IC50 of 260 mM, whereas diclofenac inhibits the same channels with an IC50 of only 92 mM. In addition to a direct inhibition of the ASIC activity, NSAIDs also prevent inflammation-induced increase of ASIC expression in sensory neurons [72]. The inhibitory action of non-steroid anti-inflammatory drugs has been investigated on ASICs in isolated hippocampal interneurons and on recombinant ASICs expressed in CHO cells. Diclofenac and ibuprofen inhibited proton-induced currents in hippocampal interneurons (IC50 were 622 34 mM and 3.42 0.50 mM, respectively) [30]. This non-competitive effect is fast and fully reversible for both drugs. Interestingly, simultaneous application of acid solution and diclofenac is required for its inhibitory effect. Unlike amiloride, the action of diclofenac is voltage independent and no competition between two drugs has been found. Analysis of the action of diclofenac and ibuprofen on activation and desensitization of ASICs showed that diclofenac but not ibuprofen shifted the steady-state desensitization curve to more alkaline pH values. The reason for this shift was slowing down the recovery from desensitization of ASICs. Thus, diclofenac may serve as a neuroprotective agent during pathological conditions associated with acidification [30].

7 Physiological Role 7.1 Activation of ASICs Induces Membrane Depolarization and Increased Intracellular Ca2+ in Neurons Since all ASICs are Naþ-selective channels, which have a reversal potential near Naþ equilibrium potential (þ60 mV), activations of ASICs at normal resting potentials produce exclusively inward currents which result in membrane depolarization and neuron excitation [10, 52]. For homomeric ASIC1a channels, acid activation also induces Ca2þ entry directly through these channels [75, 83, 86]. In addition, the ASIC-mediated membrane

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depolarization may facilitate the activation of voltage-gated Ca2þ channels and NMDA receptor-gated channels [78], further promoting neuronal excitation and [Ca2þ]i accumulation. The Ca2þ permeability of ASICs in native neurons has been characterized using fluorescent Ca2þ imaging and ion-substitution protocol [83, 86]. In mouse cortical and hippocampal neurons, activation of ASICs by decreasing pHo induces increases of [Ca2þ]i. This acid-induced increase of [Ca2þ]i could be recorded in the presence of a cocktail of drugs blocking other voltage-gated and ligand-gated Ca2þ channels [83, 86], indicating Ca2þ entry directly through ASICs. The acid-induced increase of [Ca2þ]i is eliminated by specific and nonspecific ASIC1a blocker, or by ASIC1 gene knockout [83, 86]. Consistent with the finding of fluorescent imaging, acid-activated inward current is activated when extracellular solution contains Ca2þ as the only conducting cations [83, 86]. Thus, homomeric ASIC1a channels constitute an additional and important Ca2þ entry pathway for neurons.

8 Pathophysiological Role in Stroke and in Other Neurodegenerative Diseases 8.1 ASIC1a Activation in Acidosis-mediated and Ischemic Neuronal Injury During ischemia, increased anaerobic glycolysis due to reduced oxygen supply leads to lactic acid accumulation [62]. Accumulation of lactic acid, alone with increased Hþ release from ATP hydrolysis, causes a decrease in brain pH, or acidosis. During brain ischemia, pHo falls to 6.5 or lower [55, 62]. Acidosis has long been recognized to play an important role in ischemic brain injury [67, 71]. However, the cellular and molecular mechanism remained unclear. The widespread expression of ASIC1a in the brain, its activation by pH drop to the level commonly seen in ischemic brain, and its demonstrated permeability to Ca2þ strongly suggested that activation of ASIC1a might be involved in the pathology of brain injury. Indeed, a series of recent studies have clearly demonstrated a role of ASIC1a activation in acidosis-mediated and ischemic neuronal injury [40, 57, 83, 86] (Fig. 4). In cultured mouse cortical neurons, activation of ASICs by brief acid incubation induces glutamate receptor-independent Ca2þ-dependent neuronal injury that is inhibited by specific and non-specific ASIC1a blockade, and by ASIC1 gene knockout [83]. Reducing [Ca2þ]o, which lowers the driving force for Ca2þ entry through ASICs, also decreases the acid-induced neuronal injury. Intracerebroventricular injection of ASIC1a blocker in rodents reduces the infarct volume induced by transient or permanent focal ischemia by up to 60% [57, 83]. Similarly, ASIC1 gene knockout produces a significant neuroprotection [57, 83]. The protection by ASIC1a blockade has up to5-h time window, and persists for at least 7 days

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Fig. 4 Effect of ASIC1a genetic ablation on cerebral ischemic induced in mice by transient occlusion of cerebral artery. From Benveniste and Dingledine, NEJH 352: 85–86, (2005) MMs Reference Number: Ps-2009-1953 Copyright [2005] Massachusetts Medical Society. P.T.O All rights reserved [2,11].

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[57, 83]. Furthermore, attenuating brain acidosis by intracerebroventricular administration of NaHCO3 is also protective. Since activation of NMDA receptor and subsequent Ca2þ toxicity has been known to play an important role is ischemic brain injury, the outcome of co-application of both blockers has also been investigated. Compared to ASIC1a or NMDA blockade alone, co-application of NMDA and ASIC blockade produces additional neuroprotection, and the presence of ASIC1a blockade prolongs the time window of effectiveness of NMDA blockade [57, 83]. Summarizing the results so far obtained in studies devoted to understand the role of ASICs in the stroke pathophysiology, it is possible to state that acidosis occurring in cerebral ischemia is able to activate ASICs in in vivo and in vitro models of stroke. In particular, studies conducted in cortical neurons subjected to conditions mimicking cerebral ischemia showed that when studied at resting membrane potential, acidosis activated a depolarizing current mediated by influx of cations; this required activation of ASIC1a in particular, because this response was abolished in neurons cultured from ASIC1a–/– mice. If neurons cultured from wild-type mice were deprived of oxygen and glucose, thereby mimicking some features of ischemia, acidosis resulted in a larger and more sustained current. Interestingly, Ca2þ permeated the acidosis-activated channel and the influx of Ca2þ contributed to acidosis-induced death of neurons in vitro. Most importantly, limiting activation of ASICs reduced the damage produced by ischemia in vivo by approximately 60%, a conclusion based on both pharmacological evidence and study of ASIC1a–/– mice. Furthermore, the pharmacologic and genetic interventions limiting ASIC1 activation exerted neuroprotective effects even in the presence of a glutamate receptor antagonist. Multiple molecular targets have been identified for which neuroprotective therapies have been shown to be effective in animal models including nitric oxide synthase, cannabinoid receptors, and many others. What sets apart the results obtained with ASICs is the simple yet plausible mechanism: the marked drop of pH damages the ischemic tissue and does so by activation of this ASIC receptor. Importantly, the drop of pH in the core of the ischemic lesion raises the possibility that preventing activation of the ASIC receptor may reduce damage in the core, not simply in the partially ischemic tissue surrounding the core that is targeted by most neuroprotective treatments. Thus, the discoveries made in this field of research raises the exciting possibility that small molecules could be developed to prevent acidosis-mediated activation of this receptor and thereby reduce ischemic injury.

8.2 ASIC Activation and Epileptic Seizure Activity The role of ASIC in epileptic seizure has been recently questioned. In fact, though a significant drop of brain pH during intense neuronal excitation or seizure activity [23, 28, 34, 36] would suggest that ASIC activation might play a

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detrimental role in the generation/maintenance of epileptic seizure activities, recent findings showed that ASIC1a activation is important in seizure termination. In a cell culture model of epilepsy, brief withdrawal of the NMDA antagonist kynurenic acid induces a dramatic increase in the firing of action potential, in addition to a sustained membrane depolarization [24]. Some studies showed that ASIC blockade by amiloride and PcTX1 significantly inhibit the increase of neuronal firing and the sustained membrane depolarization [20]. In hippocampal slices, high-frequency electrical stimulation or removal of extracellular Mg2þ induces triggered or spontaneous seizure-like bursting. Bath perfusion of amiloride and PcTX1 decrease the amplitude and the frequency of these seizurelike bursting activities. In in vivo model of epilepsy where intra-amygdala injection of kainic acid, KA, induces sustained polyspike activity on EEG followed by dramatic injury of CA3 neurons, intracerebroventricular injection of PcTX1 reduces both the electrographic seizure activity and the CA3 neuronal injury [20]. Together, these data suggest that activation of ASICs, particularly the ASIC1a channels, is involved in the generation or maintenance of seizure activity and seizure-mediated neuronal injury. In contrast with the above-reported results, recent data suggest that acidosis occurring after seizure is able to stop seizure and the pH sensitivity of ASIC1a. In fact, disrupting the ASIC1a gene or pharmacologically inhibiting ASIC1a increased seizure severity, whereas overexpressing ASIC1a had the opposite effect [87]. These findings suggest that ASIC1a is part of a feedback inhibition system that limits seizure severity [87]. Therefore, ASIC1a would contribute to seizure termination thus reducing seizure severity. Whether ASIC channels are protective or damaging may depend on the magnitude and duration of acidosis, the location of ASIC activation, and the presence of factors that modulate ASIC function, such as lactate. However, these latter findings suggest a new, protective function for ASIC1a in brain physiology. Thus, agents that potentiate ASIC1a activity might reduce seizure severity or duration and possibly prevent status epilepticus.

8.3 ASIC and Huntigton Despite a tremendous effort to develop therapeutic tools in several Huntington’s disease (HD) models, there is no effective cure at present. Acidosis has been observed previously in cellular and in in vivo models as well as in the brains of HD patients. In a recent study HD models were challenged with amiloride derivative benzamil to examine whether chronic acidosis is an important part of the HD pathomechanism and whether these drugs could be used as novel therapeutic agents [80]. Benzamil markedly reduced the huntingtin (htt)–polyglutamine (polyQ) aggregation in an inducible cellular system, and the therapeutic value of benzamil was successfully confirmed in the R6/2 animal

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model of HD [80]. To reveal the mechanism of action, benzamil was found to be able to alleviate the inhibition of the ubiquitin–proteasome system (UPS) activity, resulting in enhanced degradation of soluble htt–polyQ specifically in its pathological range. More importantly, it has been shown that blocking the expression of a specific isoform of ASIC1a led to an enhancement of UPS activity and this blockade also decreased htt–polyQ aggregation in the striatum of R6/2 mice [80]. Data obtained in this study propose chemical compounds that target ASIC1a as effective and promising approach to combat HD and other polyglutamine-related disorders.

9 Future Perspectives 9.1 Therapeutic Potential of Drugs Acting on ASICs Amiloride. In peripheral sensory system, amiloride has been shown to suppress acid-induced pain [50–53] whereas in CNS neurons, it reduces acid-mediated and ischemic neuronal injury [10, 12]. However, due to its non-specificity for various ion channels and ion-exchange systems, it has low potential to be used as a future analgesic or neuroprotective agent in human subjects. In a similar or lower concentration range as it inhibits ASICs, amiloride also blocks other ion channels (e.g., ENaC, T-type Ca2þ channels) and ion-exchange systems (Naþ/ Hþ and Naþ/Ca2þ exchangers). Recent studies have suggested that the normal activity of Naþ/Ca2þ exchanger, for example, is critical for maintaining the cellular calcium homeostasis and the survival of neurons against delayed calcium deregulation and injury caused by glutamate receptor activation [5, 8, 56, 58]. Conversely, inhibition of Naþ/Ca2þ by amiloride is expected to compromise normal neuronal Ca2þ handling, which may transform the Ca2þ transient elicited by non-toxic glutamate concentrations into a lethal Ca2þ overload. The inhibition of Naþ/Ca2þ exchanger by amiloride may partially explain its reduced neuroprotective efficacy in vivo compared with the ASIC1a-specific blocker PcTX1 [83]. It may also explain the finding that prolonged incubation with amiloride (e.g., 5 h) itself induces the injury of cultured neurons [83]. Given that, amiloride has been tested in several animal models of neurodegenerative diseases like Parkinson’s Disease (PD) associated with cerebral lactic acidosis [6]. Amiloride was found to protect substantia nigra (SNc) neurons from MPTP-induced degeneration, as determined by attenuated reductions in striatal tyrosine hydroxylase (TH) and dopamine transporter (DAT) immunohistochemistry, as well as smaller declines in striatal DAT radioligand binding and dopamine levels. More significantly, amiloride also preserved dopaminergic cell bodies in the SNc [6]. Amiloride was equally neuroprotective in in vitro and in vivo models of multiple sclerosis [39], HD [80] and in other acidosis-related neurodegenerative disease. Although

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amiloride itself is unlikely to be ideal as a neuroprotective agent for stroke therapy, one direction for future drug development could be to chemically modify the structure of amiloride (Fig. 2) to achieve a molecule that specifically blocks ASICs, or at least has more selectivity for these channels. A-317567. Compared with amiloride, A-317567 appears to have better potential to be established as a future analgesic agent. Its inhibition of sustained ASIC3 current suggests that it might be useful in suppressing acidosis-mediated chronic pain. Currently, it is unknown whether A-317567 is also an effective neuroprotective agent. Future study should be conducted in the attempt to demonstrate whether it is effective in reducing acidosis-induced neuronal injury in in vitro and in vivo models of stroke and of other neurodegenerative diseases. When applied peripherally, A-317567 shows minimal brain penetration in normal condition [31]. It would be interesting to know whether it can reach the brain in pathological conditions (e.g., ischemia) where blood–brain barrier may have been compromised. Finally, additional studies are necessary to elucidate the mechanism underlying A-317567-mediated ASIC inhibition. It is also important to know whether A-317567 affects the activities of other ion channels and ion-exchanger systems. PcTX1. Targeting Ca2þ permeable ASIC1a by intracerebroventricular administration of PcTX1 has been demonstrated to be effective in reducing ischemic brain injury in rat and mouse models of ischemia, with neuroprotective time window of 5 h [57, 83]. These findings suggest that a specific ASIC1a blocker may be useful as a neuroprotective agent for various acid-related neurological diseases. However, as discussed below, PcTX1 itself may not be an ideal pharmacological agent for human subject: [1] PcTX1 consists of 40 amino acids linked by three disulfide bonds. Synthesis of this toxin in a large amount might be a challenge for the pharmaceutical companies. [2] Due to the presence of three disulfide bonds, which are subjected to modification by oxidizing/reducing conditions, the long-term stability of the toxin could be an issue. [3] The large molecule of this toxin (5 kDa) makes it difficult to across the blood–brain-barrier (BBB) thus preventing its use by conventional routine of administration (e.g., i.v. or i.p.). Indeed, it has not been demonstrated in animal studies that a peripheral administration of this toxin (i.v.) is sufficient in reducing the ischemic brain injury [57, 83]. Future effort may be attempted in making shorter or truncated peptide which can pass BBB, has long-term stability, but still blocks the ASIC1a channels. Additional effort should be focused on the search of new heterocyclic small molecules that specifically blocks the ASIC1a channels. Since activation of ASIC1a is also involved in the processes of learning, memory, and fear [28, 78], blocking these channels in patients is expected to induce some behavioral changes. APETx2. Since activation of ASIC3 has been implicated in various pain processes [22, 59, 68, 85], APETX2 may be a useful analgesic agent in the treatment or prevention of pain in peripheral sensory system. However, its lack of inhibition of sustained ASIC3 current suggests that it may not be effective in suppressing the chronic pain stimuli. It is also unknown whether it

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inhibits inflammation-induced increased expression of ASICs. The lack of inhibition of ASIC expression, and likely the COXs activity, would suggest that APETx2 will not be as effective as NSAIDs (see below) in suppressing the pain by tissue inflammation. Non-Steroidal Anti-Inflammatory Drugs. During tissue inflammation, ischemia, and infection, or in tumors, hematomas, and blisters, pHo can drop from 7.4 to as low as 5.0 [45, 61]. The combined inhibition of NSAIDs on PG synthesis, ASIC currents, and ASIC expression make them ideal for a large spectrum of pain conditions, particularly the pain caused by tissue inflammation. In the acute phase of tissue inflammation, for example, the rapid inhibition of NSAIDs on ASIC currents blocks the activation of pain-sensing neurons by inflammatory acidosis. Later, the NSAIDs further suppress the inflammation and pain by their effect on COXs, limiting the production of prostaglandins. In the chronic phase, they may reduce the sensitization to pain by combined inhibition of COXs, ASIC currents, and ASIC expression. In conclusion, ASICs represent new biological components and therapeutic targets in peripheral sensory and CNS neurons. Increasing evidence supports the involvement of ASIC activation in physiological processes such as synaptic plasticity, and in neurological diseases such as brain ischemia and epileptic seizures. Ongoing studies are expected to identify the involvement of ASIC activation or changes in its expression in other physiological processes and neurological disorders. Future development of potent and specific blocker for individual ASIC subunit will dramatically advance our understanding of the role of these channels in physiological and pathological processes, and it will be helpful in establishing novel therapeutic strategy for neurological diseases.

References 1. Akopian AN, Chen CC, Ding Y, Cesare P, and Wood JN. A new member of the acidsensing ion channel family. Neuroreport 11: 2217–2222, 2000. 2. Allen NJ and Attwell D. Modulation of ASIC channels in rat cerebellar Purkinje neurons by ischaemia-related signals. J Physiol 543: 521–529, 2002. 3. Alvarez de la Rosa D, Canessa CM, Fyfe GK, and Zhang P. Structure and regulation of amiloride-sensitive sodium channels. Annu Rev Physiol 62: 573–594, 2000. 4. Alvarez de la Rosa D, Krueger SR, Kolar A, Shao D, Fitzsimonds RM, and Canessa CM. Distribution, subcellular localization and ontogeny of ASIC1 in the mammalian central nervous system. J Physiol 546: 77–87, 2003. 5. Annunziato L, Pignataro G, and Di Renzo GF. Pharmacology of brain Na+/Ca2+ exchanger: from molecular biology to therapeutic perspectives. Pharmacol Rev 56: 633–654, 2004. 6. Arias RL, Sung ML, Vasylyev D, Zhang MY, Albinson K, Kubek K, Kagan N, Beyer C, Lin Q, Dwyer JM, Zaleska MM, Bowlby MR, Dunlop J, and Monaghan M. Amiloride is neuroprotective in an MPTP model of Parkinson’s disease. Neurobiol Dis 31: 334–341, 2008. 7. Askwith CC, Wemmie JA, Price MP, Rokhlina T, and Welsh MJ. Acid-sensing ion channel 2 (ASIC2) modulates ASIC1 H+-activated currents in hippocampal neurons. J Biol Chem 279: 18296–18305, 2004.

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8. Bano D, Young KW, Guerin CJ, Lefeuvre R, Rothwell NJ, Naldini L, Rizzuto R, Carafoli E, and Nicotera P. Cleavage of the plasma membrane Na+/Ca2+ exchanger in excitotoxicity. Cell 120: 275–285, 2005. 9. Baron A, Deval E, Salinas M, Lingueglia E, Voilley N, and Lazdunski M. Protein kinase C stimulates the acid-sensing ion channel ASIC2a via the PDZ domain-containing protein PICK1. J Biol Chem 277: 50463–50468, 2002. 10. Baron A, Waldmann R, and Lazdunski M. ASIC-like, proton-activated currents in rat hippocampal neurons. J Physiol 539: 485–494, 2002. 11. Bassilana F, Champigny G, Waldmann R, de Weille JR, Heurteaux C, and Lazdunski M. The acid-sensitive ionic channel subunit ASIC and the mammalian degenerin MDEG form a heteromultimeric H+-gated Na+ channel with novel properties. J Biol Chem 272: 28819–28822, 1997. 12. Bassler EL, Ngo-Anh TJ, Geisler HS, Ruppersberg JP, and Grunder S. Molecular and functional characterization of acid-sensing ion channel (ASIC) 1b. J Biol Chem 276: 33782–33787, 2001. 13. Benson CJ, Eckert SP, and McCleskey EW. Acid-evoked currents in cardiac sensory neurons: A possible mediator of myocardial ischemic sensation. Circ Res 84: 921–928, 1999. 14. Benson CJ, Xie J, Wemmie JA, Price MP, Henss JM, Welsh MJ, and Snyder PM. Heteromultimers of DEG/ENaC subunits form H+-gated channels in mouse sensory neurons. Proc Natl Acad Sci USA 99: 2338–2343, 2002. 15. Benveniste M and Dingledine R. Limiting stroke-induced damage by targeting an acid channel. N Engl J Med 352: 85–86, 2005. 16. Cadiou H, Studer M, Jones NG, Smith ES, Ballard A, McMahon SB, and McNaughton PA. Modulation of acid-sensing ion channel activity by nitric oxide. J Neurosci 27: 13251–13260, 2007. 17. Canessa CM, Schild L, Buell G, Thorens B, Gautschi I, Horisberger JD, and Rossier BC. Amiloride-sensitive epithelial Na+ channel is made of three homologous subunits. Nature 367: 463–467, 1994. 18. Chai S, Li M, Lan J, Xiong ZG, Saugstad JA, and Simon RP. A kinase-anchoring protein 150 and calcineurin are involved in regulation of acid-sensing ion channels ASIC1a and ASIC2a. J Biol Chem 282: 22668–22677, 2007. 19. Champigny G, Voilley N, Waldmann R, and Lazdunski M. Mutations causing neurodegeneration in Caenorhabditis elegans drastically alter the pH sensitivity and inactivation of the mammalian H+-gated Na+ channel MDEG1. J Biol Chem 273: 15418–15422, 1998. 20. Chang SYLM, Li TF, Chu XP, Lan JQ, Thomson S, Jessick V, Meller R, Simon RP, and Xiong ZG. Involvement of acid-sensing ion channels in the generation of epileptic seizure activity. Society for Neuroscience. San Diego, Ca: 2007. 21. Chen CC, England S, Akopian AN, and Wood JN. A sensory neuron-specific, protongated ion channel. Proc Natl Acad Sci USA 95: 10240–10245, 1998. 22. Chen CC, Zimmer A, Sun WH, Hall J, Brownstein MJ, and Zimmer A. A role for ASIC3 in the modulation of high-intensity pain stimuli. Proc Natl Acad Sci USA 99: 8992–8997, 2002. 23. Chen X, Kalbacher H, and Grunder S. The tarantula toxin psalmotoxin 1 inhibits acidsensing ion channel (ASIC) 1a by increasing its apparent H+ affinity. J Gen Physiol 126: 71–79, 2005. 24. Chesler M. The regulation and modulation of pH in the nervous system. Prog Neurobiol 34: 401–427, 1990. 25. Chesler M and Kaila K. Modulation of pH by neuronal activity. Trends Neurosci 15: 396–402, 1992. 26. Coghlan VM, Perrino BA, Howard M, Langeberg LK, Hicks JB, Gallatin WM, and Scott JD. Association of protein kinase A and protein phosphatase 2B with a common anchoring protein. Science 267: 108–111, 1995.

170

G. Pignataro

27. Colledge M, Dean RA, Scott GK, Langeberg LK, Huganir RL, and Scott JD. Targeting of PKA to glutamate receptors through a MAGUK-AKAP complex. Neuron 27: 107–119, 2000. 28. Coryell MW, Ziemann AE, Westmoreland PJ, Haenfler JM, Kurjakovic Z, Zha XM, Price M, Schnizler MK, and Wemmie JA. Targeting ASIC1a reduces innate fear and alters neuronal activity in the fear circuit. Biol Psychiatry 62: 1140–1148, 2007. 29. Diochot S, Baron A, Rash LD, Deval E, Escoubas P, Scarzello S, Salinas M, and Lazdunski M. A new sea anemone peptide, APETx2, inhibits ASIC3, a major acidsensitive channel in sensory neurons. Embo J 23: 1516–1525, 2004. 30. Dorofeeva NA, Barygin OI, Staruschenko A, Bolshakov KV, and Magazanik LG. Mechanisms of non-steroid anti-inflammatory drugs action on ASICs expressed in hippocampal interneurons. J Neurochem 106: 429–441, 2008. 31. Dube GR, Lehto SG, Breese NM, Baker SJ, Wang X, Matulenko MA, Honore P, Stewart AO, Moreland RB, and Brioni JD. Electrophysiological and in vivo characterization of A-317567, a novel blocker of acid sensing ion channels. Pain 117: 88–96, 2005. 32. Duggan A, Garcia-Anoveros J, and Corey DP. The PDZ domain protein PICK1 and the sodium channel BNaC1 interact and localize at mechanosensory terminals of dorsal root ganglion neurons and dendrites of central neurons. J Biol Chem 277: 5203–5208, 2002. 33. Ekholm A, Kristian T, and Siesjo BK. Influence of hyperglycemia and of hypercapnia on cellular calcium transients during reversible brain ischemia. Exp Brain Res 104: 462–466, 1995. 34. Escoubas P, De Weille JR, Lecoq A, Diochot S, Waldmann R, Champigny G, Moinier D, Menez A, and Lazdunski M. Isolation of a tarantula toxin specific for a class of protongated Na+ channels. J Biol Chem 275: 25116–25121, 2000. 35. Eskandari S, Snyder PM, Kreman M, Zampighi GA, Welsh MJ, and Wright EM. Number of subunits comprising the epithelial sodium channel. J Biol Chem 274: 27281–27286, 1999. 36. Ettaiche M, Deval E, Cougnon M, Lazdunski M, and Voilley N. Silencing acid-sensing ion channel 1a alters cone-mediated retinal function. J Neurosci 26: 5800–5809, 2006. 37. Faux MC and Scott JD. Regulation of the AKAP79-protein kinase C interaction by Ca2+/Calmodulin. J Biol Chem 272: 17038–17044, 1997. 38. Firsov D, Gautschi I, Merillat AM, Rossier BC, and Schild L. The heterotetrameric architecture of the epithelial sodium channel (ENaC). Embo J 17: 344–352, 1998. 39. Friese MA, Craner MJ, Etzensperger R, Vergo S, Wemmie JA, Welsh MJ, Vincent A, and Fugger L. Acid-sensing ion channel-1 contributes to axonal degeneration in autoimmune inflammation of the central nervous system. Nat Med 13: 1483–1489, 2007. 40. Gao J, Duan B, Wang DG, Deng XH, Zhang GY, Xu L, and Xu TL. Coupling between NMDA receptor and acid-sensing ion channel contributes to ischemic neuronal death. Neuron 48: 635–646, 2005. 41. Garcia-Anoveros J, Derfler B, Neville-Golden J, Hyman BT, and Corey DP. BNaC1 and BNaC2 constitute a new family of human neuronal sodium channels related to degenerins and epithelial sodium channels. Proc Natl Acad Sci USA 94: 1459–1464, 1997. 42. Glantz SB, Amat JA, and Rubin CS. cAMP signaling in neurons: patterns of neuronal expression and intracellular localization for a novel protein, AKAP 150, that anchors the regulatory subunit of cAMP-dependent protein kinase II beta. Mol Biol Cell 3: 1215–1228, 1992. 43. Grunder S, Geissler HS, Bassler EL, and Ruppersberg JP. A new member of acid-sensing ion channels from pituitary gland. Neuroreport 11: 1607–1611, 2000. 44. Hansen AJ and Zeuthen T. Extracellular ion concentrations during spreading depression and ischemia in the rat brain cortex. Acta Physiol Scand 113: 437–445, 1981. 45. Helmlinger G, Yuan F, Dellian M, and Jain RK. Interstitial pH and pO2 gradients in solid tumors in vivo: high-resolution measurements reveal a lack of correlation. Nat Med 3: 177–182, 1997.

Acid-Sensing Ion Channels (ASICs): New Targets in Stroke Treatment

171

46. Hruska-Hageman AM, Wemmie JA, Price MP, and Welsh MJ. Interaction of the synaptic protein PICK1 (protein interacting with C kinase 1) with the non-voltage gated sodium channels BNC1 (brain Na+ channel 1) and ASIC (acid-sensing ion channel). Biochem J 361: 443–450, 2002. 47. Jasti J, Furukawa H, Gonzales EB, and Gouaux E. Structure of acid-sensing ion channel 1 at 1.9 A resolution and low pH. Nature 449: 316–323, 2007. 48. Klauck TM, Faux MC, Labudda K, Langeberg LK, Jaken S, and Scott JD. Coordination of three signaling enzymes by AKAP79, a mammalian scaffold protein. Science 271: 1589–1592, 1996. 49. Kleyman TR and Cragoe EJ Jr.. Amiloride and its analogs as tools in the study of ion transport. J Membr Biol 105: 1–21, 1988. 50. Krishtal O. The ASICs: signaling molecules? Modulators? Trends Neurosci 26: 477–483, 2003. 51. Leonard AS, Yermolaieva O, Hruska-Hageman A, Askwith CC, Price MP, Wemmie JA, and Welsh MJ. cAMP-dependent protein kinase phosphorylation of the acid-sensing ion channel-1 regulates its binding to the protein interacting with C-kinase-1. Proc Natl Acad Sci USA 100: 2029–2034, 2003. 52. Lilley S, LeTissier P, and Robbins J. The discovery and characterization of a protongated sodium current in rat retinal ganglion cells. J Neurosci 24: 1013–1022, 2004. 53. Lingueglia E, de Weille JR, Bassilana F, Heurteaux C, Sakai H, Waldmann R, and Lazdunski M. A modulatory subunit of acid sensing ion channels in brain and dorsal root ganglion cells. J Biol Chem 272: 29778–29783, 1997. 54. Miesenbock G, De Angelis DA, and Rothman JE. Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature 394: 192–195, 1998. 55. Nedergaard M, Kraig RP, Tanabe J, and Pulsinelli WA. Dynamics of interstitial and intracellular pH in evolving brain infarct. Am J Physiol 260: R581–R588, 1991. 56. Pignataro G, Gala R, Cuomo O, Tortiglione A, Giaccio L, Castaldo P, Sirabella R, Matrone C, Canitano A, Amoroso S, Di Renzo G, and Annunziato L. Two sodium/ calcium exchanger gene products, NCX1 and NCX3, play a major role in the development of permanent focal cerebral ischemia. Stroke 35: 2566–2570, 2004. 57. Pignataro G, Simon RP, and Xiong ZG. Prolonged activation of ASIC1a and the time window for neuroprotection in cerebral ischaemia. Brain 130: 151–158, 2007. 58. Pignataro G, Tortiglione A, Scorziello A, Giaccio L, Secondo A, Severino B, Santagada V, Caliendo G, Amoroso S, Di Renzo G, and Annunziato L. Evidence for a protective role played by the Na+/Ca2+ exchanger in cerebral ischemia induced by middle cerebral artery occlusion in male rats. Neuropharmacology 46: 439–448, 2004. 59. Price MP, McIlwrath SL, Xie J, Cheng C, Qiao J, Tarr DE, Sluka KA, Brennan TJ, Lewin GR, and Welsh MJ. The DRASIC cation channel contributes to the detection of cutaneous touch and acid stimuli in mice. Neuron 32: 1071–1083, 2001. 60. Price MP, Thompson RJ, Eshcol JO, Wemmie JA, and Benson CJ. Stomatin modulates gating of acid-sensing ion channels. J Biol Chem 279: 53886–53891, 2004. 61. Reeh PW and Steen KH. Tissue acidosis in nociception and pain. Prog Brain Res 113: 143–151, 1996. 62. Rehncrona S. Brain acidosis. Ann Emerg Med 14: 770–776, 1985. 63. Renard S, Lingueglia E, Voilley N, Lazdunski M, and Barbry P. Biochemical analysis of the membrane topology of the amiloride-sensitive Na+ channel. J Biol Chem 269: 12981–12986, 1994. 64. Salinas M, Rash LD, Baron A, Lambeau G, Escoubas P, and Lazdunski M. The receptor site of the spider toxin PcTx1 on the proton-gated cation channel ASIC1a. J Physiol 570: 339–354, 2006. 65. Saugstad JA, Roberts JA, Dong J, Zeitouni S, and Evans RJ. Analysis of the membrane topology of the acid-sensing ion channel 2a. J Biol Chem 279: 55514–55519, 2004.

172

G. Pignataro

66. Schild L, Schneeberger E, Gautschi I, and Firsov D. Identification of amino acid residues in the alpha, beta, and gamma subunits of the epithelial sodium channel (ENaC) involved in amiloride block and ion permeation. J Gen Physiol 109: 15–26, 1997. 67. Siesjo BK, Katsura K, and Kristian T. Acidosis-related damage. Adv Neurol 71: 209–233, 1996. 68. Sluka KA, Price MP, Breese NM, Stucky CL, Wemmie JA, and Welsh MJ. Chronic hyperalgesia induced by repeated acid injections in muscle is abolished by the loss of ASIC3, but not ASIC1. Pain 106: 229–239, 2003. 69. Smith ES, Cadiou H, and McNaughton PA. Arachidonic acid potentiates acidsensing ion channels in rat sensory neurons by a direct action. Neuroscience 145: 686–698, 2007. 70. Sutherland SP, Benson CJ, Adelman JP, and McCleskey EW. Acid-sensing ion channel 3 matches the acid-gated current in cardiac ischemia-sensing neurons. Proc Natl Acad Sci USA 98: 711–716, 2001. 71. Tombaugh GC and Sapolsky RM. Evolving concepts about the role of acidosis in ischemic neuropathology. J Neurochem 61: 793–803, 1993. 72. Voilley N, de Weille J, Mamet J, and Lazdunski M. Nonsteroid anti-inflammatory drugs inhibit both the activity and the inflammation-induced expression of acid-sensing ion channels in nociceptors. J Neurosci 21: 8026–8033, 2001. 73. Vukicevic M and Kellenberger S. Modulatory effects of acid-sensing ion channels on action potential generation in hippocampal neurons. Am J Physiol Cell Physiol 287: C682–C690, 2004. 74. Waldmann R, Bassilana F, de Weille J, Champigny G, Heurteaux C, and Lazdunski M. Molecular cloning of a non-inactivating proton-gated Na+ channel specific for sensory neurons. J Biol Chem 272: 20975–20978, 1997. 75. Waldmann R, Champigny G, Bassilana F, Heurteaux C, and Lazdunski M. A protongated cation channel involved in acid-sensing. Nature 386: 173–177, 1997. 76. Waldmann R, Champigny G, Voilley N, Lauritzen I, and Lazdunski M. The mammalian degenerin MDEG, an amiloride-sensitive cation channel activated by mutations causing neurodegeneration in Caenorhabditis elegans. J Biol Chem 271: 10433–10436, 1996. 77. Wemmie JA, Askwith CC, Lamani E, Cassell MD, Freeman JH Jr., and Welsh MJ. Acidsensing ion channel 1 is localized in brain regions with high synaptic density and contributes to fear conditioning. J Neurosci 23: 5496–5502, 2003. 78. Wemmie JA, Chen J, Askwith CC, Hruska-Hageman AM, Price MP, Nolan BC, Yoder PG, Lamani E, Hoshi T, Freeman JH Jr., and Welsh MJ. The acid-activated ion channel ASIC contributes to synaptic plasticity, learning, and memory. Neuron 34: 463–477, 2002. 79. Wemmie JA, Price MP, and Welsh MJ. Acid-sensing ion channels: advances, questions and therapeutic opportunities. Trends Neurosci 29: 578–586, 2006. 80. Wong HK, Bauer PO, Kurosawa M, Goswami A, Washizu C, Machida Y, Tosaki A, Yamada M, Knopfel T, Nakamura T, and Nukina N. Blocking acid-sensing ion channel 1 alleviates Huntington’s disease pathology via an ubiquitin-proteasome system-dependent mechanism. Hum Mol Genet 17: 3223–3235, 2008. 81. Wu LJ, Duan B, Mei YD, Gao J, Chen JG, Zhuo M, Xu L, Wu M, and Xu TL. Characterization of acid-sensing ion channels in dorsal horn neurons of rat spinal cord. J Biol Chem 279: 43716–43724, 2004. 82. Xie J, Price MP, Wemmie JA, Askwith CC, and Welsh MJ. ASIC3 and ASIC1 mediate FMRFamide-related peptide enhancement of H+-gated currents in cultured dorsal root ganglion neurons. J Neurophysiol 89: 2459–2465, 2003. 83. Xiong ZG, Zhu XM, Chu XP, Minami M, Hey J, Wei WL, MacDonald JF, Wemmie JA, Price MP, Welsh MJ, and Simon RP. Neuroprotection in ischemia: blocking calciumpermeable acid-sensing ion channels. Cell 118: 687–698, 2004. 84. Xu TL and Xiong ZG. Dynamic regulation of acid-sensing ion channels by extracellular and intracellular modulators. Curr Med Chem 14: 1753–1763, 2007.

Acid-Sensing Ion Channels (ASICs): New Targets in Stroke Treatment

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85. Yagi J, Wenk HN, Naves LA, and McCleskey EW. Sustained currents through ASIC3 ion channels at the modest pH changes that occur during myocardial ischemia. Circ Res 99: 501–509, 2006. 86. Yermolaieva O, Leonard AS, Schnizler MK, Abboud FM, and Welsh MJ. Extracellular acidosis increases neuronal cell calcium by activating acid-sensing ion channel 1a. Proc Natl Acad Sci USA 101: 6752–6757, 2004. 87. Ziemann AE, Schnizler MK, Albert GW, Severson MA, Howard MA 3rd, Welsh MJ, and Wemmie JA. Seizure termination by acidosis depends on ASIC1a. Nat Neurosci 11: 816–822, 2008.

Role of TRPM7 in Ischemic CNS Injury Michael F. Jackson*, Hong-Shuo Sun*, Michael Tymianski and John F. MacDonald

1 Introduction One of the key features noted in animal models of stroke is the progressive nature of the excitotoxic cascade. While the excessive release of glutamate and consequent overactivation of NMDARs occurs rapidly (time span of minutes to hours), the ensuing neuronal death has been noted to progress with some delay, the phenomena being appropriately referred to as delayed neuronal death (DND). Surprisingly, given the massive release of glutamate and strong NMDAR activation, during the early phases of the excitotoxic cascade, neurons are initially capable of regulating and maintaining intracellular Ca2+ near physiological levels. Only with some delay do neurons lose the ability to regulate Ca2+. This delayed rise in intracellular Ca2+ is invariably insensitive to treatment with antiexcitotoxic therapies (AETs), consisting of glutamate receptor and Ca2+ channel blockers [1–3]. The failure of AETs to prevent DND coupled with their inability to provide neuroprotection in clinical trials has led our research groups to seek additional Ca2+ influx pathways in the hopes of identifying previously overlooked sources. Our recent studies have led to the demonstration of the important contribution of TRPM7 channels, the focus of present chapter, to neuronal cell death.

2 Gene Structure The human TRPM7 gene spans 126.5 kb, is encoded by 39 exons and has been mapped to chromosome 15 in the q21.2 region, a locus linked to a form of autosomal recessive familial amyotrophic lateral sclerosis (ALS)[4]. The mouse *

These authors contributed equally to this work.

J.F. MacDonald (*) Robarts Research Institute, University of Western Ontario, 100 Perth Drive, London, Ontario, N6A 5K8, Canada e-mail: [email protected]

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homolog, located on chromosome 2 on cytoband F2, is also encoded by 39 exons spanning 84.6 kb. Human TRPM7 is 95% identical to its mouse counterpart [5]. In mouse, TRPM7 encodes a single transcript of 7.1 kb. Interestingly, a recent study suggests that a rearrangement of chromosome 15q21.2 in humans results in the generation of novel chimeric transcript consisting of exon 1 of TRPM7 spliced to the common acceptor splice site in exon 2 of the aromatase CYP19. The resulting aberrant CYP19 expression, now under control of the TRPM7 promoter, underlies androgen excess syndrome in a Russian kindred [6].

3 Structural Features Full length TRPM7 transcripts (mouse) encode a protein of 1863 amino acids having a predicted molecular weight of 212.4 kD. Kyte-Doolittle analysis reveals the presence of seven hydrophobic domains, a common finding among TRP channels [7]. However, only six of these represent membranespanning domains (Fig. 1). Both the N- and C-terminus are located intracellularly and a re-entrant loop, located between the fifth and sixth TM domains, contributes to the formation of the channel pore. By analogy with

Fig. 1 TRPM7 transmembrane topology. MHR denotes regions of high homology conserved among TRPM family members. In addition to the six identified transmembrane domains (TM1-6) a seventh hydrophobic domain (H1) has been identified within the N-terminus. The channel pore is formed by the P-loop, located between TMs 5 and 6. TRP, refers to the TRP domain, a highly conserved region of about 25 amino acids. CCD, illustrates the coiled-coil domain and a-kinase represents the enzyme domain of TRPM7

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K+ channels, TRPM7 channels are thought to form from four subunits (reviewed by [8]), either as homotetramers or as heterotetramers with TRPM6. Several distinct features have been recognized within the large N- and Cterminal regions of TRPM7. For example, the additional hydrophobic region alluded to earlier, and termed H1, localizes to the intracellular N-terminus. Ankyrin repeats, found within the N-terminus of many TRP family members are absent from TRPM7. Rather, TRPM family members’ share four regions of high homology (MHRs) within their N-terminal domains. The functional consequences of structural features within the N-terminus remain largely undefined. In contrast, the C-terminus of TRPM7 has garnered considerable interest and attention due to the identification within this region of an atypical serine/threonine protein kinase domain. Resolution of the enzyme domain structure has allowed the catalytic core to be identified as resembling that of classical protein kinases and metabolic enzymes with ATP-grasp folds [9]. This enzyme domain, homologous to a family of a-kinases [10], appears to be important for the regulation of channel function by Mg2+-nucleotides [11, 12]. In addition to the catalytic domain, a coiled-coil domain and a TRP box (amino acids VWKYQR), both conserved among TRPM family members, have been identified within the C-terminal region of TRPM7. The function of these domains has not been examined in TRPM7. However, evidence from related family members suggests that the coiled-coil domain, a common protein–protein interaction motif, is likely to be involved in channel assembly [13, 14]. A recent crystallography study has uncovered that the coiled-coil domain of TRPM7 consists of an unexpected four-stranded antiparallel arrangement [15]. Further analysis of the coiled-coil structure should facilitate the elucidation of TRPM7 channel assembly determinants. Finally, with regards to the TRP box, a recent study of TRPM8, TRPM5 and TRPV5 has suggested that this region may serve as an interacting site for PIP2, a positive regulator of several TRP channels [16], including TRPM7.

4 Molecular Biology TRPM7 was independently cloned by three groups using distinct strategies. Clapham’s group [17] performed a yeast two-hybrid screen of a rat brain library using the C2 domain of PLC-b1. In contrast, Ryazanov’s group [18] used a homology-based approach seeking to identify additional a-kinases family members. While Fleig’s group [5] used a bioinformatics approach in an attempt to indentify novel Ca2+ entry pathways in lymphocytes. The common identification of TRPM7 by these disparate approaches reflect the somewhat unusual nature of TRPM7 being at once both ion channel and enzyme. Intriguingly, a splice variant, lacking the internal exons that code for the TM domains, has been identified for TRPM7 [19]. A physiological function for this variant, consisting of only the N-terminus and kinase domain, has yet to be identified.

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5 Tissue and Cellular Distribution The original reports of TRPM7 cloning provided evidence for its widespread distribution [5, 17]. Subsequent Northern and RT-PCR analysis of various tissue samples have failed to identify a region in which transcripts for TRPM7 could not be detected. In humans, high expression levels were noted in heart, testis, and prostate [5] and in mouse, high levels were found in kidney, heart, liver, and spleen [17]. More recent studies, using quantitative PCR, confirm the widespread distribution of TRPM7 in both human [20] and mouse [21] tissues including the brain. Studies exploring the cellular distribution, suggests that TRPM7 is strongly expressed at the plasma membrane in a variety of cell types [22–24]. Our own work shows diffuse staining at the surface of hippocampal neurons (Fig. 2) with no evidence of clustering or preferential expression at synapses [25]. In contrast, TRPM7 is exclusively localized within cholinergic vesicles at the neuromuscular junction and in superior cervical ganglion neurons [26].

Fig. 2 TRPM7 expression in hippocampal CA1 region. Confocal images from coronal section of adult rat hippocampus shows that TRPM7 channels are expressed in the CA1 region (left panel). The same section is also counterstained with neuronal marker anti-NeuN antibody (right panel) showing that TRPM7 is expressed in neurons of hippocampal CA1 area. Coronal slices were sectioned into 50 mm thickness using cryostat. Images were taken using the Zeiss LSM 510 META Confocal Laser Scanning Microscope and optical stacks of 10 images from images taken at 1 mm internals through the region of interest were used for the final figures Scale bars are 100 mm for Confocal images taken with 10 power objective

6 Biophysical and Electrophysiological Properties As with many TRP family members, TRPM7 is a non-selective cation channel. Beyond this commonality, TRPM7 possesses several distinguishing biophysical features. Especially important is the relation between monovalent cation influx

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and the extracellular divalents, Ca2+ and Mg2+. This is best illustrated by considering the characteristics of the current–voltage (I–V) relation for TRPM7 (Fig. 3). In the presence of physiological concentrations of these extracellular divalents, the I–V curve for TRPM7 displays prominent outward rectification, which can entirely be attributed to a voltage-dependent block of monovalent cation influx by extracellular divalents. Indeed, in the absence of divalents, the resulting I–V relation is quasi linear and reverses at 0 mV, reflecting the absence of both voltage-dependent gating and channel selectivity with regards to Na+ or K+. Importantly, though monovalent influx is blocked, divalent entry nevertheless occurs, a phenomenon referred to as permeation block. Furthermore, and in contrast to most other Ca2+ channels, TRPM7 is more permeable to a series of divalents, including potentially toxic trace metals, than it is to Ca2+ or Mg2+. The permeability sequence is reportedly the following: Zn2+ Ni2+ >> Ba2+ > Co2+ > Mg2+ > Mn2+ > Sr2+ > Cd2+ > Ca2+ [27]. Though the resulting currents are admittedly small or even un-resolvable, divalent entry can be observed in experiments using fluorescent dyes. In this way, Monteilh-Zoller et al [27] elegantly demonstrated that TRPM7 allows entry of divalents, including Ca2+, Mn2+, Co2+, and Ni2+, despite the presence of physiological levels of extracellular Ca2+ and Mg2+. Admittedly, neurons possess numerous influx pathways for trace metal ions, most notably voltage-gated Ca2+ and NMDAR channels. However, unlike 2+

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Fig. 3 Representative traces of TRPM7-mediated currents recorded from an acutely isolated CA1 hippocampal pyramidal neuron when lowering extracellular Ca2+ from 1.3 mM to 0.1 mM. Top left trace illustrates the timing of the solution changes. Middle left trace, representative example of the increase in TRPM7-mediated inward current generated when the concentration of extracellular Ca2+ is reduced. Bottom left trace, voltage protocol illustrating the timing of voltage ramps (80 mV, 500 ms). The resulting I–V relations, generated at time points A and B, are shown on the right

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these channels, which are normally quiescent until activated (i.e., by voltage or by agonist), TRPM7 is constitutively active. Consequently, though modulated by several extracellular and intracellular factors (e.g., ROS/RNS, pH, and intracellular Mg2+, described in the following section), the current flowing through these channels at any given time is predominantly limited by the extracellular concentration of extracellular divalents. It is likely this feature, which allows TRPM7 to play an important part in maintaining intracellular Mg2+ homeostasis and also to ‘‘sense’’ changes in the extracellular concentration of divalents [25]. Such changes occur during intense neuronal activity associated, for example, with seizure and especially ischemic episodes.

7 Regulatory Mechanisms: Receptorial, Transcriptional, and Transductional In addition to regulating channel activity from the extracellular space, Mg2+ plays an important role in regulating TRPM7 intracellularly. Previous studies have clearly demonstrated that intracellular Mg2+ inhibits TRPM7 channels. Consequently, experimental conditions which reduce the intracellular concentration of this divalent cation (e.g., intracellularly applied Mg2+ chelators such as HEDTA or Na-ATP) facilitate TRPM7-mediated currents. More controversial in whether or not the intracellular block is mediated solely by Mg2+ or rather by Mg2+ nucleotides [12, 28]. Studies in which the kinase activity of the channel was modified (or entirely removed) by mutation suggest that nucleotide binding to this domain may act as a gating modifier by altering the sensitivity of the channel to inhibition by Mg2+ at a distinct site. The demonstration of an interaction between PLC-b1 and TRPM7 led Clapham’s group to explore the functional consequence of this interaction. They demonstrated that the interaction allows the concentration of phosphatidylinositol 4,5-bisphosphate (PIP2) to be regulated in the vicinity of the channel. In this way, Gaq-linked G-protein receptors that activate PLC inhibit TRPM7 through the hydrolysis of PIP2 [29]. However, a recent report [30] proposes a more complex regulation by PLC-coupled receptors. Using less invasive perforated-patch recordings, in cells expressing TRPM7 at modest levels, the authors reported that PLC-coupled receptor agonists can enhance rather than inhibit TRPM7 function. They suggest that the resting intracellular concentration of Mg2+, which may be influenced by the recording configuration used (i.e., perforated patch versus whole-cell recording techniques), may determine the outcome of PLC stimulation. TRPM7 may also be regulated through phosphorylation. Interestingly, the closely related TRPM family TRPM6, which may form heteromeric channels with TRPM7, can phosphorylate TRPM7 at threonine residues [31]. A recent study identified a variant of TRPM7 in a subset of Guamanian amyotrophic lateral sclerosis and parkinsonism dementia patients [32]. The variant harbored a missense mutation (T1482I) associated with reduced phosphorylation and

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increased sensitivity to inhibition by Mg2+. Though the specific sites have yet to be identified, these studies suggest that phosphorylation by TRPM6 could conceivably play an important role in regulating TRPM7 by altering the channel sensitivity to Mg2+.

8 Physiological Properties To date, most of the available information for the physiological functions of mammalian TRPM7 channel is drawn from cellular studies. There are unconfirmed reports that TRPM7 knockout is embryonically lethal. At the cellular level, TRPM7 overexpression in heterologous cell systems (e.g., HEK-293) has a detrimental effect on cell survival [5]. Normally adherent HEK-293 cells detach and ultimately die off upon TRPM7 overexpression [5]. Two other studies [33, 34] also confirmed that TRPM7 overexpression disrupts cell adhesion. Both studies indicated the involvement of Ca2+-dependent mechanisms. Paradoxically, TRPM7 knockout in avian B cells caused cell death indicating its vital role in cell survival [11]. TRPM7 appears to have vital role in Mg2+ homeostasis in DT-40 cells as high Mg2+-containing medium, but not high Ca2+, can restore normal cell growth and proliferation in TRPM7-deficient cells. The diffuse expression of TRPM7 suggests that it may function as a regulator for cellular Mg2+ homeostasis in numerous physiological systems [35, 36]. In addition to Mg2+, the TRPM7 channels contribute to Ca2+ influx into cells and thereby may regulate cell proliferation and G1/S cell cycle progression [24]. Similarly, TRPM7 orthologs have been identified in non-mammalian systems such as zebra fish and Caenorhabditis elegans [37] and knockdown of these channels affects cell growth and proliferation [38]. Also, in C. elegans, the TRPM7 ortholog has been implicated as a component of the defecation rhythm generator [39]. A recent study suggests a similar role for TRPM7 in mammals [23].

9 Pathophysiological Relevance in Stroke Numerous clinical trials of AETs have failed to benefit stroke patients [40]. To date, there is still no clinical or experimental treatment that can improve the outcome of stroke patients, with the exception of the thrombolytic agent, tissue plasminogen activator (tPA). This remains true despite significant advances in our understanding of ischemic mechanisms over the last two decades. A few years ago, we described two key mechanisms that cause neuronal injury when neurons are challenged by anoxia or excitotoxins. The first mechanism involves the activation of signaling pathways associated with NMDA glutamate receptors (NMDARs) through submembrane scaffolding proteins such as PSD-95 [41–43]. The second mechanism is through the activation of TRPM7 channels [25, 44]. The Ca2+ influx contributed by these channels is a major contributor to

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the neuronal damage observed in hypoxic and ischemic conditions. Since TRPM7 is widely expressed in different tissues and systems including brain, heart, kidney, and blood vessels it is likely that these channels may play an important role not only in cerebral ischemia but also in ischemic injury in the heart and kidney, for example. Our group [44] demonstrated a role for TRPM7 in neurons in an experimental model of ischemia. TRPM7 was shown to contribute to hypoxic neuronal death in rodent cultured cortical neurons subjected to prolong oxygen and glucose deprivation (OGD). In this model, Ca2+ influx through TRPM7 channels was correlated with neuronal cell death and suppression of TRPM7 expression, using siRNA, inhibited an OGD-evoked current and protected neurons against OGD-induced neuronal death. Interestingly, in the context of the recruitment of TRPM7 during ischemia, TRPM7 can be positively modulated by both protons [45] and by ROS/RNS [44]. As a result, the decrease in extracellular divalents, increase in extracellular proton concentration and the generation of ROS/RNS that occur coincidently during ischemia will act in concert to powerfully recruit TRPM7 channels (Fig. 4).

Fig. 4 Model of TRPM7 activation during cerebral ischemia. Strong NMDA receptor activation during the initial stages of an ischemic attack leads to a large influx of Ca2+. The resulting Ca2+-dependent stimulation of NOS, tethered to NMDARs through PSD-95, and O2– production from mitochondria leads to the production of ONOO–, which stimulates TRPM7 channels. The reduction of extracellular divalents and decrease in pH, both associated with ischemic episodes, further enhance TRPM7 activation. These factors act in concert to stimulate additional Ca2+ influx through TRPM7 channels leading to a potentially toxic positive feedback

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In addition to contributing to the Ca2+ loading of neurons, the permeability of TRPM7 to toxic metal ions (e.g., Zn2+, Ni+2, and Cd2+) may also play a role. In particular, Zn2+ is highly toxic to cells if its concentration exceeds the trace level. In this way, the Zn2+ permeability of TRPM7 may contribute not only to acute neurotoxicity in ischemia [46, 47] but may also be a factor in the pathogenesis of chronic neurodegenerative conditions like Alzheimer’s disease where Zn2+ has been reported to accumulate in amyloid plaques [48]. Despite compelling in vitro evidence, the role of TRPM7 in stroke will need to be clarified in in vivo studies. Indeed, even though the molecular and cellular illustrations outlined above are crucial, hypoxia or OGD models are no substitutes for ischemic models in whole animals. Events associated with ischemic episodes in vivo may not be faithfully paralleled in the in vitro hypoxic setting [49]. Correspondingly, there is a need to study animal ischemia models to confirm the role of TRPM7 in stroke.

10 Pharmacological Modulation To date, there are no selective agonists and antagonists for TRPM7 channels. Accordingly, we and others have used gene silencing by small interfering RNA (siRNA) to knock down TRPM7 in CNS neurons [25, 44], PNS neurons [26], vascular smooth muscle cells [50], interstitial pacemaker cells of Cajal in the gastrointestinal tract [23], and human epithelial cells [51]. Despite the usefulness of these powerful approaches, there is a pressing need for the development of specific pharmacological agents for studying TRPM7 function.

11 Preliminary Clinical Trials As there are no studies of TRPM7 function in vivo and no specific pharmacological blockers available, we do not expect to see any preliminary clinical trials in the near future. Until drug screening efforts for TRPM7 blockers are successful, siRNA-induced gene silencing may be a potential approach for stroke treatment, as gene therapy has been utilized in other systems [52]. Other approaches aimed at modulating TRPM7 by interrupting specific signaling pathways, as we did in interrupting the NMDA receptors and downstream signal pathway PSD95 [41, 42], may also prove useful.

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12 Therapeutic Perspectives In animal studies, the most effective way to reduce stroke damage is by interrupting the injury process before any damage becomes irreversible. To date the most promising strategies for stroke treatment involves attempts either to improve perfusion using thrombolytic agents (such as tPA) or to limit the injury area using so-called neuroprotective agents. Using tPA is likely able to reduce damage to neurons by re-establishing the blood flow to the ischemic region and this drug is the only treatment currently available for treatment of acute cerebral stroke [53]. Approved by the FDA in 1996, tPA was initially developed for the treatment of heart attack. In spite of being the only approved drug therapy for stroke, less than 1% of stroke patients benefit from tPA because of its limited 3-h therapeutic window (if applied by i.v. administration). Treatment within 90 min of the onset of cerebral stroke maximizes the potential benefit and minimizes the potential risks of tPA. The most dangerous risk is of excess cerebral bleeding from the inappropriate use of tPA, which further complicates the problem of cerebral ischemia. There are also many other limitations for tPA treatment: (1) only 5% of stroke patients are able to make it to a hospital within the 3h therapeutic window; (2) a brain scan must be performed to rule out any sign of intracranial cerebral bleeding; (3) there is an increased risk of intracerebral hemorrhage; and (4) there is a lack of trained personnel available to perform the pretreatment assessment together with tPA administration. In the end, only 1–2% of stroke patients are treated with tPA and, of those, only 30–35% show any beneficial effect of the tPA treatment. Thus, it is of great importance to identify effective neuroprotective medicines and therapeutic measures in order to protect the brain and improve recovery in patients suffering a stroke. In the future, TRPM7 channel modulators might be used in combination therapy for the treatment of cerebral ischemic stroke. Combination therapy for ischemic stroke would ideally help restore cerebral blood flow, while selected pharmacological agents, targeting select targets and applied at empirically determined time points of stroke onset, would provide neuroprotection and potentially facilitate recovery of function (Fig. 5). One potential caveat is that the widespread expression of TRPM7 may limit the selectivity of pharmacological agents developed to target this channel. Since reductions in extracellular divalents are associated with ischemic episodes, it may be possible to improve upon the selectivity by developing activity-dependent blockers that would preferentially target TRPM7 channels strongly recruited during stroke.

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Fig. 5 Ischemic cell death cascade. The recruitment of NMDARs is central to the excitotoxic model of ischemic cell death. The failure of NMDAR antagonists to provide protection in clinical stroke trials has forced a re-evaluation of this model. We propose that the contribution of NMDARs, though critical as an early trigger of death cascades, is limited in time to a brief period immediately following a stroke. Our hypothesis is that the progressive recruitment of additional non-selective cation channels plays an equal, if not greater role, in mediating neuronal death. Rational polytherapy combining drugs that target various contributors of neuronal death may prove to be the most effective means of providing neuroprotection for stroke patients

Conclusions Knockdown of TRPM7 in vitro using siRNA gene silencing has been shown to provide neuroprotection in in vitro models pointing toward an alternative therapeutic approach to stroke treatment. Future studies will likely proceed via a rational polytherapy for stroke treatment involving pharmacological agents that target different ion channels, including TRPM7, receptors and exchangers, as well as acting to restore cerebral blood flow.

References 1. Manev H, Favaron M, Guidotti A, and Costa E. Delayed increase of Ca2+ influx elicited by glutamate: role in neuronal death. Mol Pharmacol 36: 106–112, 1989. 2. Tymianski M, Charlton MP, Carlen PL, and Tator CH. Secondary Ca2+ overload indicates early neuronal injury which precedes staining with viability indicators. Brain Res 607: 319–323, 1993. 3. Randall RD and Thayer SA. Glutamate-induced calcium transient triggers delayed calcium overload and neurotoxicity in rat hippocampal neurons. J Neuropsychiatry 12: 1882–1895, 1992.

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M.F. Jackson et al.

4. Hermosura MC and Garruto RM. TRPM7 and TRPM2-Candidate susceptibility genes for Western Pacific ALS and PD? Biochim Biophys Acta 1772: 822–835, 2007. 5. Nadler MJ, Hermosura MC, Inabe K, Perraud AL, Zhu Q, Stokes AJ, Kurosaki T, Kinet JP, Penner R, Scharenberg AM, and Fleig A. LTRPC7 is a Mg. ATP-regulated divalent cation channel required for cell viability. Nature 411: 590–595, 2001. 6. Tiulpakov A, Kalintchenko N, Semitcheva T, Polyakov A, Dedov I, Sverdlova P, Kolesnikova G, Peterkova V, and Rubtsov P. A potential rearrangement between CYP19 and TRPM7 genes on chromosome 15q21.2 as a cause of aromatase excess syndrome. J Clin Endocrinol Metab 90: 4184–4190, 2005. 7. Vannier B, Zhu X, Brown D, and Birnbaumer L. The membrane topology of human transient receptor potential 3 as inferred from glycosylation-scanning mutagenesis and epitope immunocytochemistry. J Biol Chem 273: 8675–8679, 1998. 8. Clapham DE, Runnels LW, and Strubing C. The TRP ion channel family. Nat Rev Neurosci 2: 387–396, 2001. 9. Yamaguchi H, Matsushita M, Nairn AC, and Kuriyan J. Crystal structure of the atypical protein kinase domain of a TRP channel with phosphotransferase activity. Mol Cell 7: 1047–1057, 2001. 10. Ryazanov AG. Elongation factor-2 kinase and its newly discovered relatives. FEBS Lett 514: 26–29, 2002. 11. Schmitz C, Perraud AL, Johnson CO, Inabe K, Smith MK, Penner R, Kurosaki T, Fleig A, and Scharenberg AM. Regulation of vertebrate cellular Mg2+ homeostasis by TRPM7. Cell 114: 191–200, 2003. 12. Demeuse P, Penner R, and Fleig A. TRPM7 channel is regulated by magnesium nucleotides via its kinase domain. J Gen Physiol 127: 421–434, 2006. 13. Mei ZZ, Xia R, Beech DJ, and Jiang LH. Intracellular coiled-coil domain engaged in subunit interaction and assembly of melastatin-related transient receptor potential channel 2. J Biol Chem 281: 38748–38756, 2006. 14. Tsuruda PR, Julius D, and Minor DL Jr.. Coiled coils direct assembly of a cold-activated TRP channel. Neuron 51: 201–212, 2006. 15. Fujiwara Y and Minor DL Jr.. X-ray Crystal Structure of a TRPM Assembly Domain Reveals an Antiparallel Four-stranded Coiled-coil. J Mol Biol. 383: 854–870, 2008. 16. Rohacs T, Lopes CM, Michailidis I, and Logothetis DE. PI(4,5)P2 regulates the activation and desensitization of TRPM8 channels through the TRP domain. Nat Neurosci 8: 626–634, 2005. 17. Runnels LW, Yue L, and Clapham DE. TRP-PLIK, a bifunctional protein with kinase and ion channel activities. Science 291: 1043–1047, 2001. 18. Riazanova LV, Pavur KS, Petrov AN, Dorovkov MV, and Riazanov AG. Novel type of signaling molecules: protein kinases covalently linked to ion channels. Mol Biol (Mosk) 35: 321–332, 2001. 19. Chubanov V, Schnitzler M, Waring J, Plank A, and Gudermann T. Emerging roles of TRPM6/TRPM7 channel kinase signal transduction complexes. Naunyn Schmiedebergs Arch Pharmacol 371: 334–341, 2005. 20. Fonfria E, Murdock PR, Cusdin FS, Benham CD, Kelsell RE, and McNulty S. Tissue distribution profiles of the human TRPM cation channel family. J Recept Signal Transduct Res 26: 159–178, 2006. 21. Kunert-Keil C, Bisping F, Kruger J, and Brinkmeier H. Tissue-specific expression of TRP channel genes in the mouse and its variation in three different mouse strains. BMC Genomics 7: 159, 2006. 22. Oancea E, Wolfe JT, and Clapham DE. Functional TRPM7 channels accumulate at the plasma membrane in response to fluid flow. Circ Res 98: 245–253, 2006. 23. Kim BJ, Lim HH, Yang DK, Jun JY, Chang IY, Park CS, So I, Stanfield PR, and Kim KW. Melastatin-type transient receptor potential channel 7 is required for intestinal pacemaking activity. Gastroenterology 129: 1504–1517, 2005.

Role of TRPM7 in Ischemic CNS Injury

187

24. Hanano T, Hara Y, Shi J, Morita H, Umebayashi C, Mori E, Sumimoto H, Ito Y, Mori Y, and Inoue R. Involvement of TRPM7 in cell growth as a spontaneously activated Ca2+ entry pathway in human retinoblastoma cells. J Pharmacol Sci 95: 403–419, 2004. 25. Wei WL, Sun HS, Olah ME, Sun X, Czerwinska E, Czerwinski W, Mori Y, Orser BA, Xiong ZG, Jackson MF, Tymianski M, and MacDonald JF. TRPM7 channels in hippocampal neurons detect levels of extracellular divalent cations. Proc Natl Acad Sci USA 104: 16323–16328, 2007. 26. Krapivinsky G, Mochida S, Krapivinsky L, Cibulsky SM, and Clapham DE. The TRPM7 Ion Channel Functions in Cholinergic Synaptic Vesicles and Affects Transmitter Release. Neuron 52: 485–496, 2006. 27. Monteilh-Zoller MK, Hermosura MC, Nadler MJ, Scharenberg AM, Penner R, and Fleig A. TRPM7 provides an ion channel mechanism for cellular entry of trace metal ions. J Gen Physiol 121: 49–60, 2003. 28. Kozak JA and Cahalan MD. MIC channels are inhibited by internal divalent cations but not ATP. Biophys J 84: 922–927, 2003. 29. Runnels LW, Yue L, and Clapham DE. The TRPM7 channel is inactivated by PIP(2) hydrolysis. Nat Cell Biol 4: 329–336, 2002. 30. Langeslag M, Clark K, Moolenaar WH, van Leeuwen FN, and Jalink K. Activation of TRPM7 channels by PLC-coupled receptor agonists. J Biol Chem. 282: 232–239, 2007. 31. Schmitz C, Dorovkov MV, Zhao X, Davenport BJ, Ryazanov AG, and Perraud AL. The channel kinases TRPM6 and TRPM7 are functionally non-redundant. J Biol Chem. 280: 37763–37771, 2005. 32. Hermosura MC, Nayakanti H, Dorovkov MV, Calderon FR, Ryazanov AG, Haymer DS, and Garruto RM. A TRPM7 variant shows altered sensitivity to magnesium that may contribute to the pathogenesis of two Guamanian neurodegenerative disorders. Proc Natl Acad Sci USA 102: 11510–11515, 2005. 33. Su LT, Agapito MA, Li M, Simonson WT, Huttenlocher A, Habas R, Yue L, and Runnels LW. TRPM7 Regulates Cell Adhesion by Controlling the Calcium-dependent Protease Calpain. J Biol Chem 281: 11260–11270, 2006. 34. Clark K, Langeslag M, van Leeuwen B, Ran L, Ryazanov AG, Figdor CG, Moolenaar WH, Jalink K, and van Leeuwen FN. TRPM7, a novel regulator of actomyosin contractility and cell adhesion. EMBO J 25: 290–301, 2006. 35. Fleig A and Penner R. The TRPM ion channel subfamily: molecular, biophysical and functional features. Trends Pharmacol Sci 25: 633–639, 2004. 36. Montell C. Mg2+ homeostasis: the Mg2+nificent TRPM chanzymes. Curr Biol 13: R799–R801, 2003. 37. Estevez AY, Roberts RK, and Strange K. Identification of store-independent and storeoperated Ca2+ conductances in Caenorhabditis elegans intestinal epithelial cells. J Gen Physiol 122: 207–223, 2003. 38. Elizondo MR, Arduini BL, Paulsen J, MacDonald EL, Sabel JL, Henion PD, Cornell RA, and Parichy DM. Defective skeletogenesis with kidney stone formation in dwarf zebrafish mutant for trpm7. Curr Biol 15: 667–671, 2005. 39. Vriens J, Owsianik G, Voets T, Droogmans G, and Nilius B. Invertebrate TRP proteins as functional models for mammalian channels. Pflugers Arch 449: 213–226, 2004. 40. Davis SM, Lees KR, Albers GW, Diener HC, Markabi S, Karlsson G, and Norris J. Selfotel in acute ischemic stroke: possible neurotoxic effects of an NMDA antagonist. Stroke 31: 347–354, 2000. 41. Sattler R, Xiong Z, Lu WY, Hafner M, MacDonald JF, and Tymianski M. Specific coupling of NMDA receptor activation to nitric oxide neurotoxicity by PSD-95 protein. Science 284: 1845–1848, 1999. 42. Aarts M, Liu Y, Liu L, Besshoh S, Arundine M, Gurd JW, Wang YT, Salter MW, and Tymianski M. Treatment of ischemic brain damage by perturbing NMDA receptorPSD-95 protein interactions. Science 298: 846–850, 2002.

188

M.F. Jackson et al.

43. Sun HS, Doucette TA, Liu Y, Fang Y, Teves L, Aarts M, Ryan CL, Bernard PB, Lau A, Forder JP, Salter MW, Wang YT, Tasker RA, and Tymianski M. Effectiveness of PSD95 inhibitors in permanent and transient focal ischemia in the rat. Stroke 39: 2544–2553, 2008. 44. Aarts M, Iihara K, Wei WL, Xiong ZG, Arundine M, Cerwinski W, MacDonald JF, and Tymianski M. A key role for TRPM7 channels in anoxic neuronal death. Cell 115: 863–877, 2003. 45. Jiang J, Li M, and Yue L. Potentiation of TRPM7 inward currents by protons. J Gen Physiol 126: 137–150, 2005. 46. Lee JM, Grabb MC, Zipfel GJ, and Choi DW. Brain tissue responses to ischemia. J Clin Invest 106: 723–731, 2000. 47. Lee JM, Zipfel GJ, Park KH, He YY, Hsu CY, and Choi DW. Zinc translocation accelerates infarction after mild transient focal ischemia. Neuroscience 115: 871–878, 2002. 48. Maynard CJ, Bush AI, Masters CL, Cappai R, and Li QX. Metals and amyloid-beta in Alzheimer’s disease. Int J Exp Pathol 86: 147–159, 2005. 49. Lipton P. Ischemic cell death in brain neurons. Physiol Rev 79: 1431–1568, 1999. 50. He Y, Yao G, Savoia C, and Touyz RM. Transient receptor potential melastatin 7 ion channels regulate magnesium homeostasis in vascular smooth muscle cells: role of angiotensin II. Circ Res 96: 207–215, 2005. 51. Numata T, Shimizu T, and Okada Y. TRPM7 is a stretch- and swelling-activated cation channel involved in volume regulation in human epithelial cells. Am J Physiol Cell Physiol 292: C460–C467, 2007. 52. Zhu T, Zhou L, Mori S, Wang Z, McTiernan CF, Qiao C, Chen C, Wang DW, Li J, and Xiao X. Sustained whole-body functional rescue in congestive heart failure and muscular dystrophy hamsters by systemic gene transfer. Circulation 112: 2650–2659, 2005. 53. Marler JR and Goldstein LB. Medicine. Stroke – tPA and the clinic. Science 301: 1677, 2003.

Subtypes of Voltage-Gated Ca2þ Channels and Ischemic Brain Injury Soon-Tae Lee, Daejong Jeon, and Kon Chu

1 Introduction Intracellular Ca2þ ([Ca2þ]i) initiates physiologic events as diverse as gene transcription, muscle contraction, cell division, and exocytosis. Disruption of Ca2þ regulation in the brain accompanies numerous neurological dysfunctions, including epilepsy and ischemia. Although the metabolic machinery that elicits changes in intracellular Ca2þ concentration is heterogenous, voltage-gated calcium channels (VGCCs) are one of the most important components of neurons and other excitable cells, and they play major roles in both normal functioning and various pathological processes, such as cerebral ischemia, epilepsy, migraine, muscle weakness, and cerebellar ataxia. VGCCs are a group of voltage-gated ion channels found in excitable cells with permeability to the Ca2þ ion. Since VGCCs were first identified in crustacean muscle in 1953 by Paul Fatt and Bernard Katz [29], researchers have identified several different kinds of VGCCs, including L-, N-, P/Q-, R-, and T-type calcium channels. VGCCs are normally closed at physiologic or resting membrane potential. When they are activated (i.e., opened) at depolarized membrane potentials, VGCCs allow Ca2þ to enter the cell, resulting in muscular contraction, excitation of neurons, upregulation of gene expression, activation of Ca2þ-dependent enzyme, neurite growth, or release of neurotransmitters. Given their importance in cerebral ischemia, this review describes the structures, functions, phenotypes of knockout mice, and pathophysiologic relevance of VGCCs in cerebral ischemia.

K. Chu (*) Department of Neurology, Seoul National University Hospital, 101, Daehangno, Jongno-Gu, Seoul, 110-744, South Korea e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_11, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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2 Structure and Subunits of VGCCs VGCCs have multi-subunit structures consisting of a major pore-forming subunit (a1) and several auxiliary subunits (a2–d1–4, b1–4, and g1–8) (Fig. 1). The purification of VGCCs was started from 1,4-dihydropyridines (DHPs)-binding Ca2þ channels, which are highly expressed in T-tubules [15, 52]. The a1 protein was identified as the component that bound to DHPs. Cloning of the cDNA for the DHP-binding channels was accomplished initially from skeletal muscle [121], and subsequently from heart, by homology with the skeletal muscle sequence [80]. The general structure of a calcium channel is biochemically complex and composed of four or five distinct subunits [15]. The various VGCCs are structurally homologous, but not identical. The a1 subunit is the largest subunit (190–250 kDa), and the a1 subunit comprises four homologous (or conserved) domains (I–IV) that are linked by variable cytoplasmic sequences (intracellular connecting loop). Each homologous domain (I–IV) contains six transmembrane a-helix segments (S1–S6), and the S4 segment is the voltage sensor (Fig. 1). The a1 subunit harbors the molecular machinery to produce a functional channel, such as p-loops that permit ion permeation and selectivity, voltage sensors that allow the channel to respond to membrane depolarization, intrinsic inactivation machinery, and the binding sites for all known Ca2þ channel blockers. The pore of VGCCs is

Fig. 1 Structural organization of VGCCs. VGCCs have multi-subunit structures consisting of a major pore-forming subunit (a1) and several auxiliary subunits (a2–d, b, and g) [107]

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exquisitely Ca2þ selective, and this property has been attributed to key negatively charged residues, usually glutamate, in each pore loop [26, 131]. Thus, the a1 subunit pore is the primary subunit required for channel functioning. Most Ca2þ channels contain an intracellular b subunit and a transmembrane, disulfide-linked a2d subunit complex. Among the auxiliary subunits, the b subunits are entirely cytosolic (Fig. 1). Four b subunits (Cavb1–4) have been cloned and each Cavb has distinctive properties [25]. Structurally, Cavb has five different domains, with the two conserved domains sharing significant homology among b subunits. The conserved domains were revealed as a src homology-3 (SH3) domain and a guanylate kinase (GK) domain [18, 41, 90, 125], and thus Cavb is included in membrane-associated guanylate kinase (MAGUK) family that has scaffolding functions. Interestingly, recent several reports suggest that Cavb can bind to other molecules and work without marked influence on VGCCs [6, 8, 32, 39, 46, 47, 59, 99]. Cavb has marked effects on the properties of VGCCs a1 subunits, including trafficking of Ca2þ channel complexes to the plasma membrane, voltage-dependence, and activation/inactivation kinetics of Ca2þ currents [9, 25]. The a2 and d auxiliary subunits are encoded by a single gene, and their polypeptide is post-translationally cleaved into an a2 and a d (Fig. 2). The post-translationally cleaved products form disulfide-linked a2–d. a2 is entirely extracellular and d has a single transmembrane domain, which serves to anchor a2d complex in the plasma membrane. a2 is extensively glycosylated, and this post-translational modification gives the stability of the interaction with a1. Four a2d subunit genes have now been cloned. a2d subunit was also shown to modify the biophysical and pharmacological properties of the a1 subunit [4, 68]. Eight g subunit has four transmembrane domains and eight g subunit genes have now been identified (Fig. 1). The g1 subunit is known to be associated with skeletal muscle VGCC complexes, and other g subunits are expressed in the

Fig. 2 The phylogenetic tree illustrates the amino acid sequence homology of various VGCCs (modified from [28])

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heart and brain [16]. The pharmacological and electrophysiological diversity of VGCCs arises primarily from the presence of multiple a1 subunits, and the remaining auxiliary subunits (b, a2d, g) modify the properties of the channel complex [51].

3 Classification of VGCCs The a1 subunits are encoded by at least 10 distinct genes, called CACNA1 (voltage-gated calcium channel a1). The Ca2þ channel genes were named using the chemical symbol of the principal permeating ion (Ca) with ‘‘voltage’’ indicated as a subscript (CaV) [28]. The numerical identifier corresponds to the CaV channel a1 subunit gene subfamily (1–3) and the order in which the a1 subunit was discovered within its respective subfamily (1–n). According to this nomenclature, L-type Ca2þ currents are mediated by the CaV1 subfamily (CaV1.1–CaV1.4), which includes channels containing CACNA1S, CACNA1C, CACNA1D, and CACNA1F (also referred to as a1S, a1C, a1D and a1F, respectively) (Table 1). P/Q-type, N-type, and R-type Ca2þ currents are mediated by the CaV2 subfamily (CaV2.1–CaV2.3), which includes channels containing CACNA1A, CACNA1B, and CACNA1E (referred to as a1A, a1B, a1E, respectively) (Table 1). T-type Ca2þ currents are mediated by the CaV3 subfamily (CaV3.1–CaV3.3), which includes channels containing CACNA1G, CACNA1H, and CACNA1I (referred to as a1G, a1H and a1I, respectively). This classification of VGCCs is based on electrophysiological and pharmacological properties [15, 28, 52] (Tables 1 and 2). The amino acid sequences of the a1 subunits within a subfamily share more than 70% identity, but they share less than 40% identity among the three subfamilies [16] (Fig. 2). The genes for the various a1 subunits have become widely dispersed in the genome and are not clustered on single chromosomes in mammals.

4 Biophysical and Electrophysiological Properties Researchers classified two current components (low- and high-voltageactivated current) in mammalian sensory neurons according to their biophysical properties, showing different conductances in the single channels [14, 30] (Table 2). At first, pharmacological studies using DHPs defined L-type VGCC with ‘long lasting’ conductance as high-voltage activated (HVA) channels present in skeletal muscle, heart, smooth muscle, and neurons [45]. Pharmacological studies further revealed the presence of non-L-type HVA channels [110]. The DHP-insensitive, non-L-type current component was then subdivided according to its biophysical properties [88] and several irreversible toxins [89]. Two subtypes of non-L-type HVA channels were identified. First, N-type (for non-L and neuronal) channels were blocked by a toxin fraction

SNX-482 None

CACNA1D

CACNA1F

CACNA1A

CACNA1B

CACNA1E

CACNA1G CACNA1H CACNA1I

Cav1.3

Cav1.4

Cav2.1

Cav2.2

Cav2.3

Cav3.1 Cav3.2 Cav3.3

P/Q-type

N-type

R-type

T-type

Specific antagonists

o-Conotoxin-GVIA

o-Agatoxin IVA

Dihydropyridines, phenylalkylamines, benzothiazepines

Cav1.1 Cav1.2

L-type

Protein

CACNA1S CACNA1C

Gated by

Current type Skeletal muscle, transverse tubules. Cardiac myocytes, smooth muscles, endocrine cells, neuronal cell bodies, proximal dendrites Endocrine cells, neuronal cell bodies and dendrites, cardiac atrial cells and pacemaker cells, cochlear hair cells Retinal rod and bipolar cells, spinal cord, adrenal gland, mast cells Purkinje neurons and granule cells in the cerebellum, nerve terminals and dendrites, neuroendocrine cells Throughout the brain, nerve terminals and dendrites, neuroendocrine cells Neuronal bodies and dendrites, cerebellar granule cells Neuronal cell bodies and dendrites, cardiac and smooth muscle myocytes

Distribution

Repetitive firing, dendritic calcium transients. Pacemaking, repetitive firing

Neurotransmitter release, dendritic Ca2þ transients, hormone release

Excitation-contraction coupling Excitation-contraction coupling, hormone release, transcriptional regulation, synaptic integration Hormone release, transcriptional regulation, cardiac pacemaking, hearing, neurotransmitter release from sensory cells Neurotransmitter release from photoreceptors Neurotransmitter release, dendritic Ca2þ transients, hormone release

Cellular function

Table 1 Subtypes and physiologic functions of voltage-gated calcium channels

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Table 2 Electrophysiological properties of HVA and LVA VGCCs. Each type of VGCCs has a distinct kinetics and a conductance [48] HVA LVA L P/Q, N, R T Activation range

Positive to –30 mV

Inactivation range Inactivation

–60 to –10 mV Very slow (t > 500 ms) Rapid 25 pS

Deactivation rate Single-channel conductance Single-channel openings Relative conductance Divalent block

Continual reopening Ba2þ > Ca2þ Cd2þ > Ni2þ

Positive to –20 mV –120 to –30 mV Partial (t = 5080 ms) Slow 13 pS Long burst Ba2þ > Ca2þ Cd2þ > Ni2þ

Positive to –70 mV –100 to –60 mV Complete (t = 2050 ms) Rapid 8 pS Brief burst, inactivation Ba2þ = Ca2þ Cd2+ < Ni2+

from the fish-eating cone shell, termed o-conotoxin GVIA. Second, a very slowly inactivating Ca2þ current that was insensitive to both DHP and oconotoxin GVIA was identified in Purkinje neurons of the cerebellum [49]. This current (P-type for Purkinje) was found to be sensitive to a component of the venom (o-agatoxin IVA) from the American funnel web spider, Agelenopsis aperta [19, 81]. Another o-agatoxin IVA-sensitive current showing more rapid inactivation was subsequently identified in cerebellar granule cells. This was originally termed Q-type current [97]. However, it was found that these two currents are elicited by different splicing forms of the same gene, and now these currents are termed as P/Q-type [11, 61]. There was also a residual or R-type Ca2þ current that is resistant to DHPs and the N- and P/Q-type VGCCs toxins [97]. Originally R-type Ca2þ current was included in low-voltage-activated (LVA) channels according to its activation or inactivation voltage ranges. But, now this is included in HVA channels, due to the clear difference from LVA channels. R-type Ca2þ channel is an intermdiate voltage-activated channel and is certainly more inactivating than the other high-votage-activated channels. In comparison of the homology, LVA T-type VGCCs are clearly divergent in terms of structure from the HVA channels [92] (Fig. 2). T-type VGCCs have a tiny conductance and make a transient Ca2þ current. LVA T-type VGCCs also show distinct electrophysiological and pharmacological properties compared to those of HVA VGCCs (Table 2). Neuronal T-type Ca2þ channels (Cav3.1, 3.2, and 3.3) are concentrated on the cell body and dendrites and are important in modulating neuronal excitability. They have several unique properties, including activation at very negative potentials, and the ability to promote burst firing in neurons [57, 77, 112]. T-type VGCCs require strong hyperpolarization to recover from inactivation.

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5 Genetics and Molecular Biology The genetic analysis of VGCCs provides an approach to defining channel function that is complimentary to pharmacological, electrophysiological, and other molecular ways. For an understanding of in vivo functions of VGCCs, mice lacking various a1 subunits were generated in the last few years, and then the functions of VGCCs at the whole animal level have been well described [74, 85, 93]. To study the physiological or pathological roles of L-type Ca2þ channels in vivo, knockout mice were generated. The null homozygous Cav1.2 (a1C) mutant mice died before day 14.5 postcoitum (p.c.) due to the problem of heart development, which suggests that the Cav1.2 VGCC has a critical role in the developmental stage of heart [105]. To overcome this embryonic lethal condition, Cav1.2 mouse was generated using Cre/LoxP site-specific recombination system, which allows time and tissue-specific gene deletion [79, 83, 96]. Cav1.2 was selectively deleted in forebrain including hippocampal formation and cerebral cortex using Nex-Cre donor mice [104]. Forebrain-specific Cav1.2 mutant mice were viable, and showed normal N-methyl-D-aspartate receptor (NMDAR)-dependent long-term potentiation (LTP) in the Schaffer collateral/ CA1 pathway. However, these mice showed the impairment of NMDARindependent LTP. The Cav1.2 mutant mice showed not only reduced late phase of LTP (L-LTP) that requires protein synthesis, but also exhibited impairment in spatial learning in the water maze and labyrinth maze tasks. These results demonstrate the functional relevance of Cav1.2 channel-mediated Ca2þ influx for synaptic plasticity, transcriptional activation, and learning and memory [84]. The Cav1.3 (a1D), another L-type Ca2þ channel, null homozygote mice were generated. These mutant mice were deaf, and both inner and outer hair cell populations were degenerated in the mutant mice by postnatal day 35 and had degeneration of outer and inner hair cells [95]. Notably, however, these mutant mice did not show any effects on LTP in the hippocampus and learning/ memory [23]. Mice lacking Cav2.1 (a1A) exhibited several neurological dysfunctions. Although mutant mice were not embryonically lethal, they showed ataxia and dystonia at about 10 days after birth and could not survive over 4 weeks [33, 61]. In addition, absence seizures, characterized by 3–5 Hz cortical spike-wave discharges (SWDs), which were revealed by electroencephalogram (EEG) recording, associated with behavioral immobility appeared [109]. EEG recordings also showed persistent absence status and a dramatic reduction of gamma-band activity, indicating P/Q-type Ca2þ channels are essential in the generation of gamma-band activity and resultant cognitive function [75]. Disruption of the Ca2þ influx mediated by the gene encoding Cav2.1 altered the expression and functional properties of other Ca2þ channels in neurons. With respect to

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synaptic transmission, Cav2.1 mutant mice displayed impaired excitatory transmission in the hippocampus, but they showed enhanced reliance on Ca2þ influx through N-type or R-type Ca2þ channels in neurons [61]. The Cav2.1 mutant also displayed several alterations in presynaptic function in the neuromuscular junction [123]. The N-type Ca2þ channels are predominantly localized in the nervous system and have been considered to play a vital role in transmitter release at presynaptic terminals, along with P/Q-type Ca2þ channels. The null knockout mice that are genetically deficient in the a1B subunit, unexpectedly, had a normal life span, unlike Cav2.1 mutant mice [44, 56, 64, 100]. However, the functional analysis in vivo revealed several behavioral aberrations and alterations of their nervous system. A large number of studies suggested that the Cav2.2 subunit of N-type calcium channel plays an important role in pain perception at the spinal level, as well as at the supraspinal level. The mutant mice exhibited a significant decrease in anxiety-related behaviors [10, 100, 117]. It was found that the sensitivity to halothane was significantly increased in Cav2.2 mutant mice compared with the wild-type littermates [120], but voluntary ethanol consumption was reduced [86]. With respect to synaptic transmission, the mutant mice exhibited reduced brain-derived neurotrophic factor (BDNF)-induced synaptic potentiation and facilitation. In addition, they showed impaired learning and memory in the Morris water maze and the social transmission of food preference tasks, suggesting that an interaction between N-type VGCCs and BDNF may be involved in the presynaptic enhancement of LTP and normal hippocampus-dependent learning and memory [60]. Otherwise, the role of N-type VGCC in regulation of aggression by modulating inhibitory neurotransmission in dorsal raphe nucleus was revealed [63]. Some compensatory activities were reported in Cav2.2 mutant mice. The mRNA expression of Cav2.1 subunit was increased in cerebellum, olfactory bulb, and adrenal gland [114–116]. The physiological function of R-type Ca2þ channels was not clear, compared with other types of Ca2þ channels. However, Cav2.3 mutant mice gave an advance to understanding of R-type Ca2þ channels. Cav2.3 mutant mice showed reduced spontaneous locomotor activities and increased anxiety level [71, 101]. They also displayed a reduced response to somatic inflammatory pain and an altered response to visceral inflammatory pain, without any difference in acute pain responses [101], suggesting the involvement of R-type Ca2þ channels in controlling the excitability of neurons in the descending antinociceptive pathways (periaqueductal gray matter and/or raphe magnus neurons) [102]. Mutant mice also showed decreased anesthetic sensitivities to propofol and halothane [119]. Acute administration of cocaine normally enhances the locomotor activity in animals, but failed to produce any response in Cav2.3 mutant mice [40]. The studies of synaptic plasticity in Cav2.3 mutant mice revealed that presynaptic Ca2þ entry through R-type Ca2þ channels contributes to the

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induction of mossy fiber LTP, while they did not play a role in fast synaptic transmission [24]. They also showed deficits in long-term depression (LTD) at the parallel fiber–Purkinje cell synapse and motor learning in rotarod tests, suggesting the contribution of Cav2.3 Ca2þ channels to the regulation of activity of Purkinje cells related to motor learning [91]. Mice with a global deletion of the Cav3.1 channels showed absence of the rebound burst-mode potentials in the thalamic neurons, as no difference was observed in tonic-mode firings [66]. Cav3.1 mutant mice were resistant to the genesis of spike and wave discharges induced by baclofen or g-butyrolactone. However, the mutant mice exhibited behavioral seizures in response to systemic administration of bicuculline or 4-aminopyridine. Cav3.1 mutant mice showed hyperalgesia to visceral pain induced by acetic acid [65]. The role of T-type Ca2þ channels in brain rhythms during sleep and their contribution to behavioral states of sleep were revealed [2, 69]. With EEG recordings, Cav3.1 mutant mice showed a loss of delta (1–4 Hz) waves and a reduction of sleep spindles (7–14 Hz) in the thalamus [70]. Furthermore, conditional deletion of Cav3.1 in the rostral-midline thalamus led to frequent and prolonged arousal, which fragmented and reduced sleep in mice. However, sleep was not disturbed in the mice with deletion of Cav3.1 in cortical pyramidal neurons. These findings provide that the Cav3.1 in the thalamus, but not the cortex, is required for stabilized sleep, and this is consistent with a notion that sensory suppression in the thalamus prevents disruption of sleep [2]. Coronary arteries isolated from Cav3.2 mutant arteries showed normal contractile responses, but reduced relaxation in response to acetylcholine and nitroprusside, indicating Ca2þ influx through Cav3.2 Ca2þ channels is essential for normal relaxation of coronary arteries [17]. Cav3.2 mutant mice were used to study pain susceptibility. In acute pain tests, Cav3.2 mutant mice showed decreased pain responses to both acute (mechanical, thermal, and chemical stimuli) and tonic noxious stimuli (intraperitoneal injections of irritant agents and intradermal injections of formalin) [20]. In addition, Cav3.2 mutant mice did not show any responses for GABA-induced cell excitability and intracellular Ca2þ increase in the medium-sized dorsal root ganglia neurons [3]. These results suggest that Cav3.2 T-type Ca2þ channels may be involved in both spinal reflex and supraspinal pain modulation. There was little study of the roles of Cav3.3 T-type Ca2þ channels in the nervous system and Cav3.3 Ca2þ channel-deficient mice have not been reported up to now.

6 Regulatory Mechanisms: Receptorial, Transcriptional, and Transductional As described, VGCCs are comprised of five different protein subunits: a1, a2d, b, and g [118]. The a1 and a2 subunits were co-purified with equimolar stoichiometry [106, 118]. The a2d subunit was first discovered in rabbit skeletal muscle

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[27] and later in the rat brain [67] and human brain [129]. The a2d subunits are derived from a single gene [27], and alternative splicing mechanisms have been proposed to give rise to variants of the a2 subunit [67]. Other details of the protein subunits comprising VGCCs are summarized in Table 1. There are several means by which the levels of calcium channels are regulated by second messenger pathways, such that L-type calcium channels are upregulated by cAMP-dependent kinase [98]. The regulation of N- and P/Q-type channels are inhibited by the activation of heterotrimeric G-proteins by Gprotein-coupled receptors (GPCRs). GPCRs involved in this regulation include two adenoreceptors, m- and d-opioid receptors, GABA-B receptors, and adenosine A1 receptors. These receptors or GTP analogues slow the current activation kinetics of N- and P/Q-type channels.

7 Physiological Properties VGCCs play a major role in both normal functioning and various pathological processes. The calcium action potential and subsequent calcium currents are detected in mammalian skeletal, cardiac, and all excitable cells. For example, depolarization of smooth muscle opens L-type calcium channels and causes an influx of extracellular Ca2þ, which binds calmodulin. The activated calmodulin molecule activates myosin light-chain kinase, which phosphorylates myosin. Phosphorylated myosin form cross-bridges with actin thin filaments, and thus contraction of smooth muscle fibers is driven by the sliding filament mechanism. L-type VGCCs on the neuronal cell body provide the Ca2þ signals necessary for gene activation, cell survival, and protein expression. Calcium channels are also present at low levels in many non-excitable cells, such as immune cells, though their role has not been elucidated [12].

8 Pathophysiological Relevance in Stroke The level of intracellular Ca2þ concentration is strictly regulated by multiple complex machineries. Intracellular Ca2þ normally binds to calcium-binding proteins, such as calmodulin, or exists in storage in cellular organelles, such as the endoplasmic reticulum (ER) and mitochondria. Free Ca2þ released into the intracellular space is actively excreted by the Naþ/Ca2þ-exchange system, which is coupled to ATPase. However, cerebral ischemia causes breakdown of the regulatory mechanisms, and unrestricted increases in the concentration of calcium ions in the intracellular space cause cell death. Excessive influx of Ca2þ via glutamate receptors, and excessive release of Ca2þ from the ER are involved in various cytotoxic processes, such as proteolysis, oxidative damage, and apoptosis. Although the main inflow of calcium into neurons in ischemia is mediated by

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glutamate receptors, VGCCs also participate in Ca2þ entry. Thus, the L-type VGCC activator Bay K 8644 enhances and the blocker nimodipine inhibits cellular apoptosis and mitochondrial damage, suggesting that Ca2þ entry through L-type channels is responsible for apoptosis [13, 55]. L-type VGCCs also ensure an early rise in intracellular calcium in ischemia. L-type Ca2þ channels are upregulated in reactive astrocytes after ischemia [127]. Upregulation of L-type Ca2þ channels in reactive astrocytes may contribute to the maintenance of ionic homeostasis in injured brain regions by increasing the release of neurotrophic factors to promote neuronal survival and differentiation, and enhancing signaling in astrocytic networks in response to injury [127]. Inhibiting L-type VGCCs with dihydropyridine abolished the rise in intracellular calcium [94]. L-type Ca2þ channels are also involved in the activation of calcium/calmodulin-dependent protein kinase II (CaMKII), which induces serine phosphorylation of GluR6 and neuronal cell death, and nifedipine can exert neuroprotective effects in this way [43]. Transient increases in N-type Ca2þ channels are typically observed 2–3 days after ischemia [22]. Delayed neuronal death in the CA1 region after 5 min of ischemia is attenuated by o-conotoxin in a dose-dependent manner [130]. oConotoxin has a neuroprotective effect against ischemic injury, probably by inhibition of N-type VGCC [22]. Several synthetic o-conopeptides, such as synthetic MVIIA (SNX-111), have allowed the evaluation of the therapeutic potential of blocking N-type Ca2þ channels in a variety of pathological conditions, including cerebral ischemia [78, 103]. T-type calcium channel inhibitors also reduce ischemic cell damage [87]. Although the inhibition of L-type VGCCs with dihydropyridine abolished the rise in intracellular calcium usually observed after 2–4 min, it had no effect on a small very early increase in intracellular calcium concentration [94]. Thus, the potential role of low-voltage-gated T-type calcium channels during brain ischemia has been suggested. Because T-type calcium channels have abilities to sustain a continuous calcium influx in neurons and glia at rest states by a window current mechanism, the increase in early intracellular calcium concentration observed after the initiation of an ischemic insult could be the expression of an unbalanced equilibrium between sustained calcium influx through T-type calcium channels and dysfunctional efflux mechanisms under metabolic distress [87]. Therefore, pharmacological inhibition of the T-type calcium current by mibefradil, kurtoxin, nickel, zinc, and pimozide during the oxygen–glucose deprivation episode could provide significant protections against delayed neuronal death [87].

9 Pharmacological Modulation The pharmacology of drugs acting on calcium channels is quite distinct. The calcium channel blockers used widely in the therapy of cardiovascular diseases are derivatives of dihydropyridines that target the Cav1 (L-type) channels, and

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act at three separate, but allosterically coupled, receptor sites [37]. Dihydropyridine calcium channel blockers are often used to reduce systemic vascular resistance and arterial pressure. These include ‘‘-dipine’’ drugs, such as amlodipine, felodipine, nifedipine, nicardipine, and nimodipine. Phenylalkylamine calcium channel blockers, such as verapamil, are relatively selective for myocardium, reduce myocardial oxygen demand and reverse coronary vasospasm, and they are often used to treat angina. Phenylalkylamines are intracellular pore blockers that are thought to enter the pore from the cytoplasmic side of the channel and block it [50, 52, 113]. The receptor site of dihydropyridine is closely apposed to that of phenylalkylamine, even though they share some common amino acid residues. Diltiazem and related benzothiazepines are thought to bind to a third receptor site, even though the amino acid residues required for their binding largely overlap with those required for phenylalkylamine binding. The CaV2 subfamily (P/Q, N, and R-type VGCC) of calcium channels is relatively insensitive to dihydropyridine calcium channel blockers. Several newly developed anti-epileptic drugs such as lamotrigine, levetiracetam, and pregabalin have some affinities to these receptors, which will be discussed below. The CaV3 subfamily (T-type) of calcium channels is insensitive to dihydropyridines and toxins, and there are no widely useful pharmacological agents that block T-type calcium currents [92]. Mibefradil is somewhat selective for T-type over L-type calcium currents (threefold to fivefold). Development of more specific and high-affinity blockers of the CaV3 family would be useful for both therapy and research.

10 Preliminary Clinical Trials Many clinical trials with Ca2þ antagonists acting on VGCCs have been conducted for neuroprotection. However, a critical and extensive review has shown that Ca2þ antagonists do not reduce the risk of death or dependency after acute ischemic stroke [54]. The L-type Ca2þ channels blocker nimodipine is one such example. Initially, a randomized, placebo-controlled trial in stroke patients showed a significant reduction in death and neurological impairment after administration of the calcium antagonist nimodipine [36]. Similar results were found in subarachnoid hemorrhage, in which administration of nimodipine before the onset of ischemia was associated with a reduced occurrence of cerebral ischemia and improved clinical outcome [31]. However, further randomized studies with nimodipine or other calcium antagonists (e.g., flunarizine, isradipine) in ischemic stroke did not confirm the beneficial effect of the earlier study [124]. However, subgroup analyses by the American Nimodipine Study Group of patients treated with nimodipine within 18 h after stroke onset in the American Nimodipine Trial suggested an improved outcome after treatment [1]. The

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notion that a short time interval between stroke onset and the start of treatment is crucial was supported by results of a meta-analysis of nine trials of nimodipine in patients with ischemic stroke [82]. Although the overall analysis did not show any beneficial effect, there was a statistically significant reduction in poor outcome in patients treated with nimodipine within 12 h after stroke onset (odds ratio, 0.62; 95% CI, 0.44–0.87) [82]. Though this raised hope that the risk of death and neurological impairment after ischemic stroke could be reduced with proper treatment, a systematic review of the effects of calcium antagonists in stroke could not confirm this positive effect of early treatment [54]. Another study, the Very Early Nimodipine Use in Stroke (VENUS) trial was designed to test the hypothesis that early treatment with nimodipine has a positive effect on survival and functional outcome after stroke [53]. In this randomized, double-blind, placebo-controlled trial, treatment was started by general practitioners or neurologists within 6 h after stroke onset (oral nimodipine 30 mg QID or identical placebo, for 10 days). However, the results of the VENUS study did not support the beneficial effect of early nimodipine treatment in stroke patients [53]. Though there have been other clinical trials involving the use of other Ca2þ channel blockers, such as isradipine [5], flunarizine [34, 72], and nicardipine [73], the overall systemic review and the individual results rule out a clinically important effect of administration of a calcium channel antagonist after ischemic stroke [55]. Therefore, the use of L-type VGCC blockers in acute stroke was not accepted. Because all of these effort-consuming trials were based on insufficient preclinical designs and evidences, these results raised a lot of discussion concerning the focus of future neuroprotective preclinical experiments. However, most of the clinical trials have used L-type Ca2þ channel antagonists, such as nimodipine. On the other hand, the clinical benefits of antagonists that selectively modulate N- or P/Q-type channels have not been systemically explored.

11 Therapeutic Perspectives As discussed above, the clinical benefits of antagonists that selectively modulate Nand P/Q-type VGCCs should be investigated in future studies. Interestingly, various anti-epileptic drugs have potent inhibitory effects on N- or P/Q-type channels. Lamotrigine, which has been known to inhibit Naþ-dependent repetitive firing discharge, exerts a potent inhibitory effect on N-type VGCCs [126]. Neuroprotective effects of lamotrigine have been reported in a rat cerebral ischemia model [108, 128], although some controversy remains concerning the efficacy of lamotrigine [122]. The severity of the ischemic insult as well as the differential time schedule should be taken into account when conducting a clinical trial. Another feasible anti-epileptic drug in neuroprotection is levetiracetam, which modulates N-type Ca2þ channels [76]. The neuroprotective effect of levetiracetam is comparable to that of the NMDA antagonist MK-801, with less adverse effects

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[42]. Gabapentin also modulates P/Q-type and L-type channels in addition to its inhibitory action on Naþ-dependent firing [111]; however, its neuroprotective effect in cerebral ischemia has not been elucidated. The drugs gabapentin and pregabalin interact with the P/Q-type VGCCs and have alleviated ataxia in cerebellar atrophy and other degenerative ataxias [35]. Pregabalin has exhibited more delayed channel inactivation and greater antiepileptic power than gabapentin under experimental conditions [7, 38]. Applications of gabapentin and pregabalin on VGCCs together with a possible anti-glutamatergic action [7] suggest that they may contribute to neuroprotection against ischemia. Thus, further efforts to develop therapeutics for cerebral ischemia targeting Nor P/Q-type VGCCs are still warranted. In addition, studies should develop combination therapies of VGCC blockers with other neuroprotective agents that can be used in the early treatment of ischemic stroke, such as nitrite [62], and that orchestrate other Ca2þ-permeable ion pores involved in cerebral ischemia, such as glutamate receptors, glutamate transporters [21], and Naþ/Ca2þ exchangers [58].

12 Conclusions In this review, we discussed the characteristics of VGCCs and the therapeutic feasibility of VGCC antagonists in ischemic stroke. Although L-type Ca2þ channel blockers (i.e., nimodipine) have failed to attenuate neurologic deterioration in large clinical trials, a number of newly developed Ca2þ channel blockers that target non-L-type channels remain uninvestigated. Because ‘‘calcium’’ is the key regulator in ischemic injury and neuronal death, continuing efforts to discover clinically effective therapeutics based on VGCCs and/or calcium modulators are strongly warranted. Scientists and clinicians around the world are waiting for such data to be reported soon.

References 1. The American Nimodipine Study Group. Clinical trial of nimodipine in acute ischemic stroke. Stroke 23: 3–8, 1992. 2. Anderson MP, Mochizuki T, Xie J, Fischler W, Manger JP, Talley EM, Scammell TE, and Tonegawa S. Thalamic Cav3.1 T-type Ca2+ channel plays a crucial role in stabilizing sleep. Proc Natl Acad Sci USA 102: 1743–1748, 2005. 3. Aptel H, Hilaire C, Pieraut S, Boukhaddaoui H, Mallie S, Valmier J, and Scamps F. The Cav3.2/alpha1H T-type Ca2+ current is a molecular determinant of excitatory effects of GABA in adult sensory neurons. Mol Cell Neurosci 36: 293–303, 2007. 4. Arikkath J and Campbell KP. Auxiliary subunits: essential components of the voltagegated calcium channel complex. Curr Opin Neurobiol 13: 298–307, 2003. 5. Azcona A and Lataste X. Isradipine in patients with acute ischaemic cerebral infarction. An overview of the ASCLEPIOS Programme. Drugs 40: 52–57, 1990. 6. Beguin P, Nagashima K, Gonoi T, Shibasaki T, Takahashi K, Kashima Y, Ozaki N, Geering K, Iwanaga T, and Seino S. Regulation of Ca2+ channel expression at the cell surface by the small G-protein kir/Gem. Nature 411: 701–706, 2001.

Subtypes of Voltage-Gated Ca2þ Channels and Ischemic Brain Injury

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7. Belliotti TR, Capiris T, Ekhato IV, Kinsora JJ, Field MJ, Heffner TG, Meltzer LT, Schwarz JB, Taylor CP, Thorpe AJ, Vartanian MG, Wise LD, Zhi-Su T, Weber ML, and Wustrow DJ. Structure-activity relationships of pregabalin and analogues that target the alpha(2)-delta protein. J Med Chem 48: 2294–2307, 2005. 8. Berggren PO, Yang SN, Murakami M, Efanov AM, Uhles S, Kohler M, Moede T, Fernstrom A, Appelskog IB, Aspinwall CA, Zaitsev SV, Larsson O, de Vargas LM, Fecher-Trost C, Weissgerber P, Ludwig A, Leibiger B, Juntti-Berggren L, Barker CJ, Gromada J, Freichel M, Leibiger IB, and Flockerzi V.. Removal of Ca2+ channel beta3 subunit enhances Ca2+ oscillation frequency and insulin exocytosis. Cell 119: 273–284, 2004. 9. Berrow NS, Campbell V, Fitzgerald EM, Brickley K, and Dolphin AC. Antisense depletion of beta-subunits modulates the biophysical and pharmacological properties of neuronal calcium channels. J Physiol 482: 481–491, 1995. 10. Beuckmann CT, Sinton CM, Miyamoto N, Ino M, and Yanagisawa M. N-type calcium channel alpha1B subunit (Cav2.2) knock-out mice display hyperactivity and vigilance state differences. J Neurosci 23: 6793–6797, 2003. 11. Bourinet E, Soong TW, Sutton K, Slaymaker S, Mathews E, Monteil A, Zamponi GW, Nargeot J, and Snutch TP. Splicing of alpha 1A subunit gene generates phenotypic variants of P- and Q-type calcium channels. Nat Neurosci 2: 407–415, 1999. 12. Cahalan MD, Wulff H, and Chandy KG. Molecular properties and physiological roles of ion channels in the immune system. J Clin Immunol 21: 235–252, 2001. 13. Cano-Abad MF, Villarroya M, Garcia AG, Gabilan NH, and Lopez MG. Calcium entry through L-type calcium channels causes mitochondrial disruption and chromaffin cell death. J Biol Chem 276: 39695–39704, 2001. 14. Carbone E and Lux HD. A low voltage-activated, fully inactivating Ca channel in vertebrate sensory neurones. Nature 310: 501–502, 1984. 15. Catterall WA. Structure and regulation of voltage-gated Ca2+ channels. Annu Rev Cell Dev Biol 16: 521–555, 2000. 16. Catterall WA, Perez-Reyes E, Snutch TP, and Striessnig J. International Union of Pharmacology. XLVIII. Nomenclature and structure-function relationships of voltagegated calcium channels. Pharmacol Rev 57: 411–425, 2005. 17. Chen CC, Lamping KG, Nuno DW, Barresi R, Prouty SJ, Lavoie JL, Cribbs LL, England SK, Sigmund CD, Weiss RM, Williamson RA, Hill JA, and Campbell KP. Abnormal coronary function in mice deficient in alpha1H T-type Ca2+ channels. Science 302: 1416–1418, 2003. 18. Chen YH, Li MH, Zhang Y, He LL, Yamada Y, Fitzmaurice A, Shen Y, Zhang H, Tong L, and Yang J. Structural basis of the alpha1-beta subunit interaction of voltage-gated Ca2+ channels. Nature 429: 675–680, 2004. 19. Cherksey BD, Sugimori M, and Llinas RR. Properties of calcium channels isolated with spider toxin, FTX. Ann N Y Acad Sci 635: 80–89, 1991. 20. Choi S, Na HS, Kim J, Lee J, Lee S, Kim D, Park J, Chen CC, Campbell KP, and Shin HS. Attenuated pain responses in mice lacking Ca(V)3.2 T-type channels. Genes Brain Behav 6: 425–431, 2007. 21. Chu K, Lee ST, Sinn DI, Ko SY, Kim EH, Kim JM, Kim SJ, Park DK, Jung KH, Song EC, Lee SK, Kim M, and Roh JK. Pharmacological Induction of Ischemic Tolerance by Glutamate Transporter-1 (EAAT2) Upregulation. Stroke 38: 177–182, 2007. 22. Chung YH, Shin CM, Kim MJ, Shin DH, Yoo YB, and Cha CI. Spatial and temporal distribution of N-type Ca(2+) channels in gerbil global cerebral ischemia. Brain Res 902: 294–300, 2001. 23. Clark NC, Nagano N, Kuenzi FM, Jarolimek W, Huber I, Walter D, Wietzorrek G, Boyce S, Kullmann DM, Striessnig J, and Seabrook GR. Neurological phenotype and synaptic function in mice lacking the CaV1.3 alpha subunit of neuronal L-type voltagedependent Ca2+ channels. Neuroscience 120: 435–442, 2003.

204

S.-T. Lee et al.

24. Dietrich D, Kirschstein T, Kukley M, Pereverzev A, von der Brelie C, Schneider T, and Beck H. Functional specialization of presynaptic Cav2.3 Ca2+ channels. Neuron 39: 483–496, 2003. 25. Dolphin AC. Beta subunits of voltage-gated calcium channels. J Bioenerg Biomembr 35: 599–620, 2003. 26. Dolphin AC. A short history of voltage-gated calcium channels. Br J Pharmacol 147: S56–S62, 2006. 27. Ellis SB, Williams ME, Ways NR, Brenner R, Sharp AH, Leung AT, Campbell KP, McKenna E, Koch WJ, Hui A, et al. Sequence and expression of mRNAs encoding the alpha 1 and alpha 2 subunits of a DHP-sensitive calcium channel. Science 241: 1661–1664, 1988. 28. Ertel EA, Campbell KP, Harpold MM, Hofmann F, Mori Y, Perez-Reyes E, Schwartz A, Snutch TP, Tanabe T, Birnbaumer L, Tsien RW, and Catterall WA. Nomenclature of voltage-gated calcium channels. Neuron 25: 533–535, 2000. 29. Fatt P and Katz B. The electrical properties of crustacean muscle fibres. J Physiol 120: 171–204, 1953. 30. Fedulova SA, Kostyuk PG, and Veselovsky NS. Two types of calcium channels in the somatic membrane of new-born rat dorsal root ganglion neurones. J Physiol 359: 431–446, 1985. 31. Feigin VL, Rinkel GJ, Algra A, Vermeulen M, and van Gijn J. Calcium antagonists in patients with aneurysmal subarachnoid hemorrhage: a systematic review. Neurology 50: 876–883, 1998. 32. Finlin BS, Crump SM, Satin J, and Andres DA. Regulation of voltage-gated calcium channel activity by the Rem and Rad GTPases. Proc Natl Acad Sci USA 100: 14469–14474, 2003. 33. Fletcher CF, Tottene A, Lennon VA, Wilson SM, Dubel SJ, Paylor R, Hosford DA, Tessarollo L, McEnery MW, and Pietrobon D. Copeland NG, Jenkins NA. Dystonia and cerebellar atrophy in Cacna1a null mice lacking P/Q calcium channel activity. FASEB J 15: 1288–1290, 2001. 34. Franke CL, Palm R, Dalby M, Schoonderwaldt HC, Hantson L, Eriksson B, LangJenssen L, and Smakman J. Flunarizine in stroke treatment (FIST): a double-blind, placebo-controlled trial in Scandinavia and the Netherlands. Acta Neurol Scand 93: 56–60, 1996. 35. Gazulla J and Tintore M. The P/Q-type voltage-dependent calcium channel: a therapeutic target in spinocerebellar ataxia type 6. Acta Neurol Scand 115: 356–363, 2007. 36. Gelmers HJ and Hennerici M. Effect of nimodipine on acute ischemic stroke. Pooled results from five randomized trials. Stroke 21: IV81–IV84, 1990. 37. Glossmann H and Striessnig J. Molecular properties of calcium channels. Rev Physiol Biochem Pharmacol 114: 1–105, 1990. 38. Gong HC, Hang J, Kohler W, Li L, and Su TZ. Tissue-specific expression and gabapentin-binding properties of calcium channel alpha2delta subunit subtypes. J Membr Biol 184: 35–43, 2001. 39. Gonzalez-Gutierrez G, Miranda-Laferte E, Neely A, and Hidalgo P. The Src homology 3 domain of the beta-subunit of voltage-gated calcium channels promotes endocytosis via dynamin interaction. J Biol Chem 282: 2156–2162, 2007. 40. Han W, Saegusa H, Zong S, and Tanabe T. Altered cocaine effects in mice lacking Ca(v)2. 3 (alpha(1E)) calcium channel. Biochem Biophys Res Commun 299: 299–304, 2002. 41. Hanlon MR, Berrow NS, Dolphin AC, and Wallace BA. Modelling of a voltage-dependent Ca2+ channel beta subunit as a basis for understanding its functional properties. FEBS Lett 445: 366–370, 1999. 42. Hanon E and Klitgaard H. Neuroprotective properties of the novel antiepileptic drug levetiracetam in the rat middle cerebral artery occlusion model of focal cerebral ischemia. Seizure 10: 287–293, 2001.

Subtypes of Voltage-Gated Ca2þ Channels and Ischemic Brain Injury

205

43. Hao ZB, Pei DS, Guan QH, and Zhang GY. Calcium/calmodulin-dependent protein kinase II (CaMKII), through NMDA receptors and L-Voltage-gated channels, modulates the serine phosphorylation of GluR6 during cerebral ischemia and early reperfusion period in rat hippocampus. Brain Res Mol Brain Res 140: 55–62, 2005. 44. Hatakeyama S, Wakamori M, Ino M, Miyamoto N, Takahashi E, Yoshinaga T, Sawada K, Imoto K, Tanaka I, Yoshizawa T, Nishizawa Y, Mori Y, Niidome T, and Shoji S. Differential nociceptive responses in mice lacking the alpha(1B) subunit of N-type Ca(2+) channels. NeuroReport 12: 2423–2427, 2001. 45. Hess P, Lansman JB, and Tsien RW. Different modes of Ca channel gating behaviour favoured by dihydropyridine Ca agonists and antagonists. Nature 311: 538–544, 1984. 46. Hibino H, Pironkova R, Onwumere O, Rousset M, Charnet P, Hudspeth AJ, and Lesage F. Direct interaction with a nuclear protein and regulation of gene silencing by a variant of the Ca2+-channel beta 4 subunit. Proc Natl Acad Sci USA 100: 307–312, 2003. 47. Hidalgo P and Neely A. Multiplicity of protein interactions and functions of the voltagegated calcium channel beta-subunit. Cell Calcium 42: 389–396, 2007. 48. Hille B.. Ion Channels of Excitable Membranes, 3rd ed. Sunderland, MA: Sinauer Associates, Inc., 2001. 49. Hillman D, Chen S, Aung TT, Cherksey B, Sugimori M, and Llinas RR. Localization of P-type calcium channels in the central nervous system. Proc Natl Acad Sci USA 88: 7076–7080, 1991. 50. Hockerman GH, Peterson BZ, Johnson BD, and Catterall WA. Molecular determinants of drug binding and action on L-type calcium channels. Annu Rev Pharmacol Toxicol 37: 361–396, 1997. 51. Hofmann F, Biel M, and Flockerzi V. Molecular basis for Ca2+ channel diversity. Annu Rev Neurosci 17: 399–418, 1994. 52. Hofmann F, Lacinova L, and Klugbauer N. Voltage-dependent calcium channels: from structure to function. Rev Physiol Biochem Pharmacol 139: 33–87, 1999. 53. Horn J, de Haan RJ, Vermeulen M, and Limburg M. Very Early Nimodipine Use in Stroke (VENUS): a randomized, double-blind, placebo-controlled trial. Stroke 32: 461–465, 2001. 54. Horn J and Limburg M. Calcium antagonists for acute ischemic stroke. Cochrane Database Syst Rev 2: CD001928, 2000. 55. Horn J and Limburg M. Calcium antagonists for ischemic stroke: a systematic review. Stroke 32: 570–576, 2001. 56. Ino M, Yoshinaga T, Wakamori M, Miyamoto N, Takahashi E, Sonoda J, Kagaya T, Oki T, Nagasu T, Nishizawa Y, Tanaka I, Imoto K, Aizawa S, Koch S, Schwartz A, Niidome T, Sawada K, and Mori Y. Functional disorders of the sympathetic nervous system in mice lacking the alpha 1B subunit (Cav 2.2) of N-type calcium channels. Proc Natl Acad Sci USA 98: 5323–5328, 2001. 57. Jahnsen H and Llinas R. Voltage-dependent burst-to-tonic switching of thalamic cell activity: an in vitro study. Arch Ital Biol 122: 73–82, 1984. 58. Jeon D, Chu K, Jung KH, Kim M, Yoon BW, Lee CJ, Oh U, and Shin HS. Na(+)/ Ca(2+) exchanger 2 is neuroprotective by exporting Ca(2+) during a transient focal cerebral ischemia in the mouse. Cell Calcium 43: 482–491, 2008. 59. Jeon D, Song I, Guido W, Kim K, Kim E, Oh U, and Shin HS. Ablation of Ca2+ channel beta3 subunit leads to enhanced N-methyl-D-aspartate receptor-dependent long term potentiation and improved long term memory. J Biol Chem 283: 12093–12101, 2008. 60. Jeon D, Yang YM, Jeong MJ, Philipson KD, Rhim H, and Shin HS. Enhanced learning and memory in mice lacking Na+/Ca2+ exchanger 2. Neuron 38: 965–976, 2003. 61. Jun K, Piedras-Renteria ES, Smith SM, Wheeler DB, Lee SB, Lee TG, Chin H, Adams ME, Scheller RH, Tsien RW, and Shin HS. Ablation of P/Q-type Ca(2+) channel currents, altered synaptic transmission, and progressive ataxia in mice lacking the alpha(1A)-subunit. Proc Natl Acad Sci USA 96: 15245–15250, 1999.

206

S.-T. Lee et al.

62. Jung KH, Chu K, Ko SY, Lee ST, Sinn DI, Park DK, Kim JM, Song EC, Kim M, and Roh JK. Early intravenous infusion of sodium nitrite protects brain against in vivo ischemia-reperfusion injury. Stroke 37: 2744–2750, 2006. 63. Kim C, Jeon D, Kim YH, Lee CJ, Kim H, and Shin HS. Deletion of N-type Ca2+ channel Cav2.2 results in hyperaggressive behaviors in mice. J Biol Chem, 2008, Nov 12. [Epub ahead of print]. 64. Kim C, Jun K, Lee T, Kim SS, McEnery MW, Chin H, Kim HL, Park JM, Kim DK, Jung SJ, Kim J, and Shin HS. Altered nociceptive response in mice deficient in the alpha(1B) subunit of the voltage-dependent calcium channel. Mol Cell Neurosci 18: 235–245, 2001. 65. Kim D, Park D, Choi S, Lee S, Sun M, Kim C, and Shin HS. Thalamic control of visceral nociception mediated by T-type Ca2+ channels. Science 302: 117–119, 2003. 66. Kim D, Song I, Keum S, Lee T, Jeong MJ, Kim SS, McEnery MW, and Shin HS. Lack of the burst firing of thalamocortical relay neurons and resistance to absence seizures in mice lacking alpha(1G) T-type Ca(2+) channels. Neuron 31: 35–45, 2001. 67. Kim HL, Kim H, Lee P, King RG, and Chin H. Rat brain expresses an alternatively spliced form of the dihydropyridine-sensitive L-type calcium channel alpha 2 subunit. Proc Natl Acad Sci USA 89: 3251–3255, 1992. 68. Kogan JH, Frankland PW, Blendy JA, Coblentz J, Marowitz Z, Schutz G, and Silva AJ. Spaced training induces normal long-term memory in CREB mutant mice. Curr Biol 7: 1–11, 1997. 69. Lee J, Kim D, and Shin HS. Lack of delta waves and sleep disturbances during non-rapid eye movement sleep in mice lacking alpha1G-subunit of T-type calcium channels. Proc Natl Acad Sci USA 101: 18195–18199, 2004. 70. Lee JL, Everitt BJ, and Thomas KL. Independent cellular processes for hippocampal memory consolidation and reconsolidation. Science 304: 839–843, 2004. 71. Lee SC, Choi S, Lee T, Kim HL, Chin H, and Shin HS. Molecular basis of R-type calcium channels in central amygdala neurons of the mouse. Proc Natl Acad Sci USA 99: 3276–3281, 2002. 72. Limburg M and Hijdra A. Flunarizine in acute ischemic stroke: a pilot study. Eur Neurol 30: 121–122, 1990. 73. Lisk DR, Grotta JC, Lamki LM, Tran HD, Taylor JW, Molony DA, and Barron BJ. Should hypertension be treated after acute stroke? A randomized controlled trial using single photon emission computed tomography. Arch Neurol 50: 855–862, 1993. 74. Liu L, Zwingman TA, and Fletcher CF. In vivo analysis of voltage-dependent calcium channels. J Bioenerg Biomembr 35: 671–685, 2003. 75. Llinas RR, Choi S, Urbano FJ, and Shin HS. Gamma-band deficiency and abnormal thalamocortical activity in P/Q-type channel mutant mice. Proc Natl Acad Sci USA 104: 17819–17824, 2007. 76. Lukyanetz EA, Shkryl VM, and Kostyuk PG. Selective blockade of N-type calcium channels by levetiracetam. Epilepsia 43: 9–18, 2002. 77. McCormick DA and Bal T. Sleep and arousal: thalamocortical mechanisms. Annu Rev Neurosci 20: 185–215, 1997. 78. Menzler S, Bikker JA, Suman-Chauhan N, and Horwell DC. Design and biological evaluation of non-peptide analogues of omega-conotoxin MVIIA. Bioorg Med Chem Lett 10: 345–347, 2000. 79. Metzger D and Feil R. Engineering the mouse genome by site-specific recombination. Curr Opin Biotechnol 10: 470–476, 1999. 80. Mikami A, Imoto K, Tanabe T, Niidome T, Mori Y, Takeshima H, Narumiya S, and Numa S. Primary structure and functional expression of the cardiac dihydropyridinesensitive calcium channel. Nature 340: 230–233, 1989. 81. Mintz IM, Venema VJ, Swiderek KM, Lee TD, Bean BP, and Adams ME. P-type calcium channels blocked by the spider toxin omega-Aga-IVA. Nature 355: 827–829, 1992.

Subtypes of Voltage-Gated Ca2þ Channels and Ischemic Brain Injury

207

82. Mohr JP, Orgogozo JM, Harrison MJG, Hennerici M, Wahlgren NG, Gelmers HJ, Martinez-Vila E, Dycka J, and Tettenborn D. Meta-analysis of oral nimodipine trials in acute ischemic stoke. Cerebrovasc Dis 4: 197–203, 1994. 83. Moosmang S, Haider N, Klugbauer N, Adelsberger H, Langwieser N, Muller J, Stiess M, Marais E, Schulla V, Lacinova L, Goebbels S, Nave KA, Storm DR, Hofmann F, and Kleppisch T. Role of hippocampal Cav1.2 Ca2+ channels in NMDA receptorindependent synaptic plasticity and spatial memory. J Neurosci 25: 9883–9892, 2005. 84. Moosmang S, Kleppisch T, Wegener J, Welling A, and Hofmann F. Analysis of calcium channels by conditional mutagenesis. Handb Exp Pharmacol 178: 469–490, 2007. 85. Muth JN, Varadi G, and Schwartz A. Use of transgenic mice to study voltage-dependent Ca2+ channels. Trends Pharmacol Sci 22: 526–532, 2001. 86. Newton PM, Orr CJ, Wallace MJ, Kim C, Shin HS, and Messing RO. Deletion of Ntype calcium channels alters ethanol reward and reduces ethanol consumption in mice. J Neurosci 24: 9862–9869, 2004. 87. Nikonenko I, Bancila M, Bloc A, Muller D, and Bijlenga P. Inhibition of T-type calcium channels protects neurons from delayed ischemia-induced damage. Mol Pharmacol 68: 84–89, 2005. 88. Nowycky MC, Fox AP, and Tsien RW. Three types of neuronal calcium channel with different calcium agonist sensitivity. Nature 316: 440–443, 1985. 89. Olivera BM, Imperial JS, Cruz LJ, Bindokas VP, Venema VJ, and Adams ME. Calcium channel-targeted polypeptide toxins. Ann N Y Acad Sci 635: 114–122, 1991. 90. Opatowsky Y, Chen CC, Campbell KP, and Hirsch JA. Structural analysis of the voltage-dependent calcium channel beta subunit functional core and its complex with the alpha 1 interaction domain. Neuron 42: 387–399, 2004. 91. Osanai M, Saegusa H, Kazuno AA, Nagayama S, Hu Q, Zong S, Murakoshi T, and Tanabe T. Altered cerebellar function in mice lacking CaV2.3 Ca2+ channel. Biochem Biophys Res Commun 344: 920–925, 2006. 92. Perez-Reyes E. Molecular physiology of low-voltage-activated t-type calcium channels. Physiol Rev 83: 117–161, 2003. 93. Pietrobon D. Function and dysfunction of synaptic calcium channels: insights from mouse models. Curr Opin Neurobiol 15: 257–265, 2005. 94. Pisani A, Calabresi P, Tozzi A, D’Angelo V, and Bernardi G. L-type Ca2+ channel blockers attenuate electrical changes and Ca2+ rise induced by oxygen/glucose deprivation in cortical neurons. Stroke 29: 196–201, 1998, discussion 202. 95. Platzer J, Engel J, Schrott-Fischer A, Stephan K, Bova S, Chen H, Zheng H, and Striessnig J. Congenital deafness and sinoatrial node dysfunction in mice lacking class D L-type Ca2+ channels. Cell 102: 89–97, 2000. 96. Rajewsky K, Gu H, Kuhn R, Betz UA, Muller W, Roes J, and Schwenk F. Conditional gene targeting. J Clin Invest 98: 600–603, 1996. 97. Randall A and Tsien RW. Pharmacological dissection of multiple types of Ca2+ channel currents in rat cerebellar granule neurons. J Neurosci 15: 2995–3012, 1995. 98. Reuter H. Calcium channel modulation by beta-adrenergic neurotransmitters in the heart. Experientia 43: 1173–1175, 1987. 99. Rousset M, Cens T, and Charnet P. Alone at last! New functions for Ca2+ channel beta subunits? Sci STKE 2005: pe11, 2005. 100. Saegusa H, Kurihara T, Zong S, Kazuno A, Matsuda Y, Nonaka T, Han W, Toriyama H, and Tanabe T. Suppression of inflammatory and neuropathic pain symptoms in mice lacking the N-type Ca2+ channel. EMBO J 20: 2349–2356, 2001. 101. Saegusa H, Kurihara T, Zong S, Minowa O, Kazuno A, Han W, Matsuda Y, Yamanaka H, Osanai M, Noda T, and Tanabe T. Altered pain responses in mice lacking alpha 1E subunit of the voltage-dependent Ca2+ channel. Proc Natl Acad Sci USA 97: 6132–6137, 2000. 102. Saegusa H, Matsuda Y, and Tanabe T. Effects of ablation of N- and R-type Ca(2+) channels on pain transmission. Neurosci Res 43: 1–7, 2002.

208

S.-T. Lee et al.

103. Samii A, Badie H, Fu K, Luther RR, and Hovda DA. Effects of an N-type calcium channel antagonist (SNX 111; Ziconotide) on calcium-45 accumulation following fluidpercussion injury. J Neurotrauma 16: 879–892, 1999. 104. Schwab MH, Bartholomae A, Heimrich B, Feldmeyer D, Druffel-Augustin S, Goebbels S, Naya FJ, Zhao S, Frotscher M, Tsai MJ, and Nave KA. Neuronal basic helix-loophelix proteins (NEX and BETA2/Neuro D) regulate terminal granule cell differentiation in the hippocampus. J Neurosci 20: 3714–3724, 2000. 105. Seisenberger C, Specht V, Welling A, Platzer J, Pfeifer A, Kuhbandner S, Striessnig J, Klugbauer N, Feil R, and Hofmann F. Functional embryonic cardiomyocytes after disruption of the L-type alpha1C (Cav1.2) calcium channel gene in the mouse. J Biol Chem 275: 39193–39199, 2000. 106. Sharp AH and Campbell KP. Affinity purification of antibodies specific for 1,4-dihydropyridine Ca2+ channel blockers. Circ Res 61: 37–45, 1987. 107. Shin HS, Kim C, Kim D, and Choi S. Functional Diversity of Votage-Dependent Ca2+ Channels in Nociception: Recent Progress in Genetic Studies. Curr Top Membr 57: 415–438, 2006. 108. Smith SE and Meldrum BS. Cerebroprotective effect of lamotrigine after focal ischemia in rats. Stroke 26: 117–121, 1995, discussion 121–112. 109. Song I, Kim D, Choi S, Sun M, Kim Y, and Shin HS. Role of the alpha1G T-type calcium channel in spontaneous absence seizures in mutant mice. J Neurosci 24: 5249–5257, 2004. 110. Stanley EF and Atrakchi AH. Calcium currents recorded from a vertebrate presynaptic nerve terminal are resistant to the dihydropyridine nifedipine. Proc Natl Acad Sci USA 87: 9683–9687, 1990. 111. Stefani A, Spadoni F, and Bernardi G. Gabapentin inhibits calcium currents in isolated rat brain neurons. Neuropharmacology 37: 83–91, 1998. 112. Steriade M and Timofeev I. Neuronal plasticity in thalamocortical networks during sleep and waking oscillations. Neuron 37: 563–576, 2003. 113. Striessnig J. Pharmacology, structure and function of cardiac L-type Ca(2+) channels. Cell Physiol Biochem 9: 242–269, 1999. 114. Takahashi E, Ino M, Miyamoto N, and Nagasu T. Expression analysis of P/Q-type Ca2+ channel alpha 1A subunit mRNA in olfactory mitral cell in N-type Ca2+ channel alpha 1B subunit gene-deficient mice. Neurosci Lett 359: 37–40, 2004. 115. Takahashi E, Ino M, Miyamoto N, and Nagasu T. Increased expression of P/Q-type Ca2+ channel alpha1A subunit mRNA in cerebellum of N-type Ca2+ channel alpha1B subunit gene-deficient mice. Brain Res Mol Brain Res 124: 79–87, 2004. 116. Takahashi E and Nagasu T. Expression pattern of voltage-dependent calcium channel alpha1 and beta subunits in adrenal gland of N-type Ca2+ channel alpha1B subunit gene-deficient mice. Mol Cell Biochem 271: 91–99, 2005. 117. Takahashi E and Nagasu T. Individual and group housing in N-type Ca2+ channel alpha1B subunit-deficient mice differently affect motor activity. J Am Assoc Lab Anim Sci 45: 44–47, 2006. 118. Takahashi M, Seagar MJ, Jones JF, Reber BF, and Catterall WA. Subunit structure of dihydropyridine-sensitive calcium channels from skeletal muscle. Proc Natl Acad Sci USA 84: 5478–5482, 1987. 119. Takei T, Saegusa H, Zong S, Murakoshi T, Makita K, and Tanabe T. Anesthetic sensitivities to propofol and halothane in mice lacking the R-type (Cav2.3) Ca2+ channel. Anesth Analg 97: 96–103, 2003, table of contents. 120. Takei T, Saegusa H, Zong S, Murakoshi T, Makita K, and Tanabe T. Increased sensitivity to halothane but decreased sensitivity to propofol in mice lacking the Ntype Ca2+ channel. Neurosci Lett 350: 41–45, 2003. 121. Tanabe T, Takeshima H, Mikami A, Flockerzi V, Takahashi H, Kangawa K, Kojima M, Matsuo H, Hirose T, and Numa S. Primary structure of the receptor for calcium channel blockers from skeletal muscle. Nature 328: 313–318, 1987.

Subtypes of Voltage-Gated Ca2þ Channels and Ischemic Brain Injury

209

122. Traystman RJ, Klaus JA, DeVries AC, Shaivitz AB, and Hurn PD. Anticonvulsant lamotrigine administered on reperfusion fails to improve experimental stroke outcomes. Stroke 32: 783–787, 2001. 123. Trust Study Group. Randomised, double-blind, placebo-controlled trial of nimodipine in acute stroke. Lancet 336: 1205–1209, 1990. 124. Urbano FJ, Piedras-Renteria ES, Jun K, Shin HS, Uchitel OD, and Tsien RW. Altered properties of quantal neurotransmitter release at endplates of mice lacking P/Q-type Ca2+ channels. Proc Natl Acad Sci USA 100: 3491–3496, 2003. 125. Van Petegem F, Clark KA, Chatelain FC, and Minor DL Jr. Structure of a complex between a voltage-gated calcium channel beta-subunit and an alpha-subunit domain. Nature 429: 671–675, 2004. 126. Wang SJ, Tsai JJ, and Gean PW. Lamotrigine inhibits depolarization-evoked Ca++ influx in dissociated amygdala neurons. Synapse 29: 355–362, 1998. 127. Westenbroek RE, Bausch SB, Lin RC, Franck JE, Noebels JL, and Catterall WA. Upregulation of L-type Ca2+ channels in reactive astrocytes after brain injury, hypomyelination, and ischemia. J Neurosci 18: 2321–2334, 1998. 128. Wiard RP, Dickerson MC, Beek O, Norton R, and Cooper BR. Neuroprotective properties of the novel antiepileptic lamotrigine in a gerbil model of global cerebral ischemia. Stroke 26: 466–472, 1995. 129. Williams ME, Brust PF, Feldman DH, Patthi S, Simerson S, Maroufi A, McCue AF, Velicelebi G, Ellis SB, and Harpold MM. Structure and functional expression of an omega-conotoxin-sensitive human N-type calcium channel. Science 257: 389–395, 1992. 130. Yamada K, Teraoka T, Morita S, Hasegawa T, and Nabeshima T. Omega-conotoxin GVIA protects against ischemia-induced neuronal death in the Mongolian gerbil but not against quinolinic acid-induced neurotoxicity in the rat. Neuropharmacology 33: 251–254, 1994. 131. Yang J and Tsien RW. Enhancement of N- and L-type calcium channel currents by protein kinase C in frog sympathetic neurons. Neuron 10: 127–136, 1993.

The Diverse Roles of Kþ Channels in Brain Ischemia Hiroaki Misonou and James S. Trimmer

1 Introduction Brain cells express a wide variety of Kþ-selective ion channels. Among these are inward rectifying Kþ (Kir) channels, Ca2þ-activated Kþ channels, and voltagegated Kþ (Kv) channels. These Kþ channels are crucial regulators of membrane excitability due to their ability to stabilize the membrane potential. During ischemic stress in the brain a variety of pathological events occur, including increased extracellular levels of glutamate, increased levels of intracellular Ca2þ and Naþ, and a rapid reduction in intracellular ATP levels [1]. These events and the associated aberrant neuronal membrane excitability ultimately lead to neuronal damage and death. We have learned in the previous chapters that numerous molecular events are implicated in the deleterious effects of brain ischemia. However, brain neurons also contain numerous endogenous protective mechanisms [1]. Kþ channels are ideally suited for this job. KATP/Kir6.2 channels open in response to reduced cellular ATP levels or [ATP]i, Slo1 channels exhibit enhanced activation in response to increased intracellular Ca2þ levels or [Ca2þ]i, and the activity of Kv2.1 channels can be potentiated by either of these changes. The opening of these Kþ channels hyperpolarizes the membrane potential toward the Kþ equilibrium potential (EK) and leads to a suppression of neuronal excitability. Recent studies have revealed the importance of protective mechanisms provided by endogenous Kþ channels in the response of neurons to brain ischemia.

2 Gene Structures of Kir6.2, Slo1, and Kv2.1 The human Kir6.2 gene (KCNJ11) is located on chromosome 11 (11p15.1) and is quite simple in that the entire coding region (1,173 bp) is contained within a single exon. The human Slo1 gene (KCNMA1) is located on chromosome 10 H. Misonou (*) Department of Neural and Pain Sciences, Program in Neuroscience, Dental School, University of Maryland, Baltimore, MD 21201, USA e-mail: [email protected]

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(10q22.3) and is quite complex in that the coding sequence is spread across 770 Kbp in multiple (>20) exons. Slo1 transcripts are subjected to extensive processing, and mammalian neurons express alternatively spliced variants [2; 3]. The human Kv2.1 gene (KCNB1) is located on chromosome 20 (20q13.11), and the 2,577 bp coding region is contained in two exons within the 109 Kbp gene.

3 Protein Structures of Kir6.2, Slo1, and Kv2.1 Kþ channels are tetramers of pore-forming a subunits. The 390 amino acid Kir6.2 a subunit polypeptide contains two transmembrane segments and cytoplasmic N- and C-termini. Although a tetramer of Kir6.2 subunits confers the core of the channel and the primary ATP-binding site, the sulfonylurea receptor SUR1 subunit is a component of all brain Kir6.2-containing channels, which exist as [Kir6.2]4[SUR1]4 octameric KATP channel complexes [4]. The-858amino acid Kv2.1 a subunit contains a cytoplasmic N-terminus, six transmembrane segments, and a cytoplasmic C-terminus. The tetrameric assembly of the transmembrane segments including the pore-forming loop creates the core of the Kv channel [5]. The basic configuration of Slo1 channels (1,200 residues) is similar to that of Kv2.1 except that Slo1 has an additional transmembrane segment (S0) before the core region such that the N-terminus of Slo1 is exposed to the extracellular space [6]. Slo1 has an extremely long cytoplasmic C-terminus that contains components of the Ca2þ-sensing mechanism [7].

4 Molecular Biology of Kir6.2, Slo1, and Kv2.1 Kir6.2 was cloned from a human genomic library [8], and shares 96% amino acid identity with mouse and rat Kir6.2. Kir6.2 is mutated in patients with hyperinsulinemic hypoglycemia and permanent neonatal diabetes. Slo1 was originally isolated from a mouse brain cDNA library screened with a Drosophila Slowpoke cDNA probe [9]. Human, mouse, and rat Slo1 polypeptides are highly similar. A point mutation in human Slo1 leads to generalized epilepsy and paroxysmal dyskinesia. Kv2.1 was originally cloned from a rat brain cDNA library by expression cloning in Xenopus oocytes [10]. Mouse (97% amino acid identity) and human (94% amino acid identity) are highly similar to rat Kv2.1. Mutations in Kv2.1 have not been associated with any diseases.

5 Tissue and Cellular Distribution of Kir6.2, Slo1, and Kv2.1 in Mammalian Brain Kir6.2, Slo1, and Kv2.1 are highly expressed in mammalian brain, although they are also expressed in other tissues. Most studies of their localization have been performed in rat or mouse brain. Kir6.2 is expressed widely in brain, as is

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SUR1 [11; 12]. Immunohistochemical studies suggest that Kir6.2 is mainly found in the somata and dendrites of brain neurons [13; 14]. Slo1 is also widely expressed in the brain, and is predominantly found in neuronal axons and nerve terminals [15; 16]; this is especially apparent in the hippocampus (Fig. 1) and the cerebellum (Fig. 2). Slo1 is also found in dendrites, for example in cerebellar Purkinje cells (Fig. 2) [16]. Kv2.1 is highly expressed in virtually all neurons in the central nervous system (Fig. 1) [17]. Kv2.1 is highly concentrated in the somata and the proximal portion of dendrites (Fig. 1) where it forms discrete surface clusters [18].

Fig. 1 Localization of Slo1 and Kv2.1 in adult rat hippocampus. Double immunofluorescence staining for Slo1 (red) and Kv2.1 (green). DG: dentate gyrus; PF: perforant path; MF: mossy fibers

Fig. 2 Localization of Slo1 in the axons and terminals of cerebellar basket cells and the dendrites of Purkinje cells. Rat cerebellar sections were stained by double immunofluorescence for Slo1 (red) and phosphorylated forms of neurofilament H (as an axonal marker, green). Slo1 is highly detected not only in the basket cell terminals attached to the cell bodies of Purkinje cells, but also in Purkinje cell dendrites in the molecular layer of the cerebellar cortex

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Kir6.2 [19] and Slo1 [20] channels are also present in mitochondria where they may play a distinct role in hypoxic stress. This aspect of their function will not be discussed here.

6 Biophysical Properties of Kir6.2, Slo1, and Kv2.1 Channels Most biophysical studies on Kþ channels have been performed in heterologous expression system such as Xenopus oocytes or transfected mammalian cell lines. KATP channels formed by heterologously expressed Kir6.2 and SUR1 have a relatively large Kþ conductance of 70 pS [21]. In the presence of [ATP]i > 500 mM, the open probability of Kir6.2/SUR1 channels is nearly zero but increases as [ATP]i decreases [4]. Although Kir6.2 is a member of the Kir channel family whose ionic currents are blocked at positive membrane potentials, Kir6.2 channels display only weak rectification and are able to exhibit substantial outward Kþ currents above EK [22]. This property would allow Kir6.2 channels to confer ischemia-induced outward Kþ currents in neurons. Slo1 channels are exceptional in their large unitary conductance, which ranges from 100 to 270 pS [7]. When activated by membrane depolarization coinciding with increased [Ca2þ]i, Slo1 channels show non-inactivating Kþ currents similar to the delayed rectifier current. Both voltage and Ca2þ dramatically regulate the open probability of Slo1. In the absence of Ca2þ, Slo1 is not active within the physiological range of membrane potentials. The open probability of Slo1 channels dramatically increases when [Ca2þ]i is 1–100 mM, yielding activation at physiological membrane potentials. Kv2.1 forms delayed rectifier Kþ channels with a single channel conductance of 10pS [23] and a relatively high-threshold for voltagedependent activation (G1/2 0 to +20 mV), although this is highly dependent on Kv2.1 phosphorylation state [39]. The steady-state inactivation of Kv2.1 is unique that the voltage-dependent curve is U-shaped, with maximum inactivation at 0 mV and less inactivation observed with larger depolarizations [24]. It should be noted that these characteristics are similar to those of somatic delayed rectifier currents in hippocampal pyramidal neurons [25], of which Kv2.1 is thought to be the major component [26; 27].

7 Regulation of Kir6.2, Slo1, and Kv2.1 Channels The activity of KATP channels is highly regulated by the metabolic state of the neuron, as the channel is inhibited by the binding of ATP and activated by the binding of Mg2þ-nucleotides such as Mg2þ-ADP [4]. The binding of ATP to Kir6.2 stabilizes the closed state of the channel [28]. Because the association

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constant for ATP binding to KATP channels is 10 mM, and given a normal [ATP]i 1 mM, most of the Kir6.2 subunits are normally bound to ATP. Thus, under physiological conditions the open probability of KATP channels is negligible. Reducing [ATP]i results in loss of bound ATP from Kir6.2 and association of Mg2þ-ADP with SUR1, leads to activation of Kir6.2-containing KATP channels. Kir6.2 also binds and is regulated by the important membrane phospholipid PIP2, which is mutually exclusive with the ATP binding and stabilizes Kir6.2 in the open conformation [28]. Kir6.2 is also modulated by protein kinase A [29]. Slo1 channels can be activated by either membrane depolarization or increased [Ca2þ]i alone. Under physiological conditions, Slo1 channels need to be activated synergistically by both membrane depolarization and Ca2þ, and can act as coincidence detectors of these two important neuronal signaling events. However, in pathological conditions where [Ca2þ]i can be extremely high, Slo1 may be activated solely by Ca2þ. Slo1 is also subjected to phosphorylation by multiple kinases [30]. Recent mass spectrometric analyses of Slo1 purified from mammalian brain have revealed extensive phosphorylation on its cytoplasmic C-terminus [3]. Native Slo1 channels are associated with the auxiliary b subunits, b1–4 [31]. Intriguingly, not only do the b subunits alter the biophysical properties of Slo1 channels, but they also confer sensitivity to various steroid hormones [32], including estradiol [33], which has been implicated as a protective agent in brain ischemia. Slo1 channels are sensitive to redox state, such that a reduction in oxygen in chemoreceptors in the carotid body and in heterologous expression systems leads to inhibition of Slo1 [34]. In contrast, oxidation of Slo1 polypeptides, which could occur during reperfusion following ischemia, can activate Slo1 by shifting its voltage dependence of activation by –30 mV [35]. The contribution of the redox regulation of Slo1 in ischemia is unclear as these effects are negligible in the presence of Ca2þ above 100 mM, which is readily achieved in ischemic neurons. Heme can bind directly to Slo1 [36], conferring regulation of Slo1 by carbon monoxide [37]. Kv2.1 is activated solely by voltage. However, its voltage-dependent properties are dramatically altered by changes in Kv2.1 phosphorylation state [38]. Recent mass spectrometric analyses have revealed that Kv2.1 is constitutively phosphorylated at multiple sites (>16) in brain neurons [39]. Individual mutations of the phosphorylation sites to dephosphorylation-mimetic alanine result in significant shifts in the voltage dependence of activation and steady-state inactivation toward more hyperpolarizing membrane potentials. Interestingly, concurrent mutations of multiple sites to alanine elicit an additive effect, indicating that Kv2.1 channels are subjected to a graded regulation by changes in phosphorylation at multiple sites. Similar graded responses have also been observed for the delayed rectifier current in neurons [38], where the Ca2þ/ calmodulin-dependent protein phosphatase calcineurin plays a crucial role by dephosphorylating Kv2.1 [39; 40].

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8 Physiological Properties of the KþChannels Studies of Kir6.2 knockout mice revealed that, under physiological conditions, the firing behavior of glucose-responsive neurons in the hypothalamus is regulated by the action of Kir6.2 [41]. These hypothalamic neurons increase their firing rate when the extracellular glucose level is elevated (from a few to tens of millimolar). This behavior was completely eliminated in the Kir6.2 knockout mice, indicating that the inactivation of Kir6.2 accounts for the change in the firing phenotype. However, the exact mechanism is unknown as intracellular ATP can efficiently block Kir6.2 with an IC50 10 mM. It seems unlikely that the [ATP]i in these neurons fluctuates between the micromolar and millimolar range, such that other regulatory mechanisms, such as PIP2 binding, may be involved. The sensitivity of Slo1 to Ca2þ, the channel’s rapid activation kinetics, and its extremely large single-channel conductance makes activation of Slo1 an important response to Ca2þ entry and membrane depolarization under physiological conditions. Slo1 is implicated in the generation of the fast phase of the after-hyperpolarization potential in the somata of pyramidal neurons [42], and acts as a key determinant of the refractory period of action potential firing [43]. At dendrites and nerve terminals, Slo1 channels can limit the influx of extracellular Ca2þ through voltage-gated Ca2þ channels [44; 45]. These characteristics might also be important in the pathological state of neurons during brain ischemia. Kv2.1 channels have relatively slow activation kinetics [46] and thus may not be efficiently recruited during a single action potential in the soma. However, in response to repetitive firing of action potentials and frequency-dependent action potential broadening observed in many neurons [43], Kv2.1 can be effectively activated by Ca2þ-dependent dephosphorylation and contribute to the repolarization of membrane potential. In fact, antisense knockdown of Kv2.1 in hippocampal pyramidal neurons results in aberrant hyperexcitability of the neurons and resultant Ca2þ overload at high firing frequencies [26]. As the history of activity within neurons can affect the phosphorylation state of Kv2.1 (higher activity leading to dephosphorylation and enhanced activation of Kv2.1, lower activity the opposite) [38], the role of Kv2.1 in cell excitability is conditional.

9 Pathophysiological Relevance in Stroke For most neurons under physiological conditions, Kir6.2/SUR1 channels are presumably closed due to high [ATP]i. However, they can be rapidly activated in various in vitro models of brain ischemia. Mimicking ischemia in vitro rapidly (<1 min) leads to KATP activation, hyperpolarization of the membrane potential, and suppression of neuronal firing [47; 48]. These changes last about

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5 min after the cessation of the insult. The mechanism of KATP activation during ischemia is not yet known because KATP channels can underlie hyperpolarization even in the presence of 3 mM ATP in a patch pipette. Studies employing gene-targeting approaches, in addition to specific pharmacology, have unambiguously shown that channels containing the Kir6.2 a subunit are responsible for the transient effects of ischemia on membrane potential in neurons in the substantia nigra [47] and hippocampal CA1 [49], such that Kir6.2 knockout neurons exhibit a rapid depolarization and an increase in the firing rate rather than a suppression. Furthermore, hippocampal [49] and cortical [14] neurons from mice lacking Kir6.2 are more susceptible to ischemic damage following cerebral artery occlusion (CAO) than are neurons from wildtype mice. Conversely, cortical neurons in transgenic mice overexpressing Kir6.2 are resistant to hypoxic-ischemic injury [50]. These findings strongly support the idea that the suppression provided by endogenous KATP channels is a protective response of neurons against ischemic insults. During ischemia, there is a rapid (within 1 min) increase in extracellular glutamate to levels sufficient to activate neuronal glutamate receptors [51]. This is largely due to the dysfunction of glial glutamate uptake [52] and results in a prolonged increase in neuronal [Ca2þ]i. In striatal neurons, OGD causes a large increase in [Ca2þ]i, followed by a hyperpolarization of the membrane potential that is inhibited by a specific Slo1 blocker [53]. Transient forebrain ischemia induces persistent hyperactivity of Slo1 channels via oxidation modulation in rat hippocampal CA1 pyramidal neurons [54]. However, the mechanism is unclear as Slo1 in heterologous expression systems is inhibited by hypoxia [34]. In brain stem neurons, using cyanide treatment to mimic ischemia leads to a similar increase in [Ca2þ]i and hyperpolarization [55]. The membrane hyperpolarization is insensitive to a non-specific inhibitor of Slo1, TEA (1 mM), but fully blocked by a KATP channel blocker, indicating that Slo1 activation does not play a significant role during ischemia in these neurons. Ischemia-induced hyperpolarization seems to be mediated by both KATP and Ca2þ-activated channels in hippocampal pyramidal neurons [53]. Therefore, the contribution of Slo1 to the acute hyperpolarization and suppression of membrane excitability differs between different brain neurons. Slo1 deficient mice [44; 56] have not yet been tested for their susceptibility to ischemic stress. However, pharmacological blockade of Slo1 during ischemia aggravates subsequent neuronal damage [53]. Consistent with this, enhanced pharmacological activation decreases neuronal damage (see below). Kv2.1 undergoes dramatic functional modulation during brain ischemia. Increased [Ca2þ]i during ischemia in cultured hippocampal neurons leads to activation of calcineurin [57]. This leads to dephosphorylation of Kv2.1 in a process that takes at least 5 min to complete, but which can persist for hours. A brief episode of ischemia in vitro results in Kv2.1 dephosphorylation, which impacts the voltage-dependence of the Kv2.1 activation to potentiate somatic delayed rectifier current and effectively suppress neuronal excitability [40]. The major difference in the regulation of Kv2.1 relative to the two Kþ

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channels discussed previously is in the kinetics of the response to brain ischemia. Like KATP and Slo1 channels, Kv2.1 can also couple metabolic state and [Ca2þ]i to membrane excitability under periods of hypoxia/ischemia. However, for Kv2.1 such coupling occurs via the less direct and slower mechanism of calcineurin-dependent dephosphorylation, as opposed to the intrinsically ATP- and Ca2þ-sensitive Kþ channels Kir6.2 and Slo1. The temporal profiles of the activation of KATP and Slo1 channels fit nicely with the initial and brief hyperpolarization and resultant rapid suppression of membrane excitability. The slower but longer duration modulation of Kv2.1 may provide a secondary protection of neurons during and after a brief episode of ischemia.

10 Pharmacological Modulation As KATP channels are implicated in multiple diseases including diabetes, numerous KATP channel modulators have been developed. Most of the blockers target SUR subunits because of the initial discovery of sulfonylurea as a potent blocker of KATP channels [58]. Many chemical derivatives of sulfonylurea have been generated including tolbutamide and glibenclamide. Glibenclamide in particular exhibits sub-nanomolar binding to SUR1 and blockade of KATP channels. There are also chemicals such as diazoxide, which can activate KATP channels through their binding to the SUR1 subunit. The mechanism of the pharmacological modulation is rather complex and differs between different KATP channels types [59]. Slo1 channels are highly sensitive to TEA (IC50 of 150 mM). There are several neurotoxins and drugs that block Slo1 channels with high potency and specificity, including iberiotoxin, charybdotoxin, and slotoxin (all from scorpion venom), and paxilline (a fungal mycotoxin). The peptide scorpion toxins act as pore blockers [60], while the inhibitory mechanism of paxilline is more complex and involves allosteric modulation [61]. All of these inhibitors have nanomolar potency in modulating Slo1. Several Slo1 channel openers have also been developed, particularly for use in smooth muscle cells, with the expectation that activation of Slo1 channels should suppress membrane excitability and affect smooth muscle tone. NS1608 and NS1619 are small molecular activators of Slo1 that lead to hyperpolarization and relaxation of smooth muscle [62]. The mechanism of activation is not fully understood but seems to act independently of Ca2þ [63]. BMS-204352 is structurally similar to NS1619 and is a potent opener of Slo1 channels [64]. The action of this drug requires [Ca2þ]1 > 1 mM, suggesting that this compound could selectively target Slo1 channels in cells having elevated [Ca2þ]i. Based on the assumption that BMS-204353 could provide specific activation of Slo1 channels in ischemic neurons, BMS-204353 had been tested in the preclinical studies using rat models of brain ischemia. BMS-204353 showed a promising protective effect against the focal ischemia in

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reducing the infarct volume at 24 h after onset. Although Slo1 is highly expressed in vascular smooth muscle cells, the drug showed a minimal effect on vascular smooth muscle. BMS-204353 has more recently been tested in a phase III clinical trial (see below). Kv2.1 is also sensitive to TEA, but at millimolar concentrations, and is relatively insensitive to 4-aminopyridine (4-AP). There are no specific small molecule Kv2.1 modulators, although a number of spider toxins, including hanatoxin and stromatoxin, have been shown to act as gating modifiers of Kv2.1, as well as Kv2.2 and Kv4 channels [65].

11 Preliminary Clinical Trials KATP channel modulators such as glibenclamide have been used for years for the clinical treatment of diabetes. A recent study suggests that prior use plus post-event use of sulfonylurea agents could have a beneficial effect on stroke outcome [66]. After the discovery of the promising protective effect of the Slo1 channel opener, BMS-204352, in animal models of brain ischemia [64], the drug was subjected to a phase III clinical trial [67]. Unfortunately, there was no significant effect of the drug over placebo. No clinical trials for Kv2.1 have been undertaken.

12 Therapeutic Perspectives The rationale of targeting Kir6.2, Slo1, and Kv2.1 would be to moderate the pathological hyperexcitability of ischemic neurons and thereby increasing the survival rate of the affected neurons after an ischemic event. The major challenge in targeting these Kþ channels in therapeutic treatments is to develop compounds with sufficient specificity to localize the effect of drugs to ischemic neurons, because all of these Kþ channels are expressed widely in the brain and throughout the body. Although the clinical trial ultimately failed [67], the Slo1 opener BMS-204352 provides valuable insights into what may constitute a model drug for brain ischemia. Biophysical analyses of the drug action revealed that enhanced Slo1 channel activation occurs only at high [Ca2þ]i [64]. Therefore, this compound would be expected to elicit maximal effects in ischemic neurons where [Ca2þ]i is high relative to unaffected neurons and non-neuronal cells. Coupling the efficacy of a drug with the changes that occur in response to ischemia, such as a rise in [Ca2þ]i and a reduction in ATP, may allow for future development of drugs with specificity for ischemic neurons. One caveat of using Kþ channels as drug targets is that excessive activation of Kþ channels may have deleterious effects on neurons. Prolonged (1 h) ischemic insults, as well as other neurodegenerative insults, result in increased levels of delayed rectifier currents in cultured neurons [68], likely via enhanced

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activity of Kv2.1 [69]. Intriguingly, TEA blocks the delayed neuronal death after ischemic insult [70], indicating that the excess efflux of Kþ may trigger a signaling mechanism leading to neuronal death. Although the mechanism underlying the deleterious effect of Kþ channel activation is not fully understood, therapeutic approaches resulting in sustained enhancement of neuronal Kþ channels should be carefully considered. The aforementioned strategy of linking a drug’s efficacy to intracellular changes associated with ischemia should be effective not only for localizing the effect but also for limiting its duration. Detailed biophysical studies of the mechanism of action of channel modulators remain crucial to the development of effective new therapeutic strategies targeting brain ischemia. Acknowledgments Work from our laboratories cited above was supported by NIH (NS34383 and NS42225) and AHA.

References 1. Dirnagl U, Simon RP, and Hallenbeck JM. Ischemic tolerance and endogenous neuroprotection. Trends Neurosci 26: 248–254, 2003. 2. Fury M, Marx SO, and Marks AR. Molecular BKology: the study of splicing and dicing. Sci STKE 2002: PE12, 2002. 3. Yan J, Olsen JV, Park KS, Li W, Bildl W, Schulte U, Aldrich RW, Fakler B, and Trimmer JS. Profiling the phospho-status of the BKca channel alpha subunit in rat brain reveals unexpected patterns and complexity. Mol Cell Proteomics 7: 2188–2198, 2008. 4. Nichols CG. KATP channels as molecular sensors of cellular metabolism. Nature 440: 470–476, 2006. 5. Long SB, Tao X, Campbell EB, and MacKinnon R. Atomic structure of a voltagedependent K+ channel in a lipid membrane-like environment. Nature 450: 376–382, 2007. 6. Meera P, Wallner M, Song M, and Toro L. Large conductance voltage- and calciumdependent K+ channel, a distinct member of voltage-dependent ion channels with seven N-terminal transmembrane segments (S0–S6), an extracellular N terminus, and an intracellular (S9–S10) C terminus. Proc Natl Acad Sci USA 94: 14066–14071, 1997. 7. Salkoff L, Butler A, Ferreira G, and Wei A. High-conductance potassium channels of the SLO family. Nat Rev Neurosci 7: 921–931, 2006. 8. Inagaki N, Gonoi T, Clement JP, Namba N, Inazawa J, Gonzalez G, Aguilar–Bryan L, Seino S, and Bryan J. Reconstitution of IKATP: an inward rectifier subunit plus the sulfonylurea receptor. Science 270: 1166–1170, 1995. 9. Butler A, Tsunoda S, McCobb DP, Wei A, and Salkoff L. mSlo, a complex mouse gene encoding ‘‘maxi’’ calcium-activated potassium channels. Science 261: 221–224, 1993. 10. Frech GC, VanDongen AM, Schuster G, Brown AM, and Joho RH. A novel potassium channel with delayed rectifier properties isolated from rat brain by expression cloning. Nature 340: 642–645, 1989. 11. Karschin C, Ecke C, Ashcroft FM, and Karschin A. Overlapping distribution of K(ATP) channel-forming Kir6.2 subunit and the sulfonylurea receptor SUR1 in rodent brain. FEBS Lett 401: 59–64, 1997. 12. Zhou M, Tanaka O, Suzuki M, Sekiguchi M, Takata K, Kawahara K, and Abe H. Localization of pore-forming subunit of the ATP-sensitive K(+)-channel, Kir6.2, in rat brain neurons and glial cells. Brain Res Mol Brain Res 101: 23–32, 2002.

The Diverse Roles of Kþ Channels in Brain Ischemia

221

13. Thomzig A, Laube G, Pruss H, and Veh RW. Pore-forming subunits of K-ATP channels, Kir6.1 and Kir6.2, display prominent differences in regional and cellular distribution in the rat brain. J Comp Neurol 484: 313–330, 2005. 14. Sun HS, Feng ZP, Barber PA, Buchan AM, and French RJ. Kir6.2-containing ATPsensitive potassium channels protect cortical neurons from ischemic/anoxic injury in vitro and in vivo. Neurosci 144: 1509–1515, 2007. 15. Knaus HG, Schwarzer C, Koch RO, Eberhart A, Kaczorowski GJ, Glossmann H, Wunder F, Pongs O, Garcia ML, and Sperk G. Distribution of high-conductance Ca(2+)-activated K+ channels in rat brain: targeting to axons and nerve terminals. J Neurosci 16: 955–963, 1996. 16. Misonou H, Menegola M, Buchwalder L, Park EW, Meredith A, Rhodes KJ, Aldrich RW, and Trimmer JS. Immunolocalization of the Ca2+-activated K+ channel Slo1 in axons and nerve terminals of mammalian brain and cultured neurons. J Comp Neurol 496: 289–302, 2006. 17. Trimmer JS. Immunological identification and characterization of a delayed rectifier K+ channel polypeptide in rat brain. Proc Natl Acad Sci USA 88: 10764–10768, 1991. 18. Scannevin RH, Murakoshi H, Rhodes KJ, and Trimmer JS. Identification of a cytoplasmic domain important in the polarized expression and clustering of the Kv2.1 K+ channel. J Cell Biol 135: 1619–1632, 1996. 19. Ljubkovic M, Marinovic J, Fuchs A, Bosnjak ZJ, and Bienengraeber M. Targeted expression of Kir6.2 in mitochondria confers protection against hypoxic stress. J Physiol 577: 17–29, 2006. 20. Douglas RM, Lai JC, Bian S, Cummins L, Moczydlowski E, and Haddad GG. The calcium-sensitive large-conductance potassium channel (BK/MAXI K) is present in the inner mitochondrial membrane of rat brain. Neurosci 139: 1249–1261, 2006. 21. Drain P, Li L, and Wang J. KATP channel inhibition by ATP requires distinct functional domains of the cytoplasmic C terminus of the pore-forming subunit. Proc Natl Acad Sci USA USA 95: 13953–13958, 1998. 22. Nichols CG and Lopatin AN. Inward rectifier potassium channels. Annu Rev Physiol 59: 171–191, 1997. 23. Chapman ML and VanDongen AM. K channel subconductance levels result from heteromeric pore conformations. J Gen Physiol 126: 87–103, 2005. 24. Klemic KG, Shieh CC, Kirsch GE, and Jones SW. Inactivation of Kv2.1 potassium channels. Biophys J 74: 1779–1789, 1998. 25. Mohapatra DP and Trimmer JS. The Kv2.1 C terminus can autonomously transfer Kv2. 1-like phosphorylation-dependent localization, voltage-dependent gating, and muscarinic modulation to diverse Kv channels. J Neurosci 26: 685–695, 2006. 26. Du J, Haak LL, Phillips-Tansey E, Russell JT, and McBain CJ. Frequency-dependent regulation of rat hippocampal somato-dendritic excitability by the K+ channel subunit Kv2.1. J Physiol 522(Pt 1): 19–31, 2000. 27. Murakoshi H and Trimmer JS. Identification of the Kv2.1 K+ channel as a major component of the delayed rectifier K+ current in rat hippocampal neurons. J Neurosci 19: 1728–1735, 1999. 28. Enkvetchakul D and Nichols CG. Gating mechanism of KATP channels: function fits form. J Gen Physiol 122: 471–480, 2003. 29. Lin YF, Jan YN, and Jan LY. Regulation of ATP-sensitive potassium channel function by protein kinase A-mediated phosphorylation in transfected HEK293 cells. Embo J 19: 942–955, 2000. 30. Schubert R and Nelson MT. Protein kinases: tuners of the BKCa channel in smooth muscle. Trends Pharmacol Sci 22: 505–512, 2001. 31. Lu R, Alioua A, Kumar Y, Eghbali M, Stefani E, and Toro L. MaxiK channel partners: physiological impact. J Physiol 570: 65–72, 2006.

222

H. Misonou and J.S. Trimmer

32. King JT, Lovell PV, Rishniw M, Kotlikoff MI, Zeeman ML, and McCobb DP. Beta2 and beta4 subunits of BK channels confer differential sensitivity to acute modulation by steroid hormones. J Neurophysiol 95: 2878–2888, 2006. 33. Valverde MA, Rojas P, Amigo J, Cosmelli D, Orio P, Bahamonde MI, Mann GE, Vergara C, and Latorre R. Acute activation of Maxi-K channels (hSlo) by estradiol binding to the beta subunit. Science 285: 1929–1931, 1999. 34. Lewis A, Peers C, Ashford ML, and Kemp PJ. Hypoxia inhibits human recombinant large conductance, Ca(2+)-activated K(+) (maxi-K) channels by a mechanism which is membrane delimited and Ca(2+) sensitive. J Physiol 540: 771–780, 2002. 35. Santarelli LC, Wassef R, Heinemann SH, and Hoshi T. Three methionine residues located within the regulator of conductance for K+ (RCK) domains confer oxidative sensitivity to large-conductance Ca2+-activated K+ channels. J Physiol 571: 329–348, 2006. 36. Tang XD, Xu R, Reynolds MF, Garcia ML, Heinemann SH, and Hoshi T. Haem can bind to and inhibit mammalian calcium-dependent Slo1 BK channels. Nature 425: 531–535, 2003. 37. Hou S, Xu R, Heinemann SH, and Hoshi T. The RCK1 high-affinity Ca2+ sensor confers carbon monoxide sensitivity to Slo1 BK channels. Proc Natl Acad Sci USA 105: 4039–4043, 2008. 38. Misonou H, Mohapatra DP, Park EW, Leung V, Zhen D, Misonou K, Anderson AE, and Trimmer JS. Regulation of ion channel localization and phosphorylation by neuronal activity. Nat Neurosci 7: 711–718, 2004. 39. Park K-S, Mohapatra DP, Misonou H, and Trimmer JS. Graded regulation of the Kv2.1 potassium channel by variable phosphorylation. Science 313: 976–979, 2006. 40. Misonou H, Menegola M, Mohapatra DP, Guy LK, Park KS, and Trimmer JS. Bidirectional activity-dependent regulation of neuronal ion channel phosphorylation. J Neurosci 26: 13505–13514, 2006. 41. Miki T, Liss B, Minami K, Shiuchi T, Saraya A, Kashima Y, Horiuchi M, Ashcroft F, Minokoshi Y, Roeper J, and Seino S. ATP-sensitive K+ channels in the hypothalamus are essential for the maintenance of glucose homeostasis. Nat Neurosci 4: 507–512, 2001. 42. Gu N, Vervaeke K, and Storm JF. BK potassium channels facilitate high-frequency firing and cause early spike frequency adaptation in rat CA1 hippocampal pyramidal cells. J Physiol 580: 859–882, 2007. 43. Bean BP. The action potential in mammalian central neurons. Nat Rev Neurosci 8: 451–465, 2007. 44. Sausbier M, Hu H, Arntz C, Feil S, Kamm S, Adelsberger H, Sausbier U, Sailer CA, Feil R, Hofmann F, Korth M, Shipston MJ, Knaus HG, Wolfer DP, Pedroarena CM, Storm JF, and Ruth P. Cerebellar ataxia and Purkinje cell dysfunction caused by Ca2+-activated K+ channel deficiency. Proc Natl Acad Sci USA 101: 9474–9478, 2004. 45. Hu H, Shao LR, Chavoshy S, Gu N, Trieb M, Behrens R, Laake P, Pongs O, Knaus HG, Ottersen OP, and Storm JF. Presynaptic Ca2+-activated K+ channels in glutamatergic hippocampal terminals and their role in spike repolarization and regulation of transmitter release. J Neurosci 21: 9585–9597, 2001. 46. VanDongen AM, Frech GC, Drewe JA, Joho RH, and Brown AM. Alteration and restoration of K+ channel function by deletions at the N- and C-termini. Neuron 5: 433–443, 1990. 47. Yamada K, Ji JJ, Yuan H, Miki T, Sato S, Horimoto N, Shimizu T, Seino S, and Inagaki N. Protective role of ATP-sensitive potassium channels in hypoxia-induced generalized seizure. Science 292: 1543–1546, 2001. 48. Fujimura N, Tanaka E, Yamamoto S, Shigemori M, and Higashi H. Contribution of ATP-sensitive potassium channels to hypoxic hyperpolarization in rat hippocampal CA1 neurons in vitro. J Neurophysiol 77: 378–385, 1997. 49. Sun HS, Feng ZP, Miki T, Seino S, and French RJ. Enhanced neuronal damage after ischemic insults in mice lacking Kir6.2-containing ATP-sensitive K+ channels. J Neurophysiol 95: 2590–2601, 2006.

The Diverse Roles of Kþ Channels in Brain Ischemia

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50. Heron-Milhavet L, Xue-Jun Y, Vannucci SJ, Wood TL, Willing LB, Stannard B, Hernandez–Sanchez C, Mobbs C, Virsolvy A, and LeRoith D. Protection against hypoxic-ischemic injury in transgenic mice overexpressing Kir6.2 channel pore in forebrain. Mol Cell Neurosci 25: 585–593, 2004. 51. Jabaudon D, Scanziani M, Gahwiler BH, and Gerber U. Acute decrease in net glutamate uptake during energy deprivation. Proc Natl Acad Sci USA 97: 5610–5615, 2000. 52. Rossi DJ, Brady JD, and Mohr C. Astrocyte metabolism and signaling during brain ischemia. Nat Neurosci 10: 1377–1386, 2007. 53. Runden-Pran E, Haug FM, Storm JF, and Ottersen OP. BK channel activity determines the extent of cell degeneration after oxygen and glucose deprivation: a study in organotypical hippocampal slice cultures. Neurosci 112: 277–288, 2002. 54. Gong LW, Gao TM, Huang H, Zhuang ZY, and Tong Z. Transient forebrain ischemia induces persistent hyperactivity of large conductance Ca2+-activated potassium channels via oxidation modulation in rat hippocampal CA1 pyramidal neurons. Eur J Neurosci 15: 779–783, 2002. 55. Kulik A, Brockhaus J, Pedarzani P, and Ballanyi K. Chemical anoxia activates ATPsensitive and blocks Ca(2+)-dependent K(+) channels in rat dorsal vagal neurons in situ. Neurosci 110: 541–554, 2002. 56. Meredith AL, Thorneloe KS, Werner ME, Nelson MT, and Aldrich RW. Overactive bladder and incontinence in the absence of the BK large conductance Ca2+-activated K+ channel. J Biol Chem 279: 36746–36752, 2004. 57. Misonou H, Mohapatra DP, Menegola M, and Trimmer JS. Calcium- and metabolic state-dependent modulation of the voltage-dependent Kv2.1 channel regulates neuronal excitability in response to ischemia. J Neurosci 25: 11184–11193, 2005. 58. Aguilar-Bryan L, Nichols CG, Wechsler SW, Clement JPt, Boyd AE, 3rd, Gonzalez G, Herrera-Sosa H, Nguy K, Bryan J, and Nelson DA. Cloning of the beta cell high-affinity sulfonylurea receptor: a regulator of insulin secretion. Science 268: 423–426, 1995. 59. Bryan J, Vila-Carriles WH, Zhao G, Babenko AP, and Aguilar-Bryan L. Toward linking structure with function in ATP-sensitive K+ channels. Diabetes 53(Suppl 3): S104–S112, 2004. 60. Kaczorowski GJ and Garcia ML. Pharmacology of voltage-gated and calcium-activated potassium channels. Curr Opin Chem Biol 3: 448–458, 1999. 61. Sanchez M and McManus OB. Paxilline inhibition of the alpha-subunit of the highconductance calcium-activated potassium channel. Neuropharm 35: 963–968, 1996. 62. Hu S and Kim HS. On the mechanism of the differential effects of NS004 and NS1608 in smooth muscle cells from guinea pig bladder. Eur J Pharmacol 318: 461–468, 1996. 63. Strobaek D, Christophersen P, Holm NR, Moldt P, Ahring PK, Johansen TE, and Olesen SP. Modulation of the Ca(2+)-dependent K+ channel, hslo, by the substituted diphenylurea NS 1608, paxilline and internal Ca2+. Neuropharm 35: 903–914, 1996. 64. Gribkoff VK, Starrett JE Jr., Dworetzky SI, Hewawasam P, Boissard CG, Cook DA, Frantz SW, Heman K, Hibbard JR, Huston K, Johnson G, Krishnan BS, Kinney GG, Lombardo LA, Meanwell NA, Molinoff PB, Myers RA, Moon SL, Ortiz A, Pajor L, Pieschl RL, Post-Munson DJ, Signor LJ, Srinivas N, Taber MT, Thalody G, Trojnacki JT, Wiener H, Yeleswaram K, and Yeola SW. Targeting acute ischemic stroke with a calcium-sensitive opener of maxi-K potassium channels. Nat Med 7: 471–477, 2001. 65. Huang PT, Shiau YS, and Lou KL. The interaction of spider gating modifier peptides with voltage-gated potassium channels. Toxicon 49: 285–292, 2007. 66. Kunte H, Schmidt S, Eliasziw M, Del Zoppo GJ, Simard JM, Masuhr F, Weih M, and Dirnagl U. Sulfonylureas improve outcome in patients with type 2 diabetes and acute ischemic stroke. Stroke 38: 2526–2530, 2007. 67. Jensen BS. BMS-204352: a potassium channel opener developed for the treatment of stroke. CNS Drug Rev 8: 353–360, 2002.

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68. Yu SP, Yeh CH, Sensi SL, Gwag BJ, Canzoniero LM, Farhangrazi ZS, Ying HS, Tian M, Dugan LL, and Choi DW. Mediation of neuronal apoptosis by enhancement of outward potassium current. Science 278: 114–117, 1997. 69. Pal S, Hartnett KA, Nerbonne JM, Levitan ES, and Aizenman E. Mediation of neuronal apoptosis by Kv2.1-encoded potassium channels. J Neurosci 23: 4798–4802, 2003. 70. Wei L, Yu SP, Gottron F, Snider BJ, Zipfel GJ, and Choi DW. Potassium channel blockers attenuate hypoxia- and ischemia-induced neuronal death in vitro and in vivo. Stroke 34: 1281–1286, 2003.

Clinical Trials with Drugs Targeting Ionic Channels, Antiporters, and Pumps in Ischemic Stroke Alessandra Spinali, Giuseppe Pignataro, Gianfranco Di Renzo, and Lucio Annunziato

Despite the great concerted efforts to develop effective therapies for brain ischemia, the only successfully available therapy for acute stroke treatment is the recombinant tissue plasminogen activator (rt-PA). During the last decades, several reproducible and controlled animal models of brain ischemia have been generated and validated in an attempt to develop new effective therapies for stroke treatment. Experimental studies using neuroprotective strategies in these models have provided (1) incontrovertible proof of principle that high-grade protection of ischemic brain is an achievable goal and (2) preclinical evidence supporting potential neuroprotective efficacy [19]. However, though the preclinical evaluation of several neuroprotective agents has fostered high expectation of their clinical efficacy, translation of neuroprotective benefits from the laboratory to the clinical setting has not been successful [6]. Several preclinical and clinical factors can be considered responsible for the unsuccessful outcomes of the past clinical trials in stroke.

1 Factors to Be Considered in Preclinical Studies The main cause that has probably rendered unsuccessful the clinical trials for stroke so far has been the lack of sufficiently solid evidence-based preclinical findings. In fact, there are at least three minimal preclinical requirements to be fulfilled before an agent can be fully approved for use in human trials. First, it is necessary to demonstrate that the activity of the compound is provided with robust and clinically relevant perspectives of efficacy. Second, a proper experimental study design should be planned. Third, the confirmation of positive findings by other laboratories must be provided [6, 19, 40]. However, most of the clinical trials with neuroprotectant agents have been L. Annunziato (*) Division of Pharmacology, Department of Neuroscience, ‘‘Federico II’’ University of Naples, Via Pansini 5, 80131 Naples, Italy e-mail: [email protected]

L. Annunziato (ed.), New Strategies in Stroke Intervention, Contemporary Neuroscience, DOI 10.1007/978-1-60761-280-3_13, Ó Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009

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often launched without fully satisfying these preclinical requirements [19]. In addition to these shortcomings, a great number of variables could have negatively influenced the quality, consistency, and successful outcome of the preclinical studies. Among the most relevant of these variables are [19] the following: (a) animal-related factors: age (young but not old), sex (male but not female), comorbidities (healthy but not ill), species, and strain; (b) anesthetic agent used and co-administered drugs; (c) extent and consistency of physiological control of vital parameters; (d) model-related factors: focal versus global ischemia, transient versus permanent occlusion, method of vascular occlusion, duration and intensity of ischemia, duration and extent of reperfusion, survival time after the insult, and behavioral and pathological sequelae; (e) outcome assessment; (f) quality of study design: randomization, blinding, and analysis of the data; (g) specific properties of neuroprotective agents: stability, dose, route of administration, pharmacokinetics and pharmacodynamics, and time of administration with respect to ischemic insult (i.e., before or just immediately after the ischemic insult, in contrast to the therapeutic window of the human pathology).

2 Factors to Be Considered in Clinical Studies The main reason why clinical trials often fail to replicate the same safety and efficacy profiles of drugs tested in animal models is that the reality of clinical settings cannot reproduce the same laboratory scenarios. This is particularly true in terms of time of treatment, outcome measures, functional assessments, drug dose-regimens, and, finally, comorbidity conditions. We will now briefly describe each of these factors separately in an effort to better elucidate the reasons of their unsuccessful results in clinical trials in stroke patients. (a) Time of treatment. One of the most inconsistent factors between preclinical and clinical studies has been the time when treatment begins. Indeed, whereas in many animal studies neuroprotective agents have been given before or immediately after the onset of ischemia, in most clinical trials the time window has been beyond 3 h [6, 19]. (b) Outcome measures. The approaches to detecting the treatment outcomes have also differed considerably. Indeed, in most preclinical studies, the efficacy of neuroprotective agents has been detected by measuring the reduction of histological infarction volume. By contrast, in clinical trials neuroprotective efficacy is measured by assessing the patient’s neurological function using the NIH Stroke Scale or the modified Rankin Scale. In fact,

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infarct volumes poorly correlate with the functional outcome. Moreover, most animal stroke models used in preclinical neuroprotection studies are MCA occlusion models, whereas patients enrolled in clinical trials are often affected by damage to different brain regions. Thus, some animal models have proven to be poor predictors of clinical trial results [6, 19]. (c) Functional assessment. The reason why functional assessment may have affected the therapeutic outcomes in the clinical setting, as opposed to lab trials, is that in animal studies with neuroprotective agents, the neurocognitive function has been evaluated through different tests chosen by researchers according to their own experience. In clinical trials, although various measures have been used to determine mental and physical impairment or disability, none of these clinical scales has proven to correlate with infarct volumes or with any particular battery of preclinical functional tests [6, 19]. (d) Drug dose-regimens. To avoid toxicity, in some clinical trials a dose significantly lower than that shown to be efficient in animal studies has been administered. In addition, administration schedules for treating patients has not exactly reproduced treatment regimens used in animal studies [6, 19]. (e) Comorbidity conditions. Finally, comorbidity conditions of patients can have deeply limited the success of the therapy. In stroke models, researchers usually choose healthy young male animals. By contrast, stroke patients enrolled in ischemia clinical trials are usually old and suffer from multiple chronic diseases such as atherosclerosis, hypertension, diabetes, hyperlipemia, and prior stroke. The combination of these conditions could have eventually affected functional outcomes, thus altering the measurements of drug efficacy and safety of neuroprotectants [6, 19]. In addition, what may also have contributed to the failure of the clinical efficacy of neuoprotectants is the fact that in most cases these drugs have been tested on a number of patients by far insufficient to demonstrate clinical efficacy and without a rigorously sophisticated clinical-trial methodology [19]. Therefore, in light of these considerations, we suggest that future clinical trials involving neuroprotective agents should be designed by taking into consideration evidence derived from preclinical studies along with current guidelines for drug experimentation in stroke. These should include the following [6, 19]:

A time window of treatment compatible with the preclinical findings should be attempted, to maximize possible therapeutic benefits. If the drug is proven to be beneficial, follow-up trials with an extended therapeutic window may be designed. Drug regimens chosen for phase III efficacy studies should be compatible with preclinical doses and selected on the basis of phase I and II clinical studies. Patient selection should reflect the therapeutic target. In fact, the mechanism of injury elicited by ischemic stroke may vary because of the different brain region vulnerability. CT scans and various MRIs may be useful tools, but this has not yet been proven. Moreover, it is important to consider the severity of the stroke that may affect the study outcome. Adequate follow-up periods should be scheduled.

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Choice of clinically meaningful primary-outcome measures should be established.

Sample size should be powered to demonstrate efficacy. Choice of stratification variables should be relevant to the agent in question. Combination therapy should be taken into account. Thrombolytic therapy with rt-PA has become the standard of care. Various families of neuroprotective agents, used in combination with rt -PA, may have synergistic effects against ischemic injury. For this reason, the dose of each drug may be reduced to limit toxicity and increase tolerability; however, combining two or more drugs in the same study would render the trial design more complex. In the following tables (1–9) we will consider, among the wide array of drugs tested in humans for efficacy in stroke intervention, all clinical trials related to agents targeting ionic channels, such as calcium channel blockers, GABA antagonists, glutamate receptor antagonists, sodium channel blockers, and potassium channel openers [25]. Table 1 Clinical trials with drugs targeting ionic channels in ischemic stroke Drug class Number of clinical trials Calcium channel blockers Nimodipine Flunarizine GABA agonists Clomethiazole Glutamate receptor antagonists AMPA antagonists o YM 872 o ZK-200775 NMDA antagonists o Competitive NMDA antagonists & Selfotel o NMDA channel blockers & Aptiganel & Dextrorphan & Magnesium & NPS-1506 & Remacemide o Glycine site antagonists & ACEA 1021 & GV 150 526 Sodium channel blockers Fosphenytoin Lubeluzole Sipatrigine Potassium channel openers BMS-204352

13 2 2

2 1

2 2 1 4 1 1 1 5 1 5 3 1

Within 48 h

Within 24 h

Phase III, randomized, double-blind, placebocontrolled, multicenter clinical trial of 1,064 patients.

Prospective, multicenter, double-blind, randomized, placebocontrolled trial of 186 patients.

ANS. American Nimodipine Study [45]

Controlled trial of nimodipine in acute ischemic stroke [17]

Nimodipine

Name of the study

4 weeks

21 days

30 mg every 6 h.

60 mg, 120 mg, and 240 mg daily orally.

Table 2 Clinical trials with drugs blocking calcium channels Status of the trial: clinical Treatment: phase and number of time from Treatment patients onset duration Doses Nimodipine had no overall effect when treatment was begun within 48 h. Post hoc analysis suggested benefits for the subgroup treated with 120 mg nimodipine within 18 h, and who had negative computed tomographic scans. Mortality from all causes in the 4-week treatment group was significantly reduced with nimodipine as compared with placebo but the improvement in survival was restricted to men. A significantly better neurologic outcome was also observed in the nimodipine group. The improvement in neurological status was greatest in patients with a moderate to severe deficit at baseline.

Comments

Clinical Trials with Drugs Targeting Ionic Channels 229

Effect of nimodipine on regional cerebral glucose metabolism in patients with acute ischemic stroke as measured by positron emission tomography [23]

Double-blind study of nimodipine in non-severe stroke [4]

Name of the study

Phase II, single-center, double-blind, randomized, placebocontrolled pilot study of 60 patients. Randomized, doubleblind, placebocontrolled study of 27 patients.

Status of the trial: clinical phase and number of patients 14 days

21 days

Within 48 h

Within 48 h

Table 2 (continued) Treatment: time from Treatment onset duration

2 mg/h continuous infusion followed by 120 mg/day orally.

30 mg q.i.d. orally.

Doses

Treatment with nimodipine was well tolerated but Mathew scale scores did not differ between the two groups. The Barthel Index at the end of treatment was lower in the nimodipine group but the difference was not significant. The nimodipine group, however, showed the larger trend toward subsequent improvement. No significant differences were found in Mathew Scores. The two groups were similar in their initial metabolic depression as measured by FDG PET but glucose metabolism in non-infarcted regions increased more in the nimodipine than in the control group.

Comments

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Within 48 h

Within 24 h

Randomized, doubleblind, placebocontrolled trial of 164 patients.

Randomized, multicenter, double-blind, placebocontrolled trial of 295 patients.

INWEST. Intravenous Nimodipine West European Stroke Trial [48]

21 days

6 months

Table 2 (continued) Treatment: time from Treatment onset duration

Intravenous nimodipine in acute ischemic stroke [39]

Name of the study

Status of the trial: clinical phase and number of patients

1 or 2 mg/h intravenous infusion for 5 days followed by 120 mg/day orally for 16 days.

2 mg/h intravenous infusion for 10 days, followed by 180 mg/day orally for 6 months

Doses

Comments There were no differences between treatment groups in regard to deaths, Toronto stroke scale scores, or functional disability. There was, however, a trend toward effectiveness of nimodipine in patients treated within 12 h of stroke onset. The trial was terminated early due to safety concerns. Interim results: patients in the 2 mg/h intravenous nimodipine group showed a significantly worse outcome at 21 days compared to the placebo. Patients in the 1 mg/h intravenous nimodipine group also showed worse outcome at 21 days but the difference was not statistically.

Clinical Trials with Drugs Targeting Ionic Channels 231

28 days

Within 48 h

Within 48 h

Double-blind, placebocontrolled, multicenter trial of 164 patients.

Randomized, placebocontrolled, double-blind clinical trial of 31 patients.

Placebo-controlled trial of nimodipine in the treatment of acute ischemic cerebral infarction [31] Randomized double-blind controlled study of nimodipine in acute cerebral ischemic stroke [38] 28 days

28 days

Prospective, single-center single-blind, controlled, randomized trial of 60 patients.

28 days

Nimodipine on the clinical course of patients with acute ischemic stroke [18]

Within 12 h

Randomized, doubleblind, placebocontrolled, paralleldesigned trial of 41 patients.

Table 2 (continued) Treatment: time from Treatment onset duration

Nimodipine in acute ischemic stroke:a doubleblind controlled study [41]

Name of the study

Status of the trial: clinical phase and number of patients

120 mg/day

30 mg q.i.d. orally

120 mg/day

40 mg t.i.d.

Doses

Nimodipine was well tolerated. Mathew Scale scores showed a higher rate of improvement on nimodipine than on placebo. Mathew Scale scores revealed a highly significant difference in favor of nimodipine during the 4-week period of treatment. Mortality rates and neurologic outcome after 28 days of therapy did not differ between groups. After four weeks of treatment Mathew’s scale score improved significantly in both groups, but the difference in mean score between the two groups was insignificant.

Comments

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Within 48 h

Within 48 h

Randomized, doubleblind, multicenter, placebo-controlled trial of 350 patients.

Phase III, randomized, double-blind, placebocontrolled multicenter study of 1215 patients.

TRUST. Randomized, double-blind, placebocontrolled trial of nimodipine in acute stroke [47]

21 days

Table 2 (continued) Treatment: time from Treatment onset duration

Randomized, double-blind, placebo-controlled trial of nimodipine in acute ischemic hemispheric stroke [26]

Name of the study

Status of the trial: clinical phase and number of patients

120 mg/day orally

120 mg/day orally

Doses

There were no differences in functional outcome between the two groups. During the first month and at 3 months, there was a higher case-fatality rate in the nimodipine group than the placebo group, but at 12 months this difference had lost statistical significance. At 6 months, 55% of the nimodipine group and 58% of the placebo group were independent, the odds ratio for independence on nimodipine being 0.88 (95% confidence limits 0.70–1.10). For mortality the odds ratio with nimodipine was 1.22 (95% confidence limits 0.95–1.57).

Comments

Clinical Trials with Drugs Targeting Ionic Channels 233

Within 24 h

Within 24 h

Phase II, randomized, double-bind, placebocontrolled trial of 26 patients.

Flunarizine in acute ischemic stroke: a pilot study [29]

Within 6 h

Multicenter, double-blind, placebo-controlled trial of 331 patients.

Phase III, randomized, double-blind, placebocontrolled trial of 454 patients.

14 days

28 days

10 days

Table 2 (continued) Treatment: time from Treatment onset duration

FIST. Flunarizine in Stroke Treatment [16]

Flunarizine

VENUS.Very Early Nimodipine Use in Stroke [24]

Name of the study

Status of the trial: clinical phase and number of patients

50 mg/day intravenous infusion for 7 days followed by 3 weeks of oral treatment (week 2: 21 mg/day; week 3–4: 7 mg/day). 0.1 mg/kg intravenous bolus, followed after 3 hours by continuous infusion of 0.3 mg/kg/24 hours for 3 days, followed by 10 mg/day orally for 11 days.

30 mg q.i.d. orally.

Doses

A favorable outcome on the Rankin scale was found in 32% more patients in flunarazine group than those in control group, but this difference was not statistically significant.

No differences were found between the treatment groups.

The trial was early terminated because the results did not suggest a beneficial effect of early administration of nimodipine.

Comments

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Name of the study Within 12 h

Within 12 h

Phase III, randomized, multicenter, placebocontrolled trial of 1360 patients.

Phase III, multicenter, randomized trials involving 1200 patients (CLASS-I), 100–200 patients (CLASST) and 200 patients (CLASS-H).

CLASS. Clomethiazole Acute Stroke Study [49]

CLASS-IHT. Clomethiazole Acute Stroke Study in Ischemic, Hemorrhagic, and tPAtreated patients [25, 30]

Clomethiazole

1 day

1 day

68 mg/kg intravenous Infusion over 24 h.

75 mg/kg intravenous infusion over a 24h period.

Table 3 Clinical trails with drugs activating GABAA receptors Treatment: Status of the trial: clinical time from Treatment phase and number of patients onset duration Doses

There was no significant difference in functional outcome for either ischemic or hemorrhagic stroke patients. Clomethiazole frequently caused sedation. CLASS-I: No efficacy benefit was found in patients treated with clomethiazole. CLASS-H and T trials were not completed. Clomethiazole appeared to be safe but sedation was a common side effect. Development of clomethiazole has been halted due to the lack of efficacy.

Comments

Clinical Trials with Drugs Targeting Ionic Channels 235

Phase II safety study of ZK-200775 [15, 50]

ZK200775

ARTIST MRI. AMPA Receptor Antagonist Treatment in Ischemic Stroke [25]

ARTIST +. AMPA Receptor Antagonist Treatment in Ischemic Stroke Trial [25]

YM 872

Name of the study

Phase II, multicenter, double-blind, randomized, placebocontrolled trial of 61 patients. Dose-finding design.

Phase III, multicenter, double-blind, stratified, randomized, placebocontrolled, parallel-group trial with a planned enrollment of 600 patients. Phase II, randomized, double-blind, placebocontrolled, parallel group, multicenter trial with a planned enrollment of 260 patients.

Over 2 days

1 day

Within 6 h

Within 24 h

1 day

Within 3 h

Group 1: Total dose of 262.5 mg (i.v.) in 48 h; Group 2: Total dose of 525 mg (i.v.) in 48 h; Group 3: Total dose of 105 mg (i.v.) over a period of 6 h.

1.25 mg/kg/h intravenous infusion.

1.25 mg/kg/h intravenous infusion in combination with tPA.

Table 4 Clinical trials with drugs targeting glutamate receptors: AMPA antagonists Status of the trial: clinical Treatment: phase and number of time from Treatment patients onset duration Doses

Trial stopped prematurely due to safety concerns. ZK200775 worsened patients’ neurological condition with possible glial cell toxicity. It exerts significant sedative effects.

The trial was abandoned after failing an interim futility analysis.

The trial was abandoned after failing an interim futility analysis.

Comments

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Within 6 h

Within 12 h

Phase III, randomized, multicenter, placebocontrolled, parallel-design studies in 567 patients.

Phase II, randomized, multicenter, double-blind, placebo-controlled, ascending-dose phase IIa trial of 32 patients.

ASSIST. Acute Stroke Studies Involving Selfotel Treatment [7, 8, 32]

Safety and tolerability of the glutamate antagonist CGS 19755 (selfotel) in patients with acute ischemic stroke [20]

Selfotel

Name of the study

1 day

1 day

Intravenous bolus of 1 mg/kg 2/1 day, 2 mg/kg, 1.75 mg/ kg, or 1.5 mg/kg

Single dose of 1.5 mg/ kg intravenous infusion

Trials terminated after interim analysis showed no therapeutic benefit and increase in mortality (particularly within the first 30 days and in patients with severe stroke) in selfotel group. Selfotel-treated patients had a higher incidence of agitation and hallucinations. A single intravenous dose of 1.5 mg/kg selfotel was safe and tolerable. Agitation, hallucinations, confusion, paranoia, and delirium occurred in all patients treated with 2.0 mg/kg. A significantly higher percentage of treated patients achieved an independent Barthel Index score at day 90 than placebo patients.

Table 5 Clinical trials with drugs targeting glutamate receptors: competitive NMDA antagonists Status of the trial: clinical Treatment: phase and number of time from Treatment patients onset duration Doses Comments

Clinical Trials with Drugs Targeting Ionic Channels 237

Safety, tolerability, and pharmacokinetics of the N-methyl-D-aspartate antagonist dextrorphan in patients with acute stroke. Dextrorphan Study Group [1]

Multicenter, ascendingdose study in 67 patients.

Within 48 h

Within 24 h

Double-blind, randomized, placebo-controlled, ascending doses, multicenter trial of 46 patients.

Safety and tolerability study of aptiganel hydrochloride in patients with an acute ischemic stroke [14]

Dextrorphan

Within 6 h

Phase II–III, multicenter, double-blind, placebocontrolled, randomized, trial of 628 patients.

Aptiganel acute stroke trial [3]

Aptiganel

Name of the study

1 day

1 day

12 h

1-h loading dose (60–150 mg) followed by a 23-h ascending-dose maintenance infusion. 1-h loading dose (145–260 mg) followed by an 11-h constant rate infusion (30–70 mg/h).

High-dose: 5-mg bolus followed by 0.75 mg/h for 12 h; Low-dose: 3-mg bolus followed by 0.5 mg/h for 12 h. 3, 4.5, 6 and 7.5 mg intravenous bolus followed by constant infusion for 6–12 h.

Loading-dose escalation was stopped because of rapid-onset, reversible, symptomatic hypotension in 7 of 21 patients treated with doses of 200–260 mg/h. There was no difference in neurological outcome at 48 h between the dextrorphan-treated and placebo-treated patients.

A 4.5-mg intravenous bolus of aptiganel followed by infusion of 0.75 mg/h for 12 was founded out a tolerable dose.

Clinical trial halted because an interim analysis showed no improvement in outcome at 90 days.

Table 6 Clinical Trials with drugs targeting glutamate receptors: NMDA channel blockers Status of the trial: clinical Treatment: phase and number of time from Treatment patients onset duration Doses Comments

238 A. Spinali et al.

Within 12 h

Within 2 h

Within 12 h

Within 12 h

Phase 2, open-label, pilot study in 20 patients.

Phase III, multicenter, randomized, doubleblind, placebocontrolled trial ongoing. Study Size Planned: 1298

Phase III, multicenter, double-blind, randomized placebocontrolled trial of 2589 patients.

Phase III, single-center, randomized, doubleblind, placebocontrolled pilot trial of 60 patients.

FAST-MAG Field Administration of Stroke Therapy – Magnesium Phase III Trial [25]

IMAGES Intravenous MAGnesium Efficacy in Stroke [37]

Randomized, double-blind, placebo-controlled pilot trial of intravenous magnesium sulfate in acute stroke [34]

1 day

1 day

1 day

1 day

Table 6 (continued) Treatment: time from Treatment onset duration

FAST-MAG Pilot Study. Field Administration of Stroke TherapyMagnesium [44]

Magnesium

Name of the study

Status of the trial: clinical phase and number of patients Paramedic-administered dose of 2.5 g magnesium plus an additional 1.5 g bolus, followed by a maintenance infusion of 16 g magnesium. 4 g bolus dose of magnesium delivered over 15 min by paramedics in the field, followed by an inhospital infusion of 16 g magnesium delivered over 24 h. Intravenous magnesium sulfate in a 16-mmol bolus infused over 15 min, followed by a 65mmol maintenance dose over 24 h. 8 mmol intravenous over 15 min followed by 65 mmol over 24 h.

Doses

Magnesium was well tolerated, with no significant adverse effects and no change in blood pressure or pulse rate.

Primary outcome was not improved by intravenous magnesium sulfate.

Trial Ongoing. At this writing 651 patients have been enrolled.

Good functional outcome at 3 months occurred in 60%. Field-based magnesium intervention is safe and feasible.

Comments

Clinical Trials with Drugs Targeting Ionic Channels 239

Remacemide hydrochloride: a double-blind, placebocontrolled, safety and tolerability study in patients with acute ischemic stroke [12]

Remacemide

Phase Ib trial of NPS 1506 [33]

NPS 1506

Name of the study

Placebo-controlled, dose escalating, parallel group study in 61 patient.

Phase Ib, double-blind, placebo-controlled, ascending dose study in 36 patients.

Status of the trial: clinical phase and number of patients

Within 12 h

Within 48 h

3 days

Table 6 (continued) Treatment: time from Treatment onset duration

100, 200, 300, 400, 500, or 600 mg in 6 intravenous infusions ( 2 doses per day for 3 days).

60, 80, or 100 mg, infused over 60 min.

Doses

The most common adverse events were related to the CNS, and these events appeared to increase with dose.

NPS 1506 was well tolerated at all three doses and there were no serious adverse events.

Comments

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GAIN 2. Glycine Antagonist in Neuroprotection-2 [46, 25]

GAIN 1. Glycine Antagonist in Neuroprotection-1 [46, 25]

GV 150526 (Gavestinel)

Dose escalation study of the NMDA glycine-site antagonist licostinel in acute ischemic stroke [2]

ACEA 1021 (Licostinel)

Name of the study

Phase II, randomized, multicenter, doubleblind, placebocontrolled, dose escalation study on 56 patients. Phase II, randomized, multicenter, doubleblind, placebocontrolled study on 109 patients.

Dose-escalation, placebocontrolled, randomized study in 64 patients.

Within 12 h

Within 12 h

Within 48 h

3 days

3 days

1 day

800-mg loading dose followed by 200 mg every 12 h

Group 1: 400 mg; Group 2: 800 mg; Group 3: 1000 mg

Low-doses: 0.03–0.60 mg/ kg; High-doses: 1.2–3.0 mg/kg

The incidence of serious adverse events was similar in the drug and placebo groups. Hyperbilirubinemia was reported in 6% of GV150526-treated patients compared with 3% of placebo-treated patients. Outcome at 4 weeks after stroke was better in GV150526treated patients, but the

Higher doses were associated with neurological and gastrointestinal complaints. A similar improvement in National Institutes of Health Stroke Scale scores over time was seen in both the placebo group and the licostineltreated patients.

Table 7 Clinical Trials with drugs targeting glutamate receptors: NMDA glycine site antagonists Status of the trial: clinical Treatment: phase and number of time from Treatment patients onset duration Doses Comments

Clinical Trials with Drugs Targeting Ionic Channels 241

Glycine antagonist in neuroprotection for patients with acute Stroke: GAIN Americas: a randomized controlled trial [43] Glycine antagonist (gavestinel) in neuroprotection (GAIN International) in patients with acute stroke: a randomized controlled trial. GAIN International Investigators [27] Safety and tolerability of GV150526 (a glycine site antagonist at the Nmethyl-D-aspartate receptor) in patients with acute stroke [13]

Name of the study

Phase II, randomized, placebo-controlled, parallel-group, ascending-dose study on 66 patients and conducted in 2 phases.

Phase III, stratified, randomized, international, multicenter, doubleblind, placebo controlled trials of 1367 patients. Phase III, Multicenter, randomized, doubleblind, parallel-group, placebo-controlled trial of 1804 patients.

Status of the trial: clinical phase and number of patients

3 days

3 days

Within 6 h

Within 12 h

3 days

Within 6 h

Table 7 (continued) Treatment: time from Treatment onset duration

Part A: loading doses of 50, 100, 200, 400, or 800 mg. Part B: loading dose of 800 mg followed by maintenance infusion.

Intravenous loading dose of 800 mg followed by 200 mg every 12 h for five doses.

Intravenous loading dose of 800 mg followed by 5 maintenance doses of 200 mg every 12 h.

Doses

GV150526 was well tolerated and no serious adverse events related to drug therapy were reported. The study was designed to assess safety and not designed to test efficacy.

There were no significant differences in serious side effects between the groups. Treatment with gavestinel within 6 h of acute ischemic stroke did not improve outcome.

studies were not powered to show statistical significance. No difference between the two group has been showed.

Comments

242 A. Spinali et al.

Multinational randomised controlled trial of lubeluzole in acute ischemic stroke. European and Australian Lubeluzole Ischemic Stroke Study Group [11] Lubeluzole in acute ischemic stroke. A double-blind, placebo-controlled phase II trial. Lubeluzole International Study Group [10]

LUB-INT-13 Lubeluzole in ischemic stroke [9]

Lubeluzole

Fosphenytoin phase 3 multicenter study to evaluate the safety and efficacy of intravenous fosphenytoin in patients with acute ischemic stroke [25, 42]

Fosphenytoin

Name of the study

Randomized, doubleblind, placebocontrolled trial of 232 patients.

Phase III international, multicenter, placebocontrolled trial of 1786 patients. Multicenter, double-blind, placebo-controlled trial of 725 patients.

Phase III, randomized, double-blind, multicenter trial of 462 patients.

5 days

5 days

Within 6 h

Within 6 h

5 days

Within 8 h

Within 4 h

Infusion of 7.5 mg over 1 h followed by 10 mg/day.

Loading infusion of 7.5 mg followed by a continuous day infusion of 10 mg. Infusion of 7.5 mg over 1 h followed by 10 mg/day.

Intravenous

Table 8 Clinical trials with drugs blocking sodium channels Status of the trial: clinical Treatment: phase and number of time from Treatment patients onset duration Doses

The trial, initially aimed at a patients inclusion of 270, was terminated prematurely because of an imbalance in mortality between the treatment group.

Treatment with lubeluzole did not affect mortality or clinical outcome in the overall study population.

No significant differences in any safety or efficacy endpoints were founded.

Enrollment halted when interim analysis found no differences between groups.

Comments

Clinical Trials with Drugs Targeting Ionic Channels 243

Phase II clinical trial of sipatrigine (619C89) by continuous infusion in acute stroke [36]

Phase II, randomized, placebo-controlled study of 27 patients.

Within 12 h

Within 3 h

Phase II, multicenter, placebo-controlled, randomized trial of 200 patients.

LUB-USA-6. Combination therapy with lubeluzole and t-PA in the treatment of acute ischemic stroke [21]

Sipatrigine

Within 6 h

Phase III, multicenter, randomized, doubleblind, placebocontrolled study involving 721 patients.

over 65 h

5 days

5 days

Table 8 (continued) Treatment: time from Treatment onset duration

Lub. US and Canadian lubeluzole ischemic stroke study [22]

Name of the study

Status of the trial: clinical phase and number of patients Doses

10, 18, 27, or 36 mg/kg by continuous intravenous infusion.

Infusion of 7.5 mg followed by 10 mg/5 day.

7.5 mg over 1 h, followed by a continuous daily infusion of 10 mg for up to 5 days.

Comments

Neuropsychiatric effects occurred in 16 of 21 patients receiving sipatrigine. No effects on outcome measures were demonstrated.

The overall mortality rate at 12 weeks for lubeluzoletreated patients was 20.7% compared to 25.2% for placebotreated patients (Not significant). Lubeluzole treatment resulted in significantly greater improvements in functional status (Barthel Index) and overall disability (Rankin Scale) after 12 weeks. The study was halted after lack of efficacy demonstrated for lubeluzole alone.

244 A. Spinali et al.

Safety and tolerability of 619C89 after acute stroke [35]

SIS. Sipatrigine in Stroke [25]

Name of the study

Phase II trial of 170 patients. Part 1: Double-blind, placebo controlled, multicenter, escalating dose trial. Part 2: Double-blind, placebo controlled, parallel group, multicenter study. Randomized, ascendingdose, placebo-controlled trial of 48 patients.

Status of the trial: clinical phase and number of patients

Dose escalation was stopped after the occurrence of hallucinations in 5 of 18 patients who received 2 mg/kg/8 h or more.

Within 12 h

Comments Trial halted.

Intravenous loading dose followed by maintenance doses given 8 hourly.

Doses

Within 12 h

64 h

Table 8 (continued) Treatment: time from Treatment onset duration

Clinical Trials with Drugs Targeting Ionic Channels 245

Post-011: Efficacy and safety of MaxiPostTM in patients with acute stroke [5]

BMS-204352

Name of the study

Phase III, randomized, Multicenter, doubleblind, placebocontrolled, parallel group study in 1978 patients. Within 6 h

Over 72 h

1 mg or 0,1 mg intravenous.

Table 9 Clinical trials with drugs opening potassium channels Status of the trial: clinical Treatment: time phase and number of from onset Treatment patients duration Doses

Study halted after negative results.

Comments

246 A. Spinali et al.

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247

References 1. Albers GW, Atkinson RP, Kelley RE, and Rosenbaum DM. Safety, tolerability, and pharmacokinetics of the N-methyl-D-aspartate antagonist dextrorphan in patients with acute stroke. Dextrorphan Study Group. Stroke 26: 254–258, 1995. 2. Albers GW, Clark WM, Atkinson RP, Madden K, Data JL, and Whitehouse MJ. Dose escalation study of the NMDA glycine-site antagonist licostinel in acute ischemic stroke. Stroke 30: 508–513, 1999. 3. Albers GW, Goldstein LB, Hall D, and Lesko LM. Aptiganel Acute Stroke Investigators. Aptiganel hydrochloride in acute ischemic stroke: a randomized controlled trial. J Am Med Assoc 286: 2673–2682, 2001. 4. Bogousslavsky J, Regli F, Zumstein V, and Kobberling W. Double-blind study of ¨ nimodipine in non-severe stroke. Eur Neurol 30: 23–26, 1990. 5. Bozik M, Homme Ml, Grotta J, Fisher M, Fayad P, Bogousslavsky J, Brunell R, Fernandes L, Colby S, Noonan T, and Douglass A. Post-011: efficacy and safety of MaxiPostTM in Patients with Acute Stroke. JNS 187: S255, 2001. 6. Cheng YD, Al-Khoury L, and Zivin JA. Neuroprotection for ischemic stroke: two decades of success and failure. NeuroRx 1: 36–45, 2004. 7. Davis SM, Albers GW, Diener HC, Lees KR, and Norris J. Termination of acute stroke studies involving selfotel treatment. ASSIST Steering Committed. Lancet 349: 32, 1997. 8. Davis SM, Lees KR, Albers GW, Diener HC, Markabi S, Karlsson G, and Norris J. Selfotel in acute ischemic stroke: possible neurotoxic effects of an NMDA antagonist. Stroke 31: 347–354, 2000. 9. Diener HC, Cortens M, Ford G, Grotta J, Hacke W, Kaste M, Koudstaal PJ, and Wessel T. Lubeluzole in acute ischemic stroke treatment: A double-blind study with an 8-h inclusion window comparing a 10-mg daily dose of lubeluzole with placebo. Stroke 31: 2543–2551, 2000. 10. Diener HC, Hacke W, Hennerici M, Radberg J, Hantson L, and De Keyser J. Lubeluzole in acute ischemic stroke. A double-blind, placebo-controlled phase II trial. Lubeluzole International Study Group. Stroke 27: 76–81, 1996. 11. Diener HC. Multinational randomised controlled trial of lubeluzole in acute ischaemic stroke. European and Australian Lubeluzole Ischaemic Stroke Study Group. Cerebrovasc Dis 8: 172–181, 1998. 12. Dyker AG and Lees KR. Remacemide hydrochloride: a double-blind, placebo-controlled, safety and tolerability study in patients with acute ischemic stroke. Stroke 30: 1796–1801, 1999. 13. Dyker AG and Lees KR. Safety and tolerability of GV150526 (a glycine site antagonist at the N-methyl-D-aspartate receptor) in patients with acute stroke. Stroke 30: 986–992, 1999. 14. Dyker AG, Edwards KR, Fayad PB, Hormes JT, and Lees KR. Safety and tolerability study of aptiganel hydrochloride in patients with an acute ischemic stroke. Stroke 30: 2038–2042, 1999. 15. Elting JW, Sulter GA, Kaste M, Lees KR, Diener HC, Hommel M, Versavel M, Teelken AW, and De Keyser J. AMPA antagonist ZK200775 in patients with acute ischemic stroke: possible glial cell toxicity detected by monitoring of S-100B serum levels. Stroke 33: 2813–2818, 2002. 16. Franke CL, Palm R, Dalby M, Schoonderwaldt HC, Hantson L, Eriksson B, LangJenssen L, and Smakman J. Flunarizine in stroke treatment (FIST): a double-blind, placebo-controlled trial in Scandinavia and the Netherlands. Acta Neurol Scand 93: 56–60, 1996. 17. Gelmers HJ, Gorter K, de Weerdt CJ, and Wiezer HJ. A controlled trial of nimodipine in acute ischemic stroke. N Engl J Med 318: 203–207, 1988.

248

A. Spinali et al.

18. Gelmers HJ. The effects of nimodipine on the clinical course of patients with acute ischemic stroke. Acta Neurol Scand 69: 232–239, 1984. 19. Ginsberg MD. Neuroprotection for ischemic stroke: past, present and future. Neuropharmacology 55: 363–389, 2008. 20. Grotta J, Clark W, Coull B, Pettigrew LC, Mackay B, Goldstein LB, Meissner I, Murphy D, and La Rue L.. Safety and tolerability of the glutamate antagonist CGS 19755 (Selfotel) in patients with acute ischemic stroke. Results of a phase IIa randomized trial. Stroke 26: 602–605, 1995. 21. Grotta J. Combination Therapy Stroke Trial Investigators. Combination Therapy Stroke Trial: recombinant tissue-type plasminogen activator with/without lubeluzole. Cerebrovasc Dis 12: 258–263, 2001. 22. Grotta J. Lubeluzole treatment of acute ischemic stroke. The US and Canadian Lubeluzole Ischemic Stroke Study Group. Stroke 28: 2338–2346, 1997. 23. Heiss WD, Holthoff V, Pawlik G, and Neveling M. Effect of nimodipine on regional cerebral glucose metabolism in patients with acute ischemic stroke as measured by positron emission tomography. J Cereb Blood Flow Metab 10: 127–132, 1990. 24. Horn J, de Haan RJ, Vermeulen M, and Limburg M. Very Early Nimodipine Use in Stroke (VENUS): a randomized, double-blind, placebo-controlled trial. Stroke 32: 461–465, 2001. 25. Internet Stroke Center. Stroke Trials Registry. http://www.strokecenter.org/trials/index. aspx, 2009. 26. Kaste M, Fogelholm R, Erila¨ T, Paloma¨ki H, Murros K, Rissanen A, and Sarna S. A randomized, double-blind, placebo-controlled trial of nimodipine in acute ischemic hemispheric stroke. Stroke 25: 1348–1353, 1994. 27. Lees KR, Asplund K, Carolei A, Davis SM, Diener HC, Kaste M, Orgogozo JM, and Whitehead J. Glycine antagonist (gavestinel) in neuroprotection (GAIN International) in patients with acute stroke: a randomised controlled trial. GAIN International Investigators. Lancet 355: 1949–1954, 2000. 28. Lees KR. Cerestat and other NMDA antagonists in ischemic stroke. Neurology 49: S66–S69, 1997. 29. Limburg M and Hijdra A. Flunarizine in acute ischemic stroke: a pilot study. Eur Neurol 30: 121–2, 1990. 30. Lyden P, Jacoby M, Schim J, Albers G, Mazzeo P, Ashwood T, Nordlund A, and Odergren T. The Clomethiazole Acute Stroke Study in tissue-type plasminogen activator-treated stroke (CLASS-T): final results. Neurology 57: 1199–1205, 2001. 31. Martı´ nez-Vila E, Guille´n F, Villanueva JA, Matı´ as-Guiu J, Bigorra J, Gil P, Carbonell A, and Martı´ nez-Lage JM. Placebo-controlled trial of nimodipine in the treatment of acute ischemic cerebral infarction. Stroke 21: 1023–1028, 1990. 32. Morris GF, Bullock R, Marshall SB, Marmarou A, Maas A, and Marshall LF. Failure of the competitive N-methyl-D-aspartate antagonist Selfotel (CGS 19755) in the treatment of severe head injury: results of two phase III clinical trials. The Selfotel Investigators. J Neurosurg 91: 737–743, 1999. 33. Mueller AL, Artman LD, Balandrin MF, Brady E, Chien Y, DelMar EG, Kiersted A, Marriott TB, Moe ST, Raszkiewicz JL, VanWagenen B, and Wells D. NPS 1506, a moderate affinity uncompetitive NMDA receptor antagonist: preclinical summary and clinical experience. Amino Acids 19: 177–179, 2000. 34. Muir KW and Lees KR. A randomized, double-blind, placebo-controlled pilot trial of intravenous magnesium sulfate in acute stroke. Stroke 26: 1183–1188, 1995. 35. Muir KW, Hamilton SJ, Lunnon MW, Hobbiger S, and Lees KR. Safety and tolerability of 619C89 after acute stroke. Cerebrovasc Dis 8: 31–37, 1998. 36. Muir KW, Holzapfel L, and Lees KR. Phase II clinical trial of sipatrigine (619C89) by continuous infusion in acute stroke. Cerebrovasc Dis 10: 431–436, 2000.

Clinical Trials with Drugs Targeting Ionic Channels

249

37. Muir KW, Lees KR, Ford I, and Davis S. Intravenous Magnesium Efficacy in Stroke (IMAGES) Study Investigators. Magnesium for acute stroke (Intravenous Magnesium Efficacy in Stroke trial): randomised controlled trial. Lancet 363: 439–445, 2004. 38. Nag D, Garg RK, and Varma M. A randomized double-blind controlled study of nimodipine in acute cerebral ischemic stroke. Indian J Physiol Pharmacol 42: 555–558, 1998. 39. Norris JW, LeBrun LH, and Anderson BA. Intravenous nimodipine in acute ischaemic stroke. Cerebrovasc Dis 4: 194–196, 1994. 40. O’Collins VE, Macleod MR, Donnan GA, Horky LL, van der Worp BH, and Howells DW. 1,026 experimental treatments in acute stroke. Ann Neurol 59: 467–477, 2006. 41. Paci A, Ottaviano P, Trenta A, Iannone G, De Santis L, Lancia G, Moschini E, Carosi M, Amigoni S, and Caresia L. Nimodipine in acute ischemic stroke: a double-blind controlled study. Acta Neurol Scand 80: 282–286, 1989. 42. Pulsinelli W, Mann M, Welch M, et al. Fosphenytoin in acute ischemic stroke: efficacy results. Neurology 52: A384, 1999. 43. Sacco RL, DeRosa JT, Haley EC Jr, Levin B, Ordronneau P, Phillips SJ, Rundek T, Snipes RG, and Thompson JL. Glycine Antagonist in Neuroprotection Americas Investigators. Glycine antagonist in neuroprotection for patients with acute stroke: GAIN Americas: a randomized controlled trial. J Am Med Assoc 285: 1719–1728, 2001. 44. Saver JL, Kidwell C, Eckstein M, and Starkman S. FAST-MAG Pilot Trial Investigators. Prehospital neuroprotective therapy for acute stroke: results of the Field Administration of Stroke Therapy-Magnesium (FAST-MAG) pilot trial. Stroke 35: 106–108, 2004. 45. The American Nimodipine Study Group. Clinical trial of nimodipine in acute ischemic stroke. Stroke 23: 3–8, 1992. 46. The North American Glycine Antagonist in Neuroprotection (GAIN) Investigators. Phase II studies of the glycine antagonist GV150526 in acute stroke: the North American experience. Stroke 31: 358–365, 2000. 47. Trust Study Group. Randomised, double-blind, placebo-controlled trial of nimodipine in acute stroke. Lancet 336: 1205–1209, 1990. 48. Wahlgren NG, MacMahon DG, De Keyser J, Indredavik B, and Ryman T. Intravenous Nimodipine West European Stroke Trial (INWEST) of nimodipine in the treatment of acute ischaemic stroke. Cerebrovasc Dis 4: 204 –210, 1994. 49. Wahlgren NG, Ranasinha KW, Rosolacci T, Franke CL, van Erven PM, Ashwood T, and Claesson L. Clomethiazole acute stroke study (CLASS): results of a randomized, controlled trial of clomethiazole versus placebo in 1360 acute stroke patients. Stroke 30: 21–28, 1999. 50. Walters MR, Kaste M, Lees KR, Diener HC, Hommel M, De Keyser J, Steiner H, and Versavel M. The AMPA antagonist ZK 200775 in patients with acute ischaemic stroke: a double-blind, multicentre, placebo-controlled safety and tolerability study. Cerebrovasc Dis 20: 304–309, 2005.

Index

A Acidosis, 1, 16, 18, 117, 118, 120, 153, 154, 158, 162–164, 165, 166, 167, 168 Acid-sensing ionic channels (ASIC), vii, 3, 21, 153–168 Amiloride, 30, 73, 74, 77, 78, 120, 123, 124, 154, 158, 159, 161, 165, 166, 167 a-amino-3-hydroxy-5-methyl-4isoxazolepropionate (AMPA), 2, 4, 5, 13, 20, 21, 94, 96, 101–102, 119, 228, 236 Amlodipine, 200 Apoptosis, vi, 1, 2, 4, 5, 6, 14, 18, 36, 38, 118, 139, 140, 143, 198, 199

B BCMP1, 28, 32 Benzothiazepines, 193, 200 Blood-brain-barrier, v, 167 BMS-204353, 218, 219

C Ca2+-activated K+channels, 75, 211, 217 Ca2 cation antiporter (CaCA), 65 Ca2+/cation exchanger family (CCX), 65 CACNA1, 192, 193 Ca2+/H+exchange family (CAX), 65 Ca2+/H+exchange (YRBG), 65 Calcineurin, 53, 67, 68, 116, 117, 157, 158, 215, 217, 218 Calcium, vi, 1, 3, 15, 20, 21, 27, 29, 34, 35, 37, 38, 39, 47–58, 129, 130, 132, 136, 138, 139, 140, 142, 143, 144, 166, 189, 190, 192, 193, 198, 199, 200, 202, 228, 229 Calcium/calmodulin-dependent protein kinase II (CaMKII), 117, 157, 199

Caldendrins, 54 Calexcitin, 53 Calmodulin, 53, 116, 117, 157, 198, 199, 215 Calpains, 2, 4 Canonical (TRPC), 175 Ca2+ overload, 3, 55, 77, 100, 166, 216 Ca2 overload, 34, 39 Cariporide, 30, 39, 124 Caspases, 2, 4 Cation-chloride cotransporter superfamily (CCC), 89 CaV, 191, 192, 193, 194, 195, 196, 197, 200 Cell death, 1, 2, 3, 4, 5, 6, 7, 14, 15, 17, 19, 20, 27, 36, 38, 40, 90, 98, 101, 105, 117, 119, 121, 125, 140, 143, 175, 181, 182, 185, 198, 199 Cerebral ischemia, 1–8, 14, 16, 17, 40, 65–83, 89–106, 113–125, 153, 164, 182, 183, 184, 189, 198, 199, 201, 202 Charybdotoxin, 218 CICR, 53, 54, 55 Cl– ATPase, 7 Clot, v Conopeptides, 199 o-conotoxin, 193, 194, 199

D Dihydropyridines, 75, 190, 193, 199, 200 Diltiazem, 200

E Edema, v, 2, 5, 15, 103, 105, 118, 123, 125 Endoplasmic reticulum (ER), 4, 47–58, 130, 131, 132, 198 ER Ca2+-ATPases, 4, 75

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252 Exchanger inhibitory peptide (XIP), 69, 72, 75, 79 Excitotoxicity, 1, 5, 13, 14, 16, 17–20, 21, 36, 37, 95, 96, 97, 98, 101–102, 104, 123, 153

F Felodipine, 200 Flunarizine, 200, 201, 228, 234 Focal cerebral ischemia, 1, 7, 16, 17, 102

G GABA-gated Cl– channels, 7 Gabapentin, 202 Glibenclamide, 28, 36, 218, 219 GluRs, 53 Glutamate, vi, vii, 2, 3, 4, 5, 6, 7, 13–22, 36, 37, 53, 72, 94, 96, 97, 98, 99, 101, 106, 123, 162, 164, 166, 175, 181, 191, 198, 202, 211, 217, 228, 236, 237 Glutamate excitotoxicity, vi, 14, 36, 37

H Haemorrhage, v Hanatoxin, 219 HOE642, 120, 122, 124 Homeostasis, vi, vii, 1, 2, 3–7, 14, 15, 16, 47–58, 71, 75, 77, 78, 82, 90, 91, 93, 95, 100, 103, 105, 113, 118, 125, 139, 166, 180, 181, 199 Homer, 53

I Iberiotoxin, 28, 218 Infarction core, 1 Inositol-1,4,5-trisphosphate receptors (InsP3Rs), 48, 49, 50, 51, 52, 53, 54, 55, 57, 58 Inositol triphosphate (IP3), 4 Inward rectifying K+ (Kir) channels, 211 Ion homeostasis, 1, 15, 75, 103, 105 Ischemia, 15, 18, 21, 27–40, 129, 137, 140, 141, 142, 143, 144, 160, 162, 167, 168, 183, 200, 211–220, 225, 226, 227 Isradipine, 200, 201

K K+ channels, vii, 6, 8, 21, 32, 75, 159, 211–220 K+ channel openers, 6, 228

Index K+–Cl- cotransporters, 7 K+ dependent Na+/ Ca2+ exchanger (NCKXs), 3 K+-selective ion channels, vi, 211 KB-R7943, 74, 75, 78, 79, 101, 120 Kir6.1, 32 Kir6.2, 32, 36, 212–215, 216, 217, 219 Kir6.2 gene, 211–212 Kurtoxin, 199 Kv2.1, 212–215, 216, 217, 218, 219 Kv2.1 gene, 211–212

L Lamotrigine, 200, 201 Levetiracetam, 200, 201 L-type calcium channels, 198

M Melastatin (TRPM), 175 Membrane-associated guanylate kinase (MAGUK), 191 Mibefradil, 199, 200 Middle Cerebral Artery Occlusion (MCAO), 36, 67, 71, 76, 78, 106, 121, 142 Mitochondria, 2, 4, 5, 6, 27, 29, 30, 32, 33, 34, 35, 36, 37, 39, 40, 101, 102, 138, 182, 198, 214 Mitochondrial channels, 27–40 Mitochondrial dysfunction, 2, 8, 14 Mitochondrial Na+/H+ exchanger, 39 Mitochondrial Permeability Transition Pore (PTP), 34–35 MitoKAT, 28, 32, 35, 36–37, 39 MVIIA (SNX-111), 199

N Na+/Ca2+ exchanger K+-dependent family (NCKX), 65 Na+/Ca2+ exchangers (NCXs), vii, 3, 4, 21, 65, 66, 69, 70, 71, 72, 76–81, 82, 100, 101, 120, 125 NCX1, 66, 67, 69, 70, 71, 73, 74, 75, 77, 79, 80, 104 NCX2, 66, 67, 69, 70, 71, 73, 74, 75, 77, 79, 80, 82 NCX3, 66, 67–68, 69, 70, 71, 73, 74, 75, 77, 79, 80, 81 Na+/Ca2+–Li+ exchanger (NCLX or NCKX6), 65 Na+ dependent Cl-/HCO3– exchangers, 7

Index Na+/H+ exchanger (NHEs), vii, 3, 21, 113 Na+/K+-ATPase, vii, 5, 21, 75, 78, 92, 100, 124, 129–144 Na+/K+/Ca2+/Cl- cotransporters (NKCCs), vii, 3, 92 Na+-K+-Cl– cotransporter (NKCC), 89–106, 123 NAADP receptors, 48 Necrosis, vi, 5, 14, 17, 36, 38 Neurons, 2, 3, 7, 20, 37, 38, 39, 67, 68, 70, 71, 72, 81, 89, 91, 93, 94, 95, 102, 118, 119, 123, 129, 132, 140, 141, 143, 155, 156, 157, 161–162, 166, 168, 178, 179, 182, 189, 192, 194, 196, 197, 198, 199, 211, 212, 213, 214, 216, 217, 219 Neuroprotection, v, vii, 16, 18, 19, 39, 40, 97, 104, 129, 140, 141, 142, 143, 162, 164, 175, 185, 200, 201, 202, 227, 241, 242 Neurotrophins, 53 NHE, 3, 21, 73, 74, 75, 113, 118, 120, 124, 125 Nicardipine, 200, 201 Nickel, 199 Nicotinic acid adenine dinucleotide phosphate (NAADP), 48, 53 Nifedipine, 98, 199, 200 Nimodipine, 199, 200, 201, 202, 228, 229, 230, 231, 232, 233, 234 Nitric oxide, 4, 53, 136, 158 Nitric oxide synthase, 3, 164 NKCC, 21, 89, 90, 91, 106 NKCC1, 5, 89, 90, 91, 92–104, 123 NKCC2, 89, 106 NMDA, 2, 4, 13, 14, 15, 16, 18, 19, 20, 21, 22, 75, 78, 94, 96, 97, 98, 101, 102, 143, 162, 164, 165, 182, 183, 201, 228, 237, 238, 241 NMDARs, 175, 181 182, 185, 195 N-methyl-D-aspartate (NMDA), 2, 195, 238 NS1608, 218 NS1619, 28, 218 Nuclear factor-kB (NFkB), 67

253 Permanent middle cerebral artery occlusion (pMCAO), 76, 77, 80 Phenylalkylamine, 193, 200 Pimozide, 199 Plasma membrane, 3, 4, 16, 27, 28, 29, 30, 32, 65, 71, 72, 73, 89, 90, 93, 100, 113, 115, 116, 130, 131, 132, 136, 137, 139, 140, 154, 178, 191 Potassium channel openers, 6, 228 P/Q-type channels, 198, 201 Preconditioning, 6, 139 Pregabalin, 200, 202 Protein kinases, 53, 69, 117, 136, 157, 177, 199, 215 PTP (permeability transition pore), 27, 29, 34–35, 38–40

R RNS (Reactive Nitrogen Species), vi, 72, 180, 182 ROS (Reactive Oxygen Species), vi, 2, 4, 6, 27, 28, 36, 37, 38, 39, 72, 114, 180, 182 rTPA, 81 Ryanodine receptors (RyRs), 48, 49–55, 56–57

O Oxidative stress, 2, 3, 4, 5, 6, 33, 34, 92, 106 Oxygen glucose deprivation (OGD), 7, 36, 37, 39, 40, 78, 96, 100, 102, 141, 182, 199, 217

S Sabiporide, 123 SEA0400, 74, 75, 78, 79 SERCA, 48, 49, 50, 52, 54, 56, 58 SERCA (Sarco(Endo) plasmic reticulum calcium ATPase), 48 Slo1, 211–215, 216, 217, 218, 219 Slo1 gene, 211 Slotoxin, 218 SM-20220, 122, 123, 124 SN-6, 74, 75, 79 Stroke, v, vi, vii, 1, 13–22, 36–40, 47–58, 76–81, 95–104, 118–123, 129–144, 153–168, 181–183, 198–199, 216–218, 225–246 Stromatoxin, 219 Sulfonylurea receptors SUR1, 3, 32, 212, 213, 214, 215, 216, 218 SUR2, 32

P Paxilline, 28, 218 Penumbra, vi, 1, 5, 15, 16, 17, 18, 19, 78, 80, 81, 143

T TEA, 217, 218, 219, 220 Thrombolysis, v Tolbutamide, 218

254 tPA, v, 144, 181, 184, 235, 236 Transient receptor potential channels (TRPC), vii, 3, 21 Transient receptor potential (TRP), 3, 21, 176, 177, 178 Trimeric intracellular cation channels (TRICs), 48 TRPM7, 175–185 TRPV1 channels, 48 TWIK-related potassium channels (TREK), 3 TY-12533, 124 U UCPs, 27, 28, 32–34, 35, 36, 37–38, 39 UCP1, 32 UCP2, 28, 34, 38 UCP3, 34 UCP4, 28, 32, 34 UCP5, 28, 32, 34

Index V Vanilloid (TRPV), 57, 177 Verapamil, 200 Very Early Nimodipine Use in Stroke (VENUS), 201, 234 Voltage-dependent Ca2 channels (VDCC), 2, 3, 4, 7, 98 Voltage-dependent Cl- channels, 7 Voltage-gated calcium channels (VGCC), 189, 192, 193 Voltage-gated K+ (Kv) channels, 211 Voltage operated Ca2+ channels (VOCC), vii

Y YM-244769, 74, 75

Z Zinc, 15, 199