Free Radical Effects on Membranes (Current Topics in Membranes, Volume 61)

Current Topics in Membranes, Volume 61 Series Editors Dale J. Benos Department of Physiology and Biophysics University of Alabama Birmingham, Alabama

Sidney A. Simon Department of Neurobiology Duke University Medical Care Durham, North Carolina

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Current Topics in Membranes, Volume 61

Free Radical Effects on Membranes Edited by Sadis Matalon Department of Anesthesiology School of Medicine University of Alabama at Birmingham Birmingham, AL 35233, USA Rakesh P. Patel Division of Molecular & Cellular Pathology Department of Pathology University of Alabama at Birmingham Birmingham, AL

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Contents Contributors ix Previous Volumes in Series xiii

I. Overview

CHAPTER 1 Structure and Functions of Biomembranes James F. Collawn and Zsuzsa Bebök

I. II. III. IV.

Cell Membrane Structure and Function 1 Overview of Membrane Functions 4 Calcium Signaling 14 Oxidative Stress and Organelle Dysfunction 15 References 19

CHAPTER 2 The Interaction of Reactive Oxygen and Nitrogen Species with Membranes Matias N. Möller, Jack R. Lancaster and Ana Denicola

I. Reactive Oxygen and Nitrogen Species 23 II. Physical Interactions: Compartmentalizing Reactivity 25 III. Chemical Effects: Lipid Peroxidation 35 References 39

II. Interaction of RONS with Channels and Pumps

CHAPTER 3 Modulation of Lung Epithelial Sodium Channel Function by Nitric Oxide Weifeng Song, Ahmed Lazrak, Shipeng Wei, Phillip McArdle and Sadis Matalon

I. Introduction 44 References 62 v

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CHAPTER 4 Effects of Nitrogen Oxides on Chloride Channels Benjamin Gaston

I. Overview 71 II. Introduction 72 III. Regulation by Nitrogen Oxides of Swelling-activated Cl− Channels 73 IV. Regulation of Calcium Activated Cl− Channels by Nitrogen Oxides (NOx ) 74 V. Effects of Nitrogen Oxides on CFTR 76 VI. Future Directions 82 References 83

CHAPTER 5 A Mitochondria-AOS-Kv Channel Axis in Health and Disease; New Insights and Therapeutic Targets for Vascular Disease and Cancer Gopinath Sutendra and Evangelos D. Michelakis

I. Introduction 87 II. The Components of the Mitochondria-ROS-Kv Axis 88 III. The Mitochondria-AOS-Kv Axis in Hypoxia: HPV IV. The Mitochondria-AOS-Kv Axis, Metabolism and Apoptosis 97 References 108

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CHAPTER 6 Oxidative Modification of Ca2+ Channels, Ryanodine Receptors, and the Sarco/Endoplasmic Reticulum Ca2+ -ATPase Victor S. Sharov and Christian Schöneich

I. Introduction 114 II. Overview of Ca2+ Translocation Membrane Proteins 114 III. Ca2+ Channels 115 IV. SERCA 119 V. PMCA 124 VI. Concluding Remarks 125 References 125

CHAPTER 7 Regulation of Na,K-ATPase by Reactive Oxygen Species Guofei Zhou, Laura A. Dada and Jacob I. Sznajder

I. Na,K-ATPase 132 II. Na,K-ATPase in Alveolar Fluid Reabsorption 133

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III. Role of Reactive Oxygen Species in Signaling 134 IV. Regulation of Na,K-ATPase and Alveolar Fluid Reabsorption by ROS 136 V. Dopamine and β-adrenergic Agonists Improve ROS-Mediated Decrease in Alveolar Fluid Reabsorption 140 VI. Summary 141 References 141

III. RONS and Membrane Permeability CHAPTER 8 Reactive Oxygen Species and Endothelial Permeability Masuko Ushio-Fukai, Randall S. Frey, Tohru Fukai and Asrar B. Malik

I. Introduction 148 II. Generation and Metabolism of ROS 151 III. ROS Generating System in ECs (NADPH Oxidase) 151 IV. Regulation of Adherens Junctions (AJs) by Phosphorylation and by Rho GTPase 153 V. ROS-generating Stimulants which Regulate Endothelial Permeability 154 VI. ROS Reducing Factors/Proteins which Block Endothelial Permeability 165 VII. Molecular Targets of ROS Regulating Endothelial Permeability 165 VIII. Mediators/Regulators of ROS-dependent Endothelial Permeability 171 IX. Functional Significance of ROS-dependent Endothelial Permeability in Vivo 173 X. Summary and Conclusions 175 References 176

IV. RONS and Signal Transduction CHAPTER 9 Cell Signaling by Oxidants: Pathways Leading to Activation of Mitogen-activated Protein Kinases (MAPK) and Activator Protein-1 (AP-1) Arti Shukla and Brooke T. Mossman

I. Introduction 192 II. MAPK Signaling 195

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III. Mitogen-activated Protein Kinase Phosphatases (MKPs) 198 IV. Relationships between MAPK and Activator Protein-1 (AP-1) 199 V. Conclusions 203 References 204

CHAPTER 10 The Interaction of Mitochondrial Membranes with Reactive Oxygen and Nitrogen Species Paul S. Brookes, Andrew P. Wojtovich, Lindsay S. Burwell, David L. Hoffman and Sergiy M. Nadtochiy

I. Mitochondria as a Source of Reactive Species 212 II. Effects of ROS and RNS on Mitochondrial Respiration 217 III. Mitochondrial Membrane Lipids 221 IV. ROS, RNS & Mitochondrial Ion Transport 224 V. Complex Interactions & Concluding Remarks 230 References 232

CHAPTER 11 Oxidant Stress and Airway Epithelial Function Jenora T. Waterman and Kenneth B. Adler

I. II. III. IV. V. Index 257

Introduction 243 Sources of Reactive Oxygen Species 244 Antioxidant Defenses in Airway Epithelium 245 Oxidant-induced Airway Epithelial Responses 247 Conclusions 251 References 251

Contributors Numbers in parentheses indicate the pages on which the author’s contribution begin.

Kenneth B. Adler (245), Department of Molecular Biomedical Sciences, North Carolina State University, College of Veterinary Medicine, 4700 Hillsborough Street, Raleigh, NC 27606, USA Zsuzsa Bebök (1), Department of Cell Biology, University of Alabama at Birmingham, 1918 University Blvd, Birmingham, AL 35294-0005, USA Paul S. Brookes (213), Department of Anesthesiology, University of Rochester Medical Center Lindsay S. Burwell (213), Department of Biochemistry & Biophysics, University of Rochester Medical Center James F. Collawn (1), Department of Cell Biology, University of Alabama at Birmingham, 1918 University Avenue, Birmingham, AL 35294-0005, USA Laura A. Dada (133), Division of Pulmonary and Critical Care Medicine, Feinberg School of Medicine, Northwestern University, 240 E. Huron, McGaw Pavilion M-326, Chicago, IL 60611, USA Ana Denicola (23), Lab. Fisicoquímica Biológica, Facultad de Ciencias, Universidad de la Republica, Igua 4225, 11400 Montevideo, Uruguay David L. Hoffman (213), Department of Biochemistry & Biophysics, University of Rochester Medical Center Randall S. Frey (149), Department of Pharmacology and the Center for Lung and Vascular Biology, University of Illinois, Chicago, IL 60605, USA Tohru Fukai (149), Department of Pharmacology and the Center for Lung and Vascular Biology, University of Illinois, Chicago, IL 60605, USA Benjamin Gaston (73), University of Virginia School of Medicine, Charlottesville, VA 22908, USA Jack R. Lancaster Jr. (23), Departments of Anesthesiology, Physiology & Biophysics, and Environmental Health Sciences and Center for Free Radical Biology, University of Alabama at Birmingham, AL, USA ix

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Contributors

Ahmed Lazrak (45), Department of Anesthesiology, School of Medicine, University of Alabama at Birmingham, Birmingham, AL 35233, USA Asrar B. Malik (149), Department of Pharmacology and the Center for Lung and Vascular Biology, University of Illinois, Chicago, IL 60605, USA Sadis Matalon (45), Department of Anesthesiology, School of Medicine, University of Alabama at Birmingham, Birmingham, AL 35233, USA Phillip McArdle (45), Department of Anesthesiology, School of Medicine, University of Alabama at Birmingham, Birmingham, AL 35233, USA Evangelos D. Michelakis (89), Department of Medicine, University of Alberta, Edmonton, Canada Brooke T. Mossman (183), University of Vermont College of Medicine, Department of Pathology, 89 Beaumont Avenue, Burlington, VT 05405, USA Matías N. Möller (23), Lab. Fisicoquímica Biológica, Facultad de Ciencias, Universidad de la Republica, Igua 4225, 11400 Montevideo, Uruguay Sergiy M. Nadtochiy (213), Department of Anesthesiology, University of Rochester Medical Center Victor S. Sharov (115), Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, KS 66047, USA Christian Schöneich (115), Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, KS 66047, USA Arti Shukla (183), University of Vermont College of Medicine, Department of Pathology, 89 Beaumont Avenue, Burlington, VT 05405, USA Weifeng Song (45), Department of Anesthesiology, School of Medicine, University of Alabama at Birmingham, Birmingham, AL 35233, USA Gopinath Sutendra (89), Department of Medicine, University of Alberta, Edmonton, Canada Jacob I. Sznajder (133), Division of Pulmonary and Critical Care Medicine, Feinberg School of Medicine, Northwestern University, 240 E. Huron, McGaw Pavilion M-326, Chicago, IL 60611, USA Masuko Ushio-Fukai (149), Department of Pharmacology and the Center for Lung and Vascular Biology, University of Illinois, Chicago, IL 60605, USA

Contributors

Jenora T. Waterman (245), Department of Molecular Biomedical Sciences, North Carolina State University, College of Veterinary Medicine, 4700 Hillsborough Street, Raleigh, NC 27606, USA Shipeng Wei (45), Department of Anesthesiology, School of Medicine, University of Alabama at Birmingham, Birmingham, AL 35233, USA Andrew P. Wojtovich (213), Department of Pharmacology & Physiology, University of Rochester Medical Center Guofei Zhou (133), Division of Pulmonary and Critical Care Medicine, Feinberg School of Medicine, Northwestern University, 240 E. Huron, McGaw Pavilion M-326, Chicago, IL 60611, USA

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Previous Volumes in Series Current Topics in Membranes and Transport Volume 23 Genes and Membranes: Transport Proteins and Receptors∗ (1985) Edited by Edward A. Adelberg and Carolyn W. Slayman Volume 24 Membrane Protein Biosynthesis and Turnover (1985) Edited by Philip A. Knauf and John S. Cook Volume 25 Regulation of Calcium Transport across Muscle Membranes (1985) Edited by Adil E. Shamoo Volume 26 Na+ –H+ Exchange, Intracellular pH, and Cell Function∗ (1986) Edited by Peter S. Aronson and Walter F. Boron Volume 27 The Role of Membranes in Cell Growth and Differentiation (1986) Edited by Lazaro J. Mandel and Dale J. Benos Volume 28 Potassium Transport: Physiology and Pathophysiology∗ (1987) Edited by Gerhard Giebisch Volume 29 Membrane Structure and Function (1987) Edited by Richard D. Klausner, Christoph Kempf, and Jos van Renswoude Volume 30 Cell Volume Control: Fundamental and Comparative Aspects in Animal Cells (1987) Edited by R. Gilles, Arnost Kleinzeller, and L. Bolis Volume 31 Molecular Neurobiology: Endocrine Approaches (1987) Edited by Jerome F. Strauss, III, and Donald W. Pfaff Volume 32 Membrane Fusion in Fertilization, Cellular Transport, and Viral Infection (1988) Edited by Nejat Düzgünes and Felix Bronner Volume 33 Molecular Biology of Ionic Channels∗ (1988) Edited by William S. Agnew, Toni Claudio, and Frederick J. Sigworth Volume 34 Cellular and Molecular Biology of Sodium Transport∗ (1989) Edited by Stanley G. Schultz ∗ Part of the series from the Yale Department of Cellular and Molecular Physiology.

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Previous Volumes in Series

Volume 35 Mechanisms of Leukocyte Activation (1990) Edited by Sergio Grinstein and Ori D. Rotstein Volume 36 Protein–Membrane Interactions∗ (1990) Edited by Toni Claudio Volume 37 Channels and Noise in Epithelial Tissues (1990) Edited by Sandy I. Helman and Willy Van Driessche

Current Topics in Membranes Volume 38 Ordering the Membrane Cytoskeleton Trilayer∗ (1991) Edited by Mark S. Mooseker and Jon S. Morrow Volume 39 Developmental Biology of Membrane Transport Systems (1991) Edited by Dale J. Benos Volume 40 Cell Lipids (1994) Edited by Dick Hoekstra Volume 41 Cell Biology and Membrane Transport Processes∗ (1994) Edited by Michael Caplan Volume 42 Chloride Channels (1994) Edited by William B. Guggino Volume 43 Membrane Protein–Cytoskeleton Interactions (1996) Edited by W. James Nelson Volume 44 Lipid Polymorphism and Membrane Properties (1997) Edited by Richard Epand Volume 45 The Eye’s Aqueous Humor: From Secretion to Glaucoma (1998) Edited by Mortimer M. Civan Volume 46 Potassium Ion Channels: Molecular Structure Function, and Diseases (1999) Edited by Yoshihisa Kurachi, Lily Yeh Jan, and Michel Lazdunski Volume 47 AmilorideSensitive Sodium Channels: Physiology and Functional Diversity (1999) Edited by Dale J. Benos Volume 48 Membrane Permeability: 100 Years since Ernest Overton (1999) Edited by David W. Deamer, Arnost Kleinzeller, and Douglas M. Fambrough Volume 49 Gap Junctions: Molecular Basis of Cell Communication in Health and Disease Edited by Camillo Peracchia

Previous Volumes in Series

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Volume 50 Gastrointestinal Transport: Molecular Physiology Edited by Kim E. Barrett and Mark Donowitz Volume 51 Aquaporins Edited by Stefan Hohmann, Søren Nielsen and Peter Agre Volume 52 Peptide–Lipid Interactions Edited by Sidney A. Simon and Thomas J. McIntosh Volume 53 CalciumActivated Chloride Channels Edited by Catherine Mary Fuller Volume 54 Extracellular Nucleotides and Nucleosides: Release, Receptors, and Physiological and Pathophysiological Effects Edited by Erik M. Schwiebert Volume 55 Chemokines, Chemokine Receptors, and Disease Edited by Lisa M. Schwiebert Volume 56 Basement Membrances: Cell and Molecular Biology Edited by Nicholas A. Kefalides and Jacques P. Borel Volume 57 The Nociceptive Membrane Edited by Uhtaek Oh Volume 58 Mechanosensitive Ion Channels, Part A Edited by Owen P. Hamill Volume 59 Mechanosensitive Ion Channels, Part B Edited by Owen P. Hamill Volume 60 Computational Modeling of Membrane Bilayers Edited by Scott E. Feller

Preface

Rakesh P. Patel, Sadis Matalon

Free radicals (reactive molecules with unpaired electrons in their outer orbits) are typically associated with cellular damage and death of biological organisms with perturbation of membrane integrity being key in their toxicity. Indeed, the concept that increased free radical production and the ensuing destruction of biomolecules, concomitant with depleted antioxidant defenses has shaped much thinking and efforts in elucidating the molecular mechanisms leading to a variety of disease. More recently however, with a better understanding of free radical biology, it has become apparent that the effects of free radicals (encompassing reactive oxygen species, reactive nitrogen species, reactive halogen intermediates) depend on their concentration, location, redox potential as well the biochemical composition of the target. Thus it is now clear that reactive species formation and metabolism are controlled events and central in redox-cell signaling pathways that impact diverse cellular, functions encompassing cellular, physiological and pathological responses. Interactions among reactive species and membrane components (including proteins and lipids) exemplify this paradigm. Moreover, emerging data implicate cellular membranes as critical foci for regulating the reactivity of free radical species by either regulating reactive species formation (e.g. mitochondria), determining vectorial reactive species production (e.g. respiratory burst in activated neutrophils) and providing a hydrophobic milieu which dramatically alters reactivity of various reactive species (e.g. nitric oxide). It was our goal to solicit contributions from leaders in the field that collectively discuss these concepts both in general terms and by focusing on specific examples of how reactive species interactions with specific membrane proteins can modulate cell-signaling in physiological and pathological contexts. Thus, this volume consist of eleven chapters from experts in the field that encompass free-radical effects on diverse membrane functions, ranging from selective barrier functions, controlling membrane protein function to discussing how the hydrophobic environment within membranes regulate free radical reactivity. In soliciting reviews we purposely stayed away from assembling a book on free-radicals and lipid peroxidation, a topic on which multiple research articles, xvii

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reviews and books have focused on and an area that remains an active focus of investigation. Instead we decided to focus on articles focused in discussing specific examples in which membranes from different cellular compartments (e.g. plasma, ER, mitochondria) and membrane proteins either regulate reactive species formation and reactivity or are specific targets of reactive species leading to alteration in function. The latter typically involves post-translation modification of specific amino acid residues resulting in selective and specific alteration of protein function. The book starts with chapters that provide overviews on biomembranes and the impact their physico-chemical properties have on reactive species reactivity (Chapters 1 and 2). This is followed by a series of chapters (Chapters 3–7) that illustrate the concept that different reactive species can modulate function of specific membrane ion channels in different tissues. This includes sodium channels, chloride channels, sodium-potassium ATPases and calcium channels in both plasma and intracellular organelle membranes. Importantly, the implications for reactive species control of ion channel function in vascular and pulmonary diseases are also discussed. We then focus on the role of reactive species in influencing a major role of membranes that being their selective barrier function and regulation of permeability (Chapter 8). In the final three chapters (Chapters 9–11) the focus is on reactive species and control of cell-signaling pathways. Specific topics that illustrate this concept include control of MAP kinases and down-stream signaling, role of the mitochondria as a source an target for reactive species signaling and finally, the role of reactive species in airway epithelial function. We would like to thank all the contributors for their efforts in compiling this book and hope that the final product provides the readership with a current-view of the how integral biological membranes are as sources, targets and transducers of reactive species biology and how a deeper understanding of this interplay is increasing our understanding of molecular mechanisms of diseases together with potentially offering new therapeutic targets.

CHAPTER 1 Structure and Functions of Biomembranes James F. Collawn and Zsuzsa Bebök Department of Cell Biology, University of Alabama at Birmingham, 1918 University Blvd, Birmingham, AL 35294-0005, USA

I. Cell Membrane Structure and Function A. Plasma Membrane B. Mitochondrial Membranes C. Peroxisomal Membranes II. Overview of Membrane Functions A. Permeability B. Ion Transport C. Signal Transduction III. Calcium Signaling IV. Oxidative Stress and Organelle Dysfunction A. Oxidative Stress and the ER B. Oxidative Stress and Peroxisomes C. Oxidative Stress and Mitochondria D. Oxidative Stress and Lysosomes E. Summary and Conclusions References

I. CELL MEMBRANE STRUCTURE AND FUNCTION Cell membrane systems provide two important functions: (1) they establish a biological barrier to the extracellular environment and (2) they compartmentalize specialized and sometimes toxic biological reactions within the cell. Although the different cellular membrane systems have diverse biological functions, they do share some common features. All membrane systems are composed of a lipid bilayer that contains a full complement of protein complexes that facilitate permeability, transport, and signaling. The focus of this chapter will be to describe recent advances in our understanding of cell membrane structure and function, and to highlight the specialized roles of mitochondria and peroxisomes. FurtherCurrent Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00201-9

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more, we will discuss how structure and function of these membrane systems are affected by oxidative stress.

A. Plasma Membrane Our contemporary view of the plasma membrane was first described more than 30 years ago by the fluid mosaic model of Singer and Nicholson (1972). Many of the features proposed by the original model still ring true today, including the basic structure. The fluid mosaic model predicted that there is a random distribution of molecular components in the membrane that have free lateral and rotational movement (Singer and Nicolson, 1972; Vereb et al., 2003). More recent studies, however, indicate that many membrane protein components are in large supramolecular complexes that are either tethered to the actin cytoskeleton or have limited lateral diffusion (Damjanovich et al., 1999; Vereb et al., 2003). Furthermore, elegant studies using single-particle tracking techniques indicate that even lipids experience transient confinement in lipid rafts (Dietrich et al., 2002). Based on these very sensitive biophysical techniques, it has become clear that the distribution of lipids and proteins in the plasma membrane is highly organized, dynamic, and nonrandom (Figure 1). Rapid advances in microscopy such as fluorescence recovery after photobleaching (FRAP), optical tracking by laser tweezers, single-particle tracking techniques, and confocal laser-scanning microscopy have aided in this revised view of the nature of the plasma membrane (reviewed in Vereb et al., 2003). Proteins, therefore, may be freely mobile or constrained by lipid rafts, by large protein complexes, or by the

FIGURE 1 Modified fluid-mosaic model of cell membrane.

1. Structure and Functions of Biomembranes

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cytoskeleton itself. The protein organization within these large complexes is certain to affect their biological function, particularly with regard to signaling within lipid rafts. Lipid raft structures are important components of the plasma membrane and consist of ∼50–70 nm membrane patches that are enriched in sphingolipids and cholesterol. Because of the nature of the fatty acid side chains found in these sphingolipids, lipid rafts result in a thicker membrane structure where certain proteins have a tendency to accumulate, particularly signaling complexes (Simons and Toomre, 2000). Furthermore, lipid rafts change their size and protein composition in response to extracellular stimuli, resulting in the activation of various signaling cascades (Simons and Toomre, 2000). Oxidative stress and its resultant effects on lipid structure in lipid rafts can therefore have profound effects on cellular signaling cascades.

B. Mitochondrial Membranes Mitochondria consist of a double membrane system, with an extensive inner membrane surface area. The mitochondrial membrane system is second only to the endoplasmic reticulum membrane surface area, often consisting of up to 40% of the total membrane in a cell. Because of this vast membrane network, mitochondria occupy a large portion of the cytoplasmic volume of cells, and in many cell types, are constantly changing shape. Rather than the rod-like structures often depicted in textbooks, they are more often seen as elongated stringlike structures. Mitochondrial movement occurs in a number of cell types and is mediated by microtubules. The dynamic nature of these organelles is best illustrated using videomicroscopy, which demonstrates that mitochondria constantly change shape, fuse with one another, or divide in two. Although the mitochondrial genome encodes for 33 genes, the vast majority of proteins are made by the cell and are imported by the mitochondrial transporters of the outer and inner membranes, the TOM and TIM complexes. The most commonly associated function of mitochondria is in the production of ATP from oxidative phosphorylation. In this process, pyruvate and fatty acids are broken down to acetyl CoA, and in the inner matrix of the mitochondria, acetyl CoA is metabolized in the citric acid cycle, generating NADH and FADH2 . Highenergy electrons from NADH and FADH2 are then passed along the electrontransport chain on the mitochondrial inner membrane surface, generating a proton gradient across the membrane. The electro-chemical gradient across the inner membrane is then used by ATP synthase, also on the inner membrane surface, to generate ATP from ADP. The electro-chemical gradient is also important for the import of newly synthesized “mitochondrial” proteins made from the cell’s genomic DNA. Given the essential function of mitochondria, it is not surprising that a large number of seemingly unrelated disorders such as schizophrenia,

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Alzheimer’s disease, epilepsy, Parkinson’s disease, cardiomyopathy, and diabetes have a common component, namely the production of reactive oxygen species and mitochondrial DNA damage (reviewed in Pieczenik and Neustadt, 2007).

C. Peroxisomal Membranes Peroxisomes are small, single membrane organelles that contain a number of oxidative enzymes. This membrane system consists of only about 1% of a cell’s membrane system and functionally overlaps with mitochondria in certain biological reactions such as fatty acid β-oxidation. One unique function is in the production of plasmalogens, the most abundant lipid found in myelin. In the liver and kidney, peroxisomes also aid in the detoxification of acids, aldehydes, and alcohols via catalase and H2 O2 . The essential nature of peroxisomes is best illustrated in the human inherited disease Zellweger syndrome, a condition in which the protein import machinery in peroxisomes is defective. This loss of function leads to “empty” peroxisomes and neurological, kidney and liver abnormalities. Death occurs soon after birth.

II. OVERVIEW OF MEMBRANE FUNCTIONS All organelle membrane systems function to maintain their distinctive composition from the cytosol by providing a barrier to most polar molecules. Transport of materials (or signals) requires protein receptors/channels/transporters that span the lipid bilayer. In the following section, we will discuss how the membrane systems facilitate transport of small molecules through permeability and ion transport, and then discuss how signal transduction is facilitated through a membrane system using mechanotransduction and MAP kinases as examples.

A. Permeability The hydrophobic character of a membrane that consists of phospholipids, cholesterol, and glycolipids prevents the passive diffusion of most polar, watersoluble molecules. This critical function maintains intracellular ion concentrations, as well as organelle identity and function. The rate at which a molecule diffuses across a membrane system depends on its size and how nonpolar it is. For example, O2 and CO2 readily diffuse across membrane systems, whereas small, uncharged polar molecules such as H2 O or urea diffuse across membrane systems much more slowly. Large uncharged molecules such as glucose, and ions such as Na+ , K+ , or Cl− require carrier proteins or channels for effective transport across the bilayer.

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B. Ion Transport Channel or carrier proteins are required for ion transport for two different types of transport, active and passive. In passive transport, ions are transported down their concentration gradient, from high concentration to low. Since ions are charged, however, membrane potential also influences their transport. For plasma membranes, the voltage gradient places a negative potential on the inner membrane surface, and a positive on the outer surface, facilitating positive ion entry, and negative ion exit. Transport of ions against their electrochemical gradient, on the other hand, requires energy expenditure, usually in the form of ATP hydrolysis. Active transporters can be classified as uniporters, symporters, or antiporters. Uniporters transport one solute from one side of the membrane to the other, while symporters transport two solutes in the same direction, and antiporters transport two solutes in opposite directions. In the case of two solutes, one of the solutes is usually transported down its electrochemical gradient, and this free energy is used to help transport the other solute against its electrochemical gradient. The bestcharacterized example of this is in glucose transport driven by a Na+ gradient. In this case, Na+ is transported down its electrochemical gradient into the cell (from 145 outside to 10 mM inside), and glucose is co-transported in the process. The Na+ gradient is maintained by an ATP-driven Na+ pump which pumps Na+ back out of the cell to maintain this gradient. This coupled carrier system provides active transport that is driven by the Na+ gradient. A similar system is used by the Na+ –H+ exchanger, which couples Na+ influx with H+ efflux as a means of maintaining a cytosolic pH of 7.2. Another approach for controlling pH within organelles such as endosomes and lysosomes is to couple ATP hydrolysis with proton transport, as is the case with ATP-driven H+ pump. A more complicated process involves the transport of solutes across epithelial cells. In transcellular transport, co-transport of glucose and Na+ from the intestinal lumen across the epithelial cell to the blood requires pumps at the apical and basolateral surfaces. At the apical surface, Na+ and glucose is co-transported into the cell as described above. At the basolateral surface, glucose is transported out of the cell down its concentration gradient by passive transport through a carrier protein, while Na+ is transported out by active transport by the Na+ –K+ pump. The tight junctions help maintain the concentration gradients generated by the pumps.

C. Signal Transduction Signal transduction pathways provide a mechanism for transmitting signals from the extracellular environment to cells or conveying signals between cells. These signals are transmitted a number of ways, but the most common is via

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ligand-receptor interactions. Ligand binding “transduces” a signal across the membrane bilayer either by activating intrinsic activity of the receptor itself, or by activating an associated protein. Once activated, this sets off an activation cascade that employs a variety of intracellular signaling proteins that convey the signal to the appropriate targets within the cell. Not all ligands bind a cell surface receptor for signal transduction, and a good example of this is nitric oxide (NO). NO readily diffuses across the cell membrane and binds to iron in the active site of guanylyl cyclase, stimulating the production of cyclic GMP, and thus activating the signaling cascade. 1. Mechanotransduction In some cases, transmission of a signal across a bilayer is not mediated by ligand binding, but rather mechanotransduction. In mechanotransduction, a mechanical force exerted on the cell membrane is converted into electrical or biochemical signals (reviewed in Martinac, 2004; Orr et al., 2006). Probably the best-studied example of this is the stretch-sensitive ion channels (Martinac, 2004). The first evidence for mechanically gated channels comes from studies of mechanosensory neurons (Katz, 1950). Patch clamp analysis first allowed for measurement of single mechanosensitive channels and demonstrated that there were two types of channels: stretch-activated and stretch-inactivated ion channels (Sachs and Morris, 1998). Mechanical forces are transduced along the plane of the cell membrane (membrane tension), rather than hydrostatic pressure (Martinac, 2004). Two models have been proposed to explain channel gating, the bilayer model and the tethered model (Hamill and McBride Jr., 1997). In the bilayer model, lipid bilayer tension is all that is required for activation. Whereas in the tethered model, the channel must be in contact with the cytoskeleton or extracellular matrix before activation occurs. The two models are not mutually exclusive since the mechanism of activation may depend on the channel type. The amount of membrane tension required for half activation of most of the known mechanosensitive channels is several dynes/cm (10−3 N/m) (Martinac, 2004; Sachs, 1988), an amount easily resulting from differences in the transmembrane osmolarity of only a few milliosmols (Martinac, 2004). Mechanotransduction, much like phosphorylation, uses conformational changes in the target molecule(s) to mediate signaling events (Orr et al., 2006). The conformational changes are often associated with channels that contain linkages to the cytoskeleton or extracellular matrix, and these interactions amplify small mechanical forces into displacement of large complexes, resulting in signal transduction (Orr et al., 2006). In stretch-sensitive channels, the mechanism for signal transduction may be even simpler. Increasing tension in the lipid bilayer from 10–12 dyn/cm to 20 dyn/cm (Evans et al., 1976) increases the open-probability of the channel (Martinac and Hamill, 2002).

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Two possibilities have been proposed to explain how membrane tension triggers channel opening (Orr et al., 2006). In the first, if an open channel occupies a greater surface area in the bilayer than in the closed state, i.e., if it is more unfolded or expanded, then the free energy of the open state will be lower (Orr et al., 2006). In the second possibility, tension causes the bilayer to be thinner by 0.15 nm, and this triggers channel opening (Martinac and Hamill, 2002). In this scenario, the hydrophobic transmembrane domain of the channel must be thinner in the open state, and channel opening therefore would be favored in order to avoid the hydrophobic mismatch that could occur between the channel and the lipid bilayer (Orr et al., 2006). In other words, preventing the hydrophobic sidechains of the transmembrane regions of the channels from being exposed to the aqueous environment would be energetically favorable. Thus, opening the channel in the thinner bilayer lowers the free energy through the hydrophobic effect (Orr et al., 2006). Abnormalities in mechanosensitive channel function can result in neuronal, muscular, cardiac, and kidney disturbances (Chen et al., 1999; Driscoll and Chalfie, 1991; Franco Jr. and Lansman, 1990; Hansen et al., 1990). In autosomal dominant polycystic kidney disease (PKD), the defect may be due to abnormal Ca+2 signaling through polycytins, cation channels that act as mechanosensory channels (Corey, 2003). Polycystins (PKD) 1 and 2 are found in primary cilia of renal epithelial cells (Delmas, 2004), at cell–cell junctions, and a number of other cell types. Together they form a complex in which PKD1 is a regulatory/anchoring subunit and PKD2 is a mechanically regulated calcium channel (Orr et al., 2006). Loss of this signaling complex, through loss of either PKD1 or PKD2, results in the loss of cilial mechanotransduction and Ca+2 signaling. Under normal conditions, urine flow triggers signals through the cilia that modulate kidney tubule growth. However, when this signal is lost, the result is cyst formation in the kidney (Delmas, 2004; Orr et al., 2006). Loss of PKD1/2 also leads to cardiovascular and skeletal defects, as well as left-right asymmetry abnormalities (Orr et al., 2006). Mechanotransduction has also been shown to be important in modulating blood pressure changes via the myogenic response, a response that regulates rapid changes in blood vesicles and provides a mechanism for protecting capillary beds from acute blood pressure changes (Orr et al., 2006). This effect is mediated though calcium-signaling pathways (Davis et al., 2001) that are activated when nonspecific cation channels open in the stretch-sensitive smooth muscle cells. Depolarization of the membrane occurs and this activates calcium entry L-type calcium channels that regulate myogenic constriction (Orr et al., 2006). The stretch-sensitive component of arteries is mediated by integrins primarily through the action of focal adhesion kinase and MAP kinases. Mechanotransduction also plays an important role in normal lung function. Mechanical forces are constantly being applied to the lung epithelium and these forces activate the synthesis and secretion of surfactant proteins by type II epithelial cells (Gutierrez et al., 2003; Wirtz and Dobbs, 2000). Surfactant proteins

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are necessary for normal lung function because they lower the surface tension at the air-liquid interface. Further, a mechanotransducing mechanism has been proposed to explain c-Src activation resulting from strain on the actin cytoskeletal (Han et al., 2004). AFAP-110, an actin filament-associated protein found in lung epithelia, binds and activates c-Src, bridging changes between the actin cytoskeleton and c-Src activation (Han et al., 2004). Another mechanism to explain mechanotransduction involves growth factor shedding (Tschumperlin et al., 2004). For example, EGFR is activated in epithelial type II cells in response to cellular contraction, leading to the activation of ERK MAP kinases (see below), and the production of surfactant proteins (Correa-Meyer et al., 2002; Sanchez-Esteban et al., 2004). ERK1/2 activation requires the shedding of heparin binding-EGF into the extracellular space. Given that increasing pressure decreases the volume of the intercellular space, the heparin binding-EGF concentration increases even if the amount of shedding does not change, resulting in EGFR activation (Orr et al., 2006; Tschumperlin et al., 2004). The preceding examples illustrate that mechanotransduction is utilized by a vast array of cell types, many of which were not discussed. Two different examples of mechanotransduction were presented. In the first, membrane tension changes promote protein unfolding, and result in activation pressure-sensitive channel gating. In the second case, external forces affect large molecular complexes by either altering the individual components relative to each other or by altering the complex relative to the actin cytoskeleton or to the extracellular matrix. PKD provides an excellent example of how loss of mechanotransduction leads to dire consequences. 2. MAP Kinase The mitogen-activated protein kinase (MAPK) signaling pathways are a family of signaling cascades that are affected by receptor-ligand interactions as well as oxidative stress (reviewed in McCubrey et al., 2006). There are five families of MAPK signaling pathways (Widmann et al., 1999), although only four of them are activated by oxidative stress (McCubrey et al., 2006). These include ERK1/2 (extracellular regulated kinases), ERK3/4, JNK (Jun N-terminal kinases), p38 kinase, and the BMK1 (big mitogen-activated protein kinase 1; also known as ERK 5) signaling pathways (McCubrey et al., 2006). The JNK and p38 pathways are often grouped together as the stress-activated kinases (McCubrey et al., 2006). Oxidative stress can either activate (McCubrey et al., 2006) or inhibit (Cross and Templeton, 2004) these pathways, with the outcome depending on cell type and magnitude of the oxidative stress. MAP kinases regulate a number of cellular processes including cell growth and differentiation, gene expression, cell survival and apoptosis (Lu and Xu, 2006). In the next section, we will first review the MAPK signaling pathways and then discuss how oxidative stress affects these pathways.

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FIGURE 2 The MAP kinase cascade and MAP kinases activated during oxidative stress. Based on McCubrey et al. (2006).

General Features of the MAP Kinase Pathways The first MAPK signaling pathway was identified based on the ability of ERK (extracellular regulated kinases) to phosphorylate microtubule-associated protein. When it became clear that there were a large number of substrates and kinases and pathways, the name became more generally derived from the ability of these kinases to be activated by mitogens (mitogen-activated protein (MAP) kinases) (McCubrey et al., 2006). Five families of MAP kinase signaling pathways have been identified to date (Figure 2). The ERK, JNK (Jun N-terminal kinases), p38, and BMK1 pathways share two common features. First, they are all serine/threonine kinases that preferentially phosphorylate substrates with a critical Pro residue in the recognition site. Second, they all operate in a cascade in which a MAP kinase kinase kinase (MAP3K) phosphorylates and activates a MAP ki-

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nase kinase (MAP2K), which phosphorylates and activates a MAPK (McCubrey et al., 2006). In mammalian cells, 12 MAP kinases, 7 MAP2Ks, and 20 MAP3Ks have been identified (Lu and Xu, 2006). Each of the MAPK families can be activated by at least two MAP2Ks and by multiple MAP3Ks (Lu and Xu, 2006), illustrating the complexity and overlap of these pathways. In the following sections, a description of each of these pathways will be detailed, with a specific focus on how each is affected by oxidative stress. ERK1/2 The prototype of the MAPK pathway, ERK, is activated following receptor tyrosine kinase activation in a cascade shown in Figure 2. ERK1 and ERK2 are more than 80% identical and are expressed in all tissues (Lu and Xu, 2006). Another ERK, ERK3/4 is not activated by oxidative stress (McCubrey et al., 2006), and therefore will not be discussed further. ERK1/2 (a MAPK) is activated by MEK1/2 (MAP/ERK Kinase (MAP2K)) (McCubrey et al., 2006), which is activated by Raf (MAP3K). A number of pathways can lead to MEK1/2 activation (Figure 2), including growth factor/hormone stimulation, stress (including oxidative stress), or cytokine activation (Lu and Xu, 2006). Activation of protein kinase C either through increases in intracellular Ca+2 or by PLCγ activity results in Ras activation, leading to c-Raf /MEK/ERK activation. The RAS/RAF/MEK/ERK pathway is likely the most-studied MAPK pathway (Figure 2). ERK activation, however, is also known to occur via RAS-independent pathways as well (Burgering et al., 1993). MAP kinases are activated by a dual phosphorylation event on a conserved Thr-Xaa-Tyr motif in their activation loop by a MAP2K (Lu and Xu, 2006). MAP2Ks show remarkable specificity, whereas MAP3K can activate multiple MAP kinase cascades (Lu and Xu, 2006). As would be expected, Raf is not the only MAP3K that regulates ERK1/2. Other MAP3Ks that include Mos, TPL2 protooncogene, MLK-like mitogen-activated protein triple kinase (MLTK), and interleukin 1 receptor-associated kinase (IRAK) have been shown to activate ERK1/2 (Gotoh et al., 2001; Gotoh and Nishida, 1995; Lu and Xu, 2006; MacGillivray et al., 2000; Salmeron et al., 1996), demonstrating that there are a number of potential start points for this cascade. More than 150 substrates have been identified for ERK1/2, with a list including transcription factors, kinases, phosphatases, cytoskeletal proteins, receptors, signaling molecules, and apoptosis-related proteins (McCubrey et al., 2006). Many of the down-stream effects appear related to cell survival. Constitutively active form of RAS supports cellular transformation (Cuadrado et al., 1993; Hoyle et al., 2000), and activated forms of RAF kinases inhibit apoptosis (Hoyle et al., 2000). Further, kinases downstream of ERK have similar anti-apoptotic effects, lending credence to the idea that this pathway facilitates cell survival. ERK1/2 activate a number of anti-apoptotic proteins (Mcl-1, Bcl-XL, IEX-1, c-Flip, CREB, and CBP) and DNA repair proteins (ERCC1, XRCC1, and ATR),

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while at the same time inhibit a number of pro-apoptotic proteins (caspase 8, caspase 9, Bim, BAD, Hid, and STAT3/5 (reviewed in Lu and Xu, 2006). JNK In the JNK (Jun-N-terminal kinase) pathway, the TNF family receptors and/or oxidative stress starts a cascade that eventually activates MKK4/7 (MAP2K 4 and 7) that phosphorylate and activate JNK on critical threonine and tyrosine residues (Davis, 1999). MKK4/7 is activated by Ras/Rac/MEKK1/4 from oxidative stress or through the TNF family of receptors and ASK1 (apoptosis signalregulating kinase 1) activation (McCubrey et al., 2006). Both tumor necrosis factor and FAS receptors are known to activate the JNK pathway. This pathway is also activated by a number of types of cellular stress including UV light, heavy metals, and reactive oxygen intermediates, with downstream targets that include Jun, ATF-2, Elk2, and NF-E2-related factor-2 (Nrf2) (reviewed in McCubrey et al., 2006). P38 P38 MAPK is activated by MKK3 or MKK6 (MAP2Ks), which are activated by MLK3, which is in turn activated by Rac1 and cdc42. Both growth factor receptors and the TNF family of receptors activate this pathway, illustrating the overlap between the P38 and JNK pathways (McCubrey et al., 2006). TNF receptors activate this pathway via cdc42, whereas growth factor receptors activate this pathway through Ras. Targets of the p38 pathway include a number of transcription factors such as MEF2, ATF-2, Elk-1 and CREB (McCubrey et al., 2006). A unique feature of this pathway is that it does not induce an antioxidant response via Nrf2 phosphorylation (McCubrey et al., 2006), which explains its ability to promote rather than inhibit apoptosis. The balance between ERK and P38 signaling, therefore, appears to be a critical factor regulating cell survival versus apoptosis (Birkenkamp et al., 1999). BMK1 This is the most recently described MAPK pathway and therefore the least characterized. It is also referred to as the ERK5 pathway and is activated by oxidative stress, G-coupled protein receptors, and growth factor receptors such as EGF. The MAP2K in the BMK1 pathway is MEK5 and the MAP3Ks are MEKK2 and MEKK3. Activation of this pathway promotes proliferation, differentiation and cell survival, and the downstream targets include Mef2C, c-Myc, and p90Rsk (McCubrey et al., 2006). Oxidative Stress and the MAPK Signaling Pathways Oxidative stress comes from a number of sources including normal cellular respiration, activation of growth factor receptors, and activation of the tumor

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necrosis factor (TNF) family of receptors. Cellular stress results from the production of free radicals and reactive oxygen species (ROS), and these molecules have harmful effects on both cell membranes and cellular functions. To deal with oxidative insults, cells have developed a number of methods to protect themselves that include glutathione and thioredoxin buffering systems and catalase (McCubrey et al., 2006). Transcriptional responses to stress occur through upregulation of antioxidant genes under the control of the antioxidant response element promoter and the action of the transcription factor Nrf2 (Cho et al., 2006). Part of the cell’s response to oxidative stress includes activation of the MAPK signaling pathways. The nature of the response depends upon the level of stress and which pathways become activated. Under moderate stress conditions, the MAPK pathways promote cell viability, however, when overwhelmed, apoptosis occurs. The consequences of oxidative stress on each of the MAPK signaling pathways are described below. Oxidative stress activates the ERK1/2 signaling pathway through activation of MEK1 and 2, ERF and PDGF receptors, and certain Src kinases (McCubrey et al., 2006). Further, hydrogen peroxide treatment can activate Ca+2 channels, leading to PKC activation and MEK1/2 activation. NO can activate Ras, again leading to MEK1/2 activation, illustrating that reactive oxygen species have a number of potential targets in cells that activate the ERK1/2 pathway (see Figure 3). Although this pathway is known to provide protection against oxidative stress, it is clear that too much stress overwhelms the system. In the JNK kinase-signaling pathway, oxidative stress acts on three components of the pathway (Figure 3). In the first, ASK1 (apoptosis signal-regulating kinase), the upstream component for MKK4/7, is regulated by its direct interaction with thioredoxin. When ASK1 is bound to thioredoxin, it is inactive. When thioredoxin is oxidized, it dissociates from ASK1, allowing ASK1 to become activated (McCubrey et al., 2006). Oxidative stress can also active Rac or the TNF receptor family directly, both of which result in JNK activation. The consequences of oxidative stress on the JNK pathway depend on the kinetics of activation. Sustained activation promotes apoptosis, whereas lower levels of reactive oxygen intermediates usually do not (McCubrey et al., 2006). Hydrogen peroxide, NO, and peroxynitrite all activate the p38 pathway by activating a number of targets that include Rac, MEK1–4, MLK3, ASK1 and the TNF receptor family (McCubrey et al., 2006). NO directly increases RAS activity and this activates the p38 pathway. Since this pathway shares common components with the JNK pathway, many of the mechanisms that active JNK also activate p38. Further, the kinetics of the activation, much like the JNK pathway, influences the fate of the cell after the activation cascade. In the BMK1 pathway, oxidative stress indirectly affects MEKK3 activity, leading to the downstream effects on MEK5 and BMK1 (ERK5). The consequences of activation of this pathway and the details of the pathway are less clear than the other signaling pathways.

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FIGURE 3 Activation of BMK1, JNK, p38 and ERK1/2 kinases by oxidative stress through different pathways and their effects on cellular processes. Based on McCubrey et al. (2006).

Reactive oxygen species can therefore be viewed as signaling molecules since they active a number of components in the four MAPK pathways outlined above. The cellular responses are often mediated by the transcription factors that are activated, and for the most part, result in proliferation or cell survival. If, on the other hand, the oxidative stress is prolonged or more severe, the consequences are pro-apoptotic, probably because significant cellular damage from the oxidative

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reactions has already occurred. Understanding the balance between cell survival and apoptosis will require a complete characterization of these complicated, overlapping MAPK pathways.

III. CALCIUM SIGNALING Reactive oxygen and nitrogen species have a number of effects on normal cellular functions, but when present in excess, trigger recovery responses, or in the worst case, apoptosis. One of the first effects of oxidative stress is to activate calcium channels in the plasma membrane and ER, facilitating calcium influx (reviewed in Ermak and Davies, 2002). Under normal conditions, cytoplasmic calcium levels are tightly maintained at about 100 nM, whereas outside the cell the concentration is high (∼mM). When a signal opens calcium channels, intracellular concentrations increase 20-fold and trigger calcium-activated cellular responses. Under normal conditions, the intracellular concentration in the cell is kept low by calcium pumps at the plasma membrane and ER, by Na+ -driven Ca+2 exchangers, by Ca+2 -binding proteins in the cytoplasm, and by active Ca+2 import into mitochondria. In the later case, mitochondria use a low-affinity, high capacity Ca+2 pump in the inner membrane that utilizes the electrochemical gradient generated by oxidative phosphorylation. ER calcium release is mediated by inositol 1,4,5-triphosphosphate (IP3 ) through IP3 -gated calcium channels. IP3 is generated through the action of G-protein-linked receptors that activate phospholipase C-β, which leaves the plasma membrane and diffuses to the ER to activate calcium channels. The wave of calcium also mediates PKC translocation from the cytosol to the plasma membrane, where it is activated by calcium, by the inner leaflet of the plasma membrane, and by diacylglycerol, the other product of phospholipase C-β. These calcium fluxes and PKC activation lead directly to activation of the ERK1/2 pathway (Ca+2 -PKC-Ras-c-Raf-MEK1/2-ERK1/2). Calcium fluxes can either be highly localized or can occur throughout the cell, providing a global effect. The pulses are usually short-lived since longlived elevations in intracellular calcium are often lethal. Some of the released ER calcium is taken up by mitochondria, which act as a temporary store before the calcium is returned to the ER (Berridge et al., 1998). Temporary calcium storage is fine, but when the ER becomes calcium depleted when calcium reuptake is blocked pharmacologically, for example, there are two cellular responses. First, loss of ER calcium leads to the induction of ER stress signals. And second, the subsequent buildup of calcium of mitochondrial calcium leads to mitochondrial dysfunction and eventually apoptosis (Dolmetsch et al., 1998; Berridge et al., 1998). This suggests that any perturbation with mitochondrial function leads to dire consequences.

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IV. OXIDATIVE STRESS AND ORGANELLE DYSFUNCTION Oxidative stress leads to a number of changes in cell membranes and organelle function. The best-characterized examples include the effects on the ER, peroxisomes, mitochondria, and lysosomes. Oxidative stress results from the buildup of reactive oxygen species (ROS) and reactive nitrogen species (RNS). High levels of ROS damage DNA, proteins and lipids and often lead to the dysregulation of signaling pathways and pathological consequences (reviewed in Schrader and Fahimi, 2006). Oxidative stress is associated with a number of diseases including atherosclerosis, cystic fibrosis, cancer, type-2 diabetes, and a number of neurodegenerative diseases such as Parkinson’s and Alzheimer’s disease. Increased oxidative stress has also been closely associated with aging (Schrader and Fahimi, 2006; Terlecky et al., 2006), suggesting that this is a normal consequence over time, and exacerbation of this process progresses to a disease state.

A. Oxidative Stress and the ER The first response to oxidative stress probably occurs in the endoplasmic reticulum. And that is because the ER has a number of signaling pathways that are based on the oxidation status of specific proteins (reviewed in Gorlach et al., 2006). Oxidative stress from ROS causes calcium release, leading to mitochondrial calcium loading, which leads to more ROS production and more calcium release. The sarco(endo)plasmic reticulum calcium ATPases (SERCAs) which mediate calcium reuptake into the ER, are inhibited by oxidation. Inhibition of calcium reuptake inhibits protein synthesis and processing and leads to the accumulation of partially folded proteins (Gorlach et al., 2006). When left unchecked, this can lead to the unfolded protein response that either facilitates cell recovery mechanisms, or promotes apoptosis, the result depending on the magnitude and duration of the unfolded protein response.

B. Oxidative Stress and Peroxisomes ROS include the superoxide anion and hydrogen peroxide, while RNS include nitric oxide. In the respiratory pathway in peroxisomes, electrons are removed from metabolites to reduce oxygen to hydrogen peroxide. Nitric oxide is also produced in peroxisomes (Stolz et al., 2002), indicating that peroxisomes play an important role in ROS/RNS production in the cell (Schrader and Fahimi, 2006). β-oxidation of fatty acids in peroxisomes is the major pathway for the production of hydrogen peroxide. Catabolism of hydrogen peroxide occurs via a number of antioxidant enzymes including catalase. Since hydrogen peroxide is membrane permeable, antioxidant enzymes that degrade it

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can also be found in the cytoplasm, nucleus, and mitochondria (Schrader and Fahimi, 2006). Problems arise when the β-oxidation enzymes are dramatically increased when activators of the peroxisome proliferators activated receptors (PPARs) such as free fatty acids overload the system. This leads to oxidative stress, and when left unchecked, cancer (Schrader and Fahimi, 2006). In other cases, transition metals such as ferrous iron, which are normally found in peroxisomes in complexed form, can be released to generate the hydroxyl radical, which causes serious lipid peroxidation, damage to the peroxisomal membrane, and eventually loss of peroxisomal function (Schrader and Fahimi, 2006; Yokota et al., 2001). Peroxisomes not only participate in the generation of ROS, but also have scavenging activities to metabolize these reactive oxygen molecules. When PPARs become activated, the expression of genes involved in lipid β-oxidation is strongly enhanced (up to 10-fold), while the induction of hydrogen peroxide scavenging genes such as catalase is not (1- to 2-fold) (Rao and Reddy, 1987). Since a number of xenobiotics induce PPARs including hypolipidemic drugs, industrial chemicals, agrochemicals, and many environmental pollutants (Beier and Fahimi, 1991), it has been proposed that the tumor-promoting properties of these compounds is caused by the disproportionate production of hydrogen peroxidegenerating enzymes over scavenging enzymes, leading to oxidative stress and eventually leading to hepatic tumors in animals (Schrader and Fahimi, 2006). An excellent review on peroxisomes and their central role as generators of oxidative stress can be found in Schrader and Fahimi (2006). Although peroxisomes are important in generating reactive oxygen species for metabolic purposes, they do not generate as much as mitochondria (Beckman and Ames, 1998; Lee and Wei, 2001). However, in peroxisomes when metal ion-catalyzed conversion of hydrogen peroxide to the hydroxide radical occurs, this can result in damage to mitochondria, promoting an escalation in ROS production (Terlecky et al., 2006), and a vicious cycle that leads to apoptosis. Peroxisome function is normally kept in check with catalase maintaining the delicate balance between hydrogen peroxide formation and degradation. When this balance is upset, there is an accumulation of hydrogen peroxide and downstream reactive oxygen species (Terlecky et al., 2006). When catalase activity is compromised, as in certain disease states or after exposure to certain xenobiotics, or even during the normal aging process, there is an increase in ROS that leads to oxidative damage to cellular constituents. That this is a part of the normal aging processing should not be considered surprising. In fact, there are new potential therapies that involve increasing catalase levels that may provide a benefit in the different aging pathologies associated with oxidant-induced cellular damage (Terlecky et al., 2006).

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C. Oxidative Stress and Mitochondria When peroxisomal balance is compromised, mitochondrial oxidative damage starts as well. This is a critical problem because mitochondrial deficiencies directly influence a number of diseases including Parkinson’s disease, diabetes mellitus, and possibly Alzheimer’s disease (Kakkar and Singh, 2007). The cell has antioxidants inside and outside mitochondria to detoxify ROS. Inside mitochondria, the hydroperoxyl radical is converted to hydrogen peroxide by matrix manganese superoxide dismutase, and the superoxide anion is partially dismutated by CuZn superoxide dismutase (reviewed in Kakkar and Singh, 2007). When left unchecked, ROS, especially very reactive radicals, attack proteins, lipids, and mitochondrial DNA. Outside mitochondria, two systems regulate the intracellular redox environment, glutathione-S transferase and thioredoxin. Since the reduced glutathione concentration is so high, it forms the strongest protection against oxidative stress within the cell. If, however, the reduced glutathione levels become depleted, then the cell is more sensitive to apoptotic stimuli (Kakkar and Singh, 2007). Thioredoxin is a thiol-specific antioxidant that is concentrated in the ER and functions to reduce disulfide bridges of proteins that have been subjected to oxidative stress. Both glutathione-S transferase and thioredoxin are required for maintaining the proper redox environment for proteins and act as thiol-specific antioxidants. When the redox system is overwhelmed, ROS damages a number of macromolecules, including cardiolipin, a mitochondrial specific lipid. This is a critical component of mitochondrial energetics since this lipid is present on the inner mitochondrial membrane and is important for the activities of the adenine nucleotide transporter and cytochrome c oxidase (Hoch, 1992; Paradies et al., 1998). Further, it has been proposed that mitochondrial DNA may be more sensitive to oxidative damage because (1) it lacks DNA associated proteins like histones; (2) several of the proteins encoded by mitochondrial are essential for the electron transport chain; (3) its DNA is close to where ROS production occurs; and (4) it is present in many thousand copies per cell (Kakkar and Singh, 2007). Mitochondrial DNA damage occurs over a lifetime, resulting in a decline in mitochondrial function, and increased ROS production. At the extreme end, overwhelming ROS production leads to release of cytochrome c and activation of apoptosis. Overproduction of ROS also mediates the oligomerization of the proapoptotic protein Bax through the introduction of intrachain disulfide bonds. Bax oligomerization is necessary for Bax insertion into the outer mitochondrial membrane, which leads to the release of cytochrome c and other proapoptotic proteins and the initiation of the apoptotic cascade (Er et al., 2006). Cytochrome c is anchored to the inner mitochondrial membrane by its affinity to cardiolipin, and this interaction is disrupted by cardiolipid oxidation by ROS (Neuzil et al., 2006). ROS, therefore, is a central mediator in several of the steps leading to cell death.

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D. Oxidative Stress and Lysosomes Many of the diseases that have been associated with ROS over-production involve the CNS, probably because of the brain’s particular susceptibility to oxidative damage (reviewed in Butler and Bahr, 2006). An interesting feature of a number of CNS diseases including Alzheimer’s disease is lysosomal activation (Butler and Bahr, 2006). Lysosomal activation is a normal consequence of in the CNS during aging, and oxidative damage appears to have profound effects on this organelle. There is an important link between lysosomal function and mitochondrial function since the accumulation of oxidized material within lysosomes during oxidative stress reduces autophagic processes, including the autophagic removal and recycling of damaged mitochondria (Brunk and Terman, 2002). Oxidative damage is a critical component of Alzheimer’s disease since the amyloidogenic peptides derived from the amyloid precursor protein form free radical species (Ditaranto et al., 2001; Goodman and Mattson, 1994; Hensley et al., 1994) and produce membrane oxidative stress (Cutler et al., 2004; Ditaranto et al., 2001). Further, oxidative damage to lysosomes by free radicals not only affects autophagy, but also disruption of the lysosomal membrane integrity results in lysosomal enzyme release which includes a number of cathepsins which are known to promote ROS production and apoptosis through their effects on mitochondria (Butler and Bahr, 2006). The data indicate that most of the harmful effects of oxidative damage to lysosomes results in mitochondrial damage, with an amplification of ROS production in a vicious cycle of cellular organelle damage that eventually leads to apoptosis.

E. Summary and Conclusions Oxidative stress is a process results from an inbalance between production and catabolism of reactive oxygen species. The cell has a number of mechanisms to protect against oxidative stress since it damages cell membranes, activates signaling cascades, and promotes mitochondrial DNA damage, along with organelle dysfunction. Many of the consequences, particularly fluxes in cytosolic calcium, oxidation of lipids, disruption of ER and lysosome function lead to mitochondrial damage. Interestingly, slow loss of mitochondrial function due to oxidative damage appears to be a component of the normal aging process. Severe insults to the cell, however, result in activation of the apoptotic cascade, probably because the oxidative damage is too pronounced for the cell’s machinery to facilitate recovery. Clearly understanding this process in more detail will allow for treatments of a number of neurological diseases, and interestingly, may also be important in understanding the normal aging process. Future therapies will be directly involved with the delicate balance between ROS production and ROS catabolism.

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Neuzil, J., Wang, X.F., Dong, L.F., Low, P., Ralph, S.J. (2006). Molecular mechanism of ‘mitocan’induced apoptosis in cancer cells epitomizes the multiple roles of reactive oxygen species and Bcl-2 family proteins. FEBS Lett. 580, 5125–5129. Orr, A.W., Helmke, B.P., Blackman, B.R., Schwartz, M.A. (2006). Mechanisms of mechanotransduction. Developmental Cell 10, 11–20. Paradies, G., Ruggiero, F.M., Petrosillo, G., Quagliariello, E. (1998). Peroxidative damage to cardiac mitochondria, cytochrome oxidase and cardiolipin alterations. FEBS Lett. 424, 155–158. Pieczenik, S.R., Neustadt, J. (2007). Mitochondrial dysfunction and molecular pathways of disease. Exp. Mol. Pathol. 83, 84–92. Rao, M.S., Reddy, J.K. (1987). Peroxisome proliferation and hepatocarcinogenesis. Carcinogenesis 8, 631–636. Sachs, F. (1988). Mechanical transduction in biological systems. Crit. Rev. Biomed. Eng. 16, 141–169. Sachs, F., Morris, C.E. (1998). Mechanosensitive ion channels in nonspecialized cells. Rev. Physiol. Biochem. Pharmacol. 132, 1–77. Salmeron, A., Ahmad, T.B., Carlile, G.W., Pappin, D., Narsimhan, R.P., Ley, S.C. (1996). Activation of MEK-1 and SEK-1 by Tpl-2 proto-oncoprotein, a novel MAP kinase kinase kinase. EMBO J. 15, 817–826. Sanchez-Esteban, J., Wang, Y., Gruppuso, P.A., Rubin, L.P. (2004). Mechanical stretch induces fetal type II cell differentiation via an epidermal growth factor receptor-extracellular-regulated protein kinase signaling pathway. Am. J. Resp. Cell Mol. Biol. 30, 76–83. Schrader, M., Fahimi, H.D. (2006). Peroxisomes and oxidative stress. Biochim. Biophys. Acta 1763, 1755–1766. Simons, K., Toomre, D. (2000). Lipid rafts and signal transduction. Nature Rev. 1, 31–39. Singer, S.J., Nicolson, G.L. (1972). The fluid mosaic model of the structure of cell membranes. Science 175, 720–731. Stolz, D.B., Zamora, R., Vodovotz, Y., Loughran, P.A., Billiar, T.R., Kim, Y.M., Simmons, R.L., Watkins, S.C. (2002). Peroxisomal localization of inducible nitric oxide synthase in hepatocytes. Hepatology 36, 81–93. Baltimore, MD. Terlecky, S.R., Koepke, J.I., Walton, P.A. (2006). Peroxisomes and aging. Biochim. Biophys. Acta 1763, 1749–1754. Tschumperlin, D.J., Dai, G., Maly, I.V., Kikuchi, T., Laiho, L.H., McVittie, A.K., Haley, K.J., Lilly, C.M., So, P.T., Lauffenburger, D.A., Kamm, R.D., Drazen, J.M. (2004). Mechanotransduction through growth-factor shedding into the extracellular space. Nature 429, 83–86. Vereb, G., Szollosi, J., Matko, J., Nagy, P., Farkas, T., Vigh, L., Matyus, L., Waldmann, T.A., Damjanovich, S. (2003). Dynamic, yet structured: The cell membrane three decades after the Singer– Nicolson model. Proc. Natl. Acad. Sci. USA 100, 8053–8058. Widmann, C., Gibson, S., Jarpe, M.B., Johnson, G.L. (1999). Mitogen-activated protein kinase, conservation of a three-kinase module from yeast to human. Physiol. Rev. 79, 143–180. Wirtz, H.R., Dobbs, L.G. (2000). The effects of mechanical forces on lung functions. Resp. Physiol. 119, 1–17. Yokota, S., Oda, T., Fahimi, H.D. (2001). The role of 15-lipoxygenase in disruption of the peroxisomal membrane and in programmed degradation of peroxisomes in normal rat liver. J. Histochem. Cytochem. 49, 613–622.

CHAPTER 2 The Interaction of Reactive Oxygen and Nitrogen Species with Membranes Matías N. Möller∗ , Jack R. Lancaster Jr.† and Ana Denicola∗ ∗ Lab. Fisicoquímica Biológica, Facultad de Ciencias, Universidad de la Republica, Igua 4225, 11400 Montevideo, Uruguay † Departments of Anesthesiology, Physiology & Biophysics, and Environmental Health Sciences and Center for Free Radical Biology, University of Alabama at Birmingham, AL, USA

I. Reactive Oxygen and Nitrogen Species II. Physical Interactions: Compartmentalizing Reactivity A. Nitric Oxide and Oxygen B. Superoxide C. Hydrogen Peroxide D. Hydroxyl Radical E. Peroxynitrite and Nitrogen Dioxide F. Carbonate Radical G. Permeability of Ionic Species and Their Conjugate Acids H. Solubility of ·NO and O2 in Membranes I. Diffusion of ·NO and O2 in Membranes III. Chemical Effects: Lipid Peroxidation A. Nitric Oxide B. Nitrogen Dioxide C. Peroxynitrite and Carbonate Radical D. Superoxide E. Hydrogen Peroxide and Hydroxyl Radical F. Biophysical Effects of Lipid Peroxidation in Membranes References

I. REACTIVE OXYGEN AND NITROGEN SPECIES The term reactive oxygen–nitrogen species (RONS) is used to group a number of molecules derived from and including molecular oxygen and nitric oxide (·NO). Most RONS are oxidizing species, free radicals (like O2 , ·NO) but some are still reactive although not radicals (like hydrogen peroxide or peroxynitrite). Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00202-0

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FIGURE 1 Reaction network of oxygen and nitrogen reactive species. It can be appreciated how RNS arise from the interaction of ·NO with ROS, either by the autoxidation of ·NO, or by the reaction of ·NO with superoxide to yield peroxynitrite. The formation of the conjugate acids of superoxide and peroxynitrite has been included because it is the neutral form of the molecule that can cross the biomembrane by simple diffusion. Furthermore, the formation of radical species from peroxynitrous acid or from nitrosoperoxycarboxylate (ONOOCO− 2 ) may participate in membrane permeation and oxidation. The metal-dependent reduction of H2 O2 (Fenton reaction) generates hydroxyl radical, a good initiator of membrane lipoperoxidation.

They are reactive towards several biomolecules, most of the times oxidizing, damaging and leading to deleterious effects on their function, although some important exceptions exist and some RONS are endogenously produced with specific physiological roles. The final effect of RONS in a biological environment will greatly depend on their interactions with biomembranes, which is the focus of the present chapter. But first, let’s overview some of the main RONS formed in vivo (Figure 1): Nitric oxide is an important intercellular messenger produced enzymatically from the oxidation of L-arginine by nitric oxide synthases (NOS) (Stuehr et al., 2004). It is involved in vasodilation, neurotransmission, and also in immune response (Beckman and Koppenol, 1996). It switches from cell signaling to cell damaging by significantly increasing the amount of ·NO produced and reacting with other molecules to form more reactive species (Beckman and Koppenol, 1996). It can react with superoxide (O·− 2 ) at diffusion-limited rates to form the more oxidizing species peroxynitrite (Radi et al., 2000). Peroxynitrite (ONOO− ) is an oxidizing species with a complex chemistry (Radi et al., 2000), that is illustrated in Figure 1. It is relevant to note its pKa = 6.8 close to physiological pH,

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25

its ability to form radicals nitrogen dioxide (·NO2 ) and hydroxyl radical (·OH) in a 30% yield or rearrange to nitrate (70%), and its reactivity with CO2 to also form ·NO and carbonate radical (CO·− ) in a 35% yield. 2 3 Alternatively, ·NO can react with O2 , to yield primarily ·NO2 and then dinitrogen trioxide (N2 O3 ), important nitrating and nitrosating species, in a reaction that is accelerated by lipid membranes (Liu et al., 1998; Möller et al., 2007a, 2007b). There are several sources of superoxide, which include NADPH oxidase, xanthine oxidase, mitochondrial electron leakage and xenobiotic metabolism (Radi et al., 2000). The superoxide anion can form its conjugate acid, the perhydroxyl radical (HOO·) with a pKa = 4.8 and it can also be reduced to hydrogen peroxide (H2 O2 ) (Green and Hill, 1984). The antioxidant enzyme responsible for O·− 2 detoxification is superoxide dismutase (SOD), a metal-dependent enzyme that catalyzes O·− 2 disproportionation to O2 and H2 O2 (Brunori and Rotilio, 1984). Hydrogen peroxide is relatively stable, but reacts with reduced metals via Fenton reaction to yield the most oxidizing species: hydroxyl radical (·OH) (Buettner, 1993). Because of this and its role in cell signaling, there are additional antioxidant enzymes to deal with H2 O2 , including metal-dependent (catalase), selenium-dependent (glutathione peroxidase), and thiol-dependent peroxidases (peroxiredoxins) (Rhee et al., 2005). All the RONS interact with biomembranes, either by directly reacting with membrane components and modifying its structure and function, or just crossing them to react with targets on the other side of the membrane. In the next section we will be mainly concerned with the ability of these different species to permeate through the lipid membrane, and then deal with chemical effects on membrane components. The chemical effects will be focused on lipids, the common building blocks of biomembranes, since the effect of RONS on membrane proteins will depend on the particular membrane studied, and will be dealt with in subsequent chapters.

II. PHYSICAL INTERACTIONS: COMPARTMENTALIZING REACTIVITY One of the main functions of biomembranes is to delimit and separate cell and organelles components and functions. In a similar way, the site of action of some RONS can be restricted by biomembranes (additional chemical reactions can also limit the site of action). Therefore, the reactivity of some RONS can be compartmentalized, according to the ability of RONS to traverse a biomembrane. As a general rule, uncharged nonelectrolytes can diffuse through the lipid part of the membrane, while charged species need either to form its conjugate acid, or need a protein channel to permeate a membrane at significant rates. Another important rule is that the smaller and less polar the molecule, the higher permeability through membranes. The order of permeability across pure lipid membranes is: ·NO, O  ONOOH, ·NO , HOO·, H O  O·− , ONOO− , CO·− (Figure 2). 2 2 2 2 3

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FIGURE 2 Classification of reactive species according to their lipid membrane permeation capacity. Pm /Pw is the ratio between membrane permeability and the permeability through an equivalent layer of water. Lipid membranes are not barriers to the passage of ·NO or O2 , while it offers a moderate barrier to the permeation of hydrophilic polar nonelectrolytes, similar to that of water. The permeability of charged molecules is too low to be significant, and will not occur at significant rates unless protein channels are present.

It should be noted that Overton’s rule, that states that the permeability is directly related to solubility in an organic solvent, is not strictly followed by small molecules (nonelectrolytes with mass <50 Da) which show a higher than expected permeability (Lieb and Stein, 1986; Walter and Gutknecht, 1986). It is not necessary for a molecule to be lipophilic (i.e. to be more soluble in the hydrophobic interior of membranes than in water) to permeate a membrane. Polar non-charged molecules like H2 O can enter and cross the membrane even though their solubility is 1000 times lower in the membrane than in water (Figure 2). However, charged molecules find a much higher barrier to permeation, and can traverse membranes only at very low rates (see Cl− in Figure 2), unless protein channels are present. For comparing the ability of different RONS to permeate lipid membranes, we will use reported permeability coefficients (Pm ) that indicate the speed at which the molecule moves across the membrane (Stein, 1986), and relates the flux of the molecule across the membrane (Jm ) to the difference in concentration at both sides of the membrane (C, Eq. (1)). Pm is determined by the diffusion coefficient of the molecule in the membrane (Dm ), the partition coefficient of the molecule between the membrane and water (KP ), and the thickness of the mem-

2. The Interaction of Reactive Oxygen and Nitrogen Species with Membranes

27

brane (δ, Eq. (2)). Jm = Pm .C

(1)

Pm = Dm .KP /δ

(2)

To compare the barrier effect of membranes to the transport of RONS, we compare Pm values with the calculated permeability through an equally thick layer of water, Pw (Eq. (3)) Pw = Dw /δ

(3)

Table 1 summarizes the reported Pm values for different RONS and the calculated Pm /Pw ratios. A. Nitric Oxide and Oxygen The barrier to ·NO and O2 membrane permeation is very low. It is so low that direct measurements of ·NO or O2 transport across membranes cannot be done, because the aqueous layer at both sides of the membrane offers a higher resistance to ·NO or O2 transport than the membrane itself (see below). However, the permeability through membranes could be determined from the interaction of these molecules with spin- or fluorescent probes located at different depths in the lipid membrane, yielding Pm values 10–100 cm/s. For comparison, the permeability of membranes to H2 O or other polar non-charged species is 3–6 orders of magnitude lower (Pm = 10−2 –10−4 cm/s, see Table 1). Thus, biomembranes are not barriers to ·NO and O2 transport. The differences in the calculated Pm /Pw ratios by different authors (Table 1) arise mainly from considering different experimentally determined Dw . As general considerations, it can be seen that the addition of cholesterol to egg yolk phosphatidylcholine (EYPC) leads to slight decrease in ·NO permeability (Subczynski et al., 1996), and also that the lowest permeability is observed in red blood cell membranes (Denicola et al., 1996), but is only 1/5 that through water. According to the assays using probes at different depths of the membrane bilayer, the main permeation barrier is located in the region of the polar headgroups of the phospholipids or in the water interface, while transport is maximal in the mid-bilayer (Denicola et al., 1996; Subczynski et al., 1996). The permeability of membranes to O2 has been studied in a wide range of conditions, and thus we can generalize some observations. As for ·NO, the main barrier to permeation in the membrane lies in the polar headgroups/water interface region, the permeability decreases when cholesterol is added and increases with temperature (Table 1 and references therein). The plasma membrane from Chinese hamster ovary and human red blood cells exhibit the lowest permeability to O2 (Subczynski et al., 1992; Denicola et al., 1996), but even in these cases the permeability of the membrane is just 1/3 that of water (Table 1).

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TABLE 1 Permeability coefficients of reactive oxygen and nitrogen species across different lipid membranes Molecule Membrane

T (°C) Pm (cm/s) Pm /Pw b

Ref.

·NO ·NO ·NO ·NO

EYPC EYPC:Chol (30% Chol) EYPC RBC (human)

20 20 20 20

73 66 47a 18a

1.28 1.17 0.36 0.2

(Subczynski et al., 1996) (Subczynski et al., 1996) (Denicola et al., 1996) (Denicola et al., 1996)

O2 O2 O2 O2 O2 O2 O2 O2 O2 O2 O2 O2 O2

EYPC RBC (human) EYPC EYPC:Chol (30% Chol) CHO CHO DMPC DMPC DOPC DOPC Lens Lipid POPC:Chol (50% Chol) POPC

20 20 20 20 20 37 18 29 10 30 35 35 35

87a 38a 62 56 21 42 12 125 33 114 51 50 157

0.62 0.42 1.08 0.85 0.34 0.45 0.4 1.8 0.8 1.6 0.96 0.94 3.0

(Denicola et al., 1996) (Denicola et al., 1996) (Subczynski et al., 1996) (Subczynski et al., 1996) (Subczynski et al., 1992) (Subczynski et al., 1992) (Subczynski et al., 1989) (Subczynski et al., 1989) (Subczynski et al., 1989) (Subczynski et al., 1989) (Widomska et al., 2007) (Widomska et al., 2007) (Widomska et al., 2007)

H2 O H2 O

EYPC RBC (human)

25 25

2 × 10−4 ∼10−5 1.2 × 10−3 ∼5 × 10−5

(Gennis, 1989) (Lieb and Stein, 1986)

H 2 O2 H 2 O2 H 2 O2 H 2 O2 H 2 O2 H 2 O2 H 2 O2 H 2 O2 O·− 2 O·− 2 O·− 2 HOO·

RBC (rat) PC12 HUVEC IMR-90 Peroxisome (rat liver) E. coli Chara chorallina Jurkat T-Cell DODAC SBPC EYPC EYPC

37 45 25 23 23

1.2 × 10−2 4 × 10−4 1.6 × 10−3 1.1 × 10−3 3 × 10−3 1.6 × 10−3 3.6 × 10−4 2 × 10−4 10−6 2.1 × 10−6 7.6 × 10−8 4.9 × 10−4

∼5 × 10−4 ∼2 × 10−5 ∼6 × 10−5 ∼4 × 10−5 ∼10−4 ∼6 × 10−5 ∼10−5 ∼10−5 ∼4 × 10−8 ∼10−7 ∼10−9 ∼2 × 10−5

(Mathai and Sitaraman, 1994) (Makino et al., 2004) (Makino et al., 2004) (Makino et al., 2004) (Makino et al., 2004) (Makino et al., 2004) (Makino et al., 2004) (Antunes and Cadenas, 2000) (Gomes et al., 1993) (Takahashi and Asada, 1983) (Gus’kova et al., 1984) (Gus’kova et al., 1984)

ONOOH ONOOH ONOOH ONOOH

DMPC EYPC DMPC DPPC

23 21 21 21

8 × 10−4 1.3 × 10−3 6.3 × 10−4 4 × 10−4

∼3 × 10−5 ∼5 × 10−5 ∼2.5 × 10−5 ∼1.6 × 10−5

(Marla et al., 1997) (Khairutdinov et al., 2000) (Khairutdinov et al., 2000) (Khairutdinov et al., 2000)

Na+

EYPC

25

10−14

∼10−16

(Gennis, 1989)

25

10−11

∼10−13

(Gennis, 1989)

Cl−

EYPC

Abbreviations: PC: phosphatidylcholine; EYPC: egg yolk PC; Chol: cholesterol; RBC: red blood cell membrane; CHO: Chinese hamster ovary plasma membrane; DMPC: dimyristoyl PC; DOPC: dioleoyl PC; POPC: palmitoyloleoyl PC; DODAC: dioctadecyldimethylammonium chloride; SBPC: soy bean PC; DPPC: dipalmitoyl PC. a Calculated from data in the reference, P = (average D m app )/d ; where d is 3.2 and 5.0 nm for EYPC and RBC membrane, respectively. b Except for O and ·NO, the rest of the P /P are estimated values using P = 25 cm/s, corresponding to a m w w 2 typical Dw = 1 × 10−5 cm2 /s and δ = 4 nm.

2. The Interaction of Reactive Oxygen and Nitrogen Species with Membranes

29

Why are there problems measuring the permeability of membranes to ·NO or O2 then, if they are just 1/5 to 1/3 that of on equivalent layer of water? Because there is normally a water layer much thicker than the membrane. For directly measuring permeability through the membrane it is necessary to make the transport across the membrane the rate limiting step. However, even with the most vigorous stirring or turbulent flow, due to hydrodynamic effects, cells (or liposomes) are surrounded by an “unstirred layer” of unmixed water, where transport is limited by diffusion (Liu et al., 1998). Thus, in any experimental condition, ·NO usually spends more time traveling through the aqueous solution than crossing the membrane. This can be illustrated by considering the total “resistance” to permeability (PT−1 ), which is the sum of membrane resistance (Pm−1 ) and the unstirred water layer resistance (Pw−1 ) (Figure 3)1 : PT−1 = Pw−1 + Pm−1

(4)

Note that a 12 nm-thick aqueous layer offers a resistance equal to that imposed by red blood cell membranes to ·NO (using Dw = 2.2 × 10−5 cm2 s−1 ) (Zacharia and Deen, 2005), while unstirred layers are usually thicker than 1 µm (1000 nm!) (Liu et al., 1998).

B. Superoxide Ions face a much higher barrier to membrane permeation than nonelectrolytes. The permeability coefficient for chloride across an EYPC membrane is 10−11 cm/s, several orders of magnitude slower than for O2 or water. To cross the membrane, the ion must lose the water hydration shell before entering the membrane and then move into a nonpolar medium. This process is energetically very unfavorable, so membranes are effective barriers to passive ion transport (Gennis, 1989). In cellular membranes, this is compensated by the presence of protein channels in the membrane that facilitate ion transport. Using anion channels (band 3), superoxide can easily cross the red blood cell membrane, and this was demonstrated by blocking O·− 2 transport with the band 3 inhibitors DIDS and SITS (Lynch and Fridovich, 1978b). The decrease in O·− 2 efflux by band 3 inhibitors indicates that basal (passive) permeability is low. It has been estimated that the permeability through a pure soy bean PC membrane (no ion channels) is 2.1 × 10−6 cm/s (Takahashi and Asada, 1983), similar to the estimated permeability through dioctadecyldimethylammonium chloride membranes (DODAC) in the liquid crystalline state (10−6 cm/s, Table 1, Gomes et al., 1993). This permeability coefficient is still too high for a ionic species like superoxide anion. In fact, Gus’kova et al. showed that O·− 2 permeation was insensitive 1 For interpretation of the references to color in this figure legend, the reader is referred to the web version of this book.

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FIGURE 3 Membrane resistance to ·NO permeation, compared with an aqueous layer of variable thickness. The resistance to ·NO permeation through an aqueous layer Pw−1 was calculated using Eq. (3) and DNO = 2.2 × 109 nm2 /s and increases linearly with layer thickness (red line). The resistance to ·NO permeation through the red blood cell membrane (5.6 × 10−9 s/nm, Table 1) and through EYPC membranes (1.4 × 10−9 s/nm, Table 1) are indicated in green. It can be appreciated that a 12.2 nm thick aqueous layer offers the same resistance than a red blood cell membrane, and an even narrower layer of water is equivalent to an EYPC membrane. Unstirred layers are usually thicker than 1 µm, offering a resistance 100 times greater than the membrane itself.

to transmembrane potential, indicating that the permeating species could not be charged, and should be the conjugated acid HOO· instead (Gus’kova et al., 1984). They estimated a permeability coefficient of 4.9 × 10−4 cm/s for HOO·, while −7 cm/s (Table 1). Pm for O·− 2 ∼10

C. Hydrogen Peroxide Hydrogen peroxide (H2 O2 ) is a hydrophilic molecule (KP between ether and water is 0.07 at 26 °C, Mathai and Sitaraman, 1994) that can cross lipid membranes in a similar manner to water. Most of the reported permeability coefficients in different cell membranes lie between 1 × 10−3 and 2 × 10−4 cm/s, very similar to the values reported for water (Table 1). Therefore, the membrane offers an intermediate barrier to H2 O2 transport. The permeability has been observed to vary according to the lipid composition of the membrane (Mathai and Sitaraman, 1994), and even more, Saccharomyces cerevisiae can regulate its membrane composition to decrease H2 O2 permeation (Branco et al., 2004). It was recently

2. The Interaction of Reactive Oxygen and Nitrogen Species with Membranes

31

TABLE 2 Partition coefficients for ·NO and O2 in different lipid systems O

Lipid system

T (°C)

KPNO

KP 2

Ref.

EYPC DLPC LDL Triton X-100 DMPC DMPC RBC DMPC DMPC DMPC

25 25 25 25 35 20

3.6 ± 0.3 3.6 ± 0.6 3.0 ± 0.4 3.1 ± 0.5

3.2 ± 0.3 3.2 2.6 ± 0.3 2.6 4.1 ± 0.6 1.0 ± 0.5 5±4 1.3 3.1 3.3

(Möller et al., 2005, 2007b) (Möller et al., 2007b) (Möller et al., 2005, 2007b) (Möller et al., 2007b) (Smotkin et al., 1991) (Smotkin et al., 1991) (Power and Stegall, 1970) (Subczynski and Hyde, 1983) (Subczynski and Hyde, 1983) (Subczynski and Hyde, 1983)

20 30 37

Abbreviations: as in Table 1, DLPC: dilauroyl PC.

demonstrated that H2 O2 can also use aquaporins (membrane water channels), which increases its permeation through membranes (Bienert et al., 2007).

D. Hydroxyl Radical This radical can be formed from ONOOH homolysis, or from metal-mediated H2 O2 reduction (Figure 1). It is a small polar nonelectrolyte, so it would be expected to permeate like water. However, ·OH is so reactive that will react with the first molecule it finds, within a few nm of the site of formation (Halliwell and Gutteridge, 2000). Facing a membrane, it will react not only with inner fatty acids, but also with the polar headgroups, which are unreactive towards other RONS. Therefore, ·OH will not traverse but react with the membrane.

E. Peroxynitrite and Nitrogen Dioxide Both peroxynitrite and its conjugate acid, peroxynitrous acid, can permeate biological membranes, but through different routes. Peroxynitrous acid can cross lipid bilayers by simple diffusion (Denicola et al., 1998; Khairutdinov et al., 2000; Marla et al., 1997), with a permeability coefficient similar to water (1.3–0.8 × 10−3 cm/s, Khairutdinov et al., 2000; Marla et al., 1997), while peroxynitrite uses anion channels like the band 3 of red blood cells membrane (Denicola et al., 1998). Although it is hard to differentiate between ONOOH and ·NO2 permeation, Khairutdinov et al. showed that ·NO2 could permeate phospholipids membranes as much as ONOOH (Khairutdinov et al., 2000).

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However, additional experiments suggest that ONOOH is even more permeable than ·NO2 . Using lipophilic tyrosine analogs, or tyrosine containing transmembrane peptides, Kalyanaraman et al. showed that intramembrane tyrosine nitration by peroxynitrite (that homolyzes to ·NO2 ) was more efficient than nitration of tyrosine in solution (Zhang et al., 2001). Moreover, the tyrosine residues located closer to the center of the bilayer resulted more nitrated by peroxynitrite, but when myeloperoxidase-derived ·NO2 was used, the nitration yields increased in the opposite direction, favoring the tyrosine residues closer to the surface (Zhang et al., 2003).

F. Carbonate Radical Peroxynitrite reacts very fast with CO2 to yield an unstable intermediate that decays to ·NO2 and carbonate radical (CO·− 3 ) (Figure 1). Carbonate radical is a more selective oxidant than ·OH, is more bulky and is negatively charged. As already discussed, the charge prevents the molecule from entering into the nonpolar lipid interior, and considerably limits its permeability. Furthermore, the pKa of carbonate radical is close to 0, so it will never form the conjugate acid that may permeate the membrane, as ONOOH or HOO· do (Augusto et al., 2002). In agreement with expected results, carbonate radical is very inefficient in crossing pure phospholipids membranes (Khairutdinov et al., 2000), and also in crossing red blood cell membranes (Romero et al., 1999). In theory, it could cross through anion channels, but its reactions with other molecules may preclude its permeation.

G. Permeability of Ionic Species and Their Conjugate Acids As mentioned above, CO·− 3 will always be ionic because of its low pKa , while ·− − ONOO and O2 can form their conjugate acids because their pKa are closer to the physiological pH. In the protonated form, both ONOOH and HOO· show similar transport properties across membranes, however, at pH 7.4, these species are mostly in the ionic state. The proportion of protonated species relative to the total species can be calculated as: −1    [HA]/ [HA] + [A− ] = 1 + 10(pH-pKa ) (5) So that using the reported pKa (6.8 and 4.8, for peroxynitrite and superoxide, respectively) we can calculate that 20% of peroxynitrite will be present as ONOOH, while only 0.16% of superoxide will be protonated. Therefore, at pH 7.4, mem− branes will restrict more the transport of O·− 2 than that of ONOO (∼100 times − more). This is why it is argued that the sites of ONOO formation are dictated mainly by the sites of O·− 2 formation (Radi et al., 2000).

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H. Solubility of ·NO and O2 in Membranes The solubility of gases in liquids is expressed in many different units, and care should be taken when interpreting solubility data. For convenience, we will use the partition coefficient (KP ), that is the ratio between the molar concentrations of the molecule in the membrane and in the aqueous solution (KP = Cm /Cw ) at equilibrium and at a defined temperature (25 °C, unless specified). For many years it was considered that ·NO solubility in membranes should be higher than in water because of the ∼9 times higher solubility of ·NO in organic solvents compared to water (Shaw and Vosper, 1976). Malinski et al. used 1octanol as a better model of the solvent properties of the lipid membrane and found that the partition coefficient for ·NO between 1-octanol and water was 6.5 (Malinski et al., 1993). More recently, we have shown that the solubility of ·NO in phospholipid membranes is actually ∼3 times greater than in water (Table 2) (Möller et al., 2005, 2007b). Likewise, O2 is approximately 3 times more soluble in membranes than in water (Table 2). The difference with model organic solvents can be explained based on the ordered and packed structure of lipid bilayers that is not randomly distributed as organic solvent molecules (De Young and Dill, 1990). Furthermore, this effect can be explained thermodynamically in terms of the Scaled Particle Theory (Pierotti, 1976; Pollack, 1991). According to this theory, the energies involved in dissolving a molecule in a solvent include not only the interaction between the molecule and the solvent (attractive), but also the work required to create a cavity in the solvent to accommodate the solute (repulsive). This work depends on the surface tension of the solvent, and due to the dense packing of lipids, membranes would be expected to offer a greater resistance to cavity creation and thus lower solubility of ·NO and O2 in the membrane compared with organic solvents. Furthermore, it has been shown that the solubility of O2 changes according to the physical state of the membrane (Table 2, Smotkin et al., 1991; Subczynski and Hyde, 1983), which supports the solute exclusion by lipid packing. On the other hand, the same theory predicts a significant solubility due to the small size of both ·NO and O2 compared with other bigger molecules. Another significant difference with organic solvents is that ·NO is not homogeneously distributed across the membrane, but prefers the acyl chain interior of the lipid bilayer. Molecular-dynamics calculations show that ·NO solubility increases towards the center of the bilayer (Figure 4). The overall solubility of ·NO in the membrane calculated using Sugii et al. data is one order of magnitude higher than experimentally determined values (Sugii et al., 2005), while the solubility calculated from Jedlovszky et al. is similar to experimental values (Jedlovszky and Mezei, 2003). Nitric oxide is mostly excluded from the polar headgroups region of the membrane, while it accumulates in the less polar, more mobile center of the lipid bilayer. This uneven distribution complicates the determination of experimental KP , since different hydrophobic volumes may be considered (Möller

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FIGURE 4 Distribution of ·NO along the lipid bilayer. The partition coefficient for ·NO at different depths of the membrane were calculated using energetic data from Sugii et al. (2005) and from Jedlovszky and Mezei (2003), according to KP = exp−(G/RT ) . The calculated solubility profiles were then scaled to yield an overall KP = 3.6 over the membrane width (40 Å). The square profile corresponds to an homogeneous membrane distribution. The regions 1, 2, 3 and 4 indicate the bulk water region (blue), the high polar headgroups density region (yellow), the high acyl chain density region (orange) and the low acyl chain density, mid bilayer (red), respectively. It can be appreciated how ·NO prefers the lipid acyl chain region over the polar headgroups region, and even more, the less dense and more flexible bilayer lipid interior.

et al., 2005). For convenience, most people use the volume defined by the partial specific volume of the membrane, although it may include some aqueous volume contribution. This is illustrated in Figure 4 by the square profile corresponding to the experimentally determined solubility. The distribution of O2 across the lipid bilayer is essentially the same to that of ·NO, so it will not be discussed in detail. I. Diffusion of ·NO and O2 in Membranes Another important factor regulating reaction rates in membranes, and transport across the membrane is diffusion. Diffusion controls how fast molecules can move within the membrane. It is a random process, with molecules moving in every direction and changing direction every time. If a concentration difference in space exists, then, there will be a net transport of molecules. The diffusion coefficient (D) can be used to calculate how far will a molecule travel (x) in a given time t

2. The Interaction of Reactive Oxygen and Nitrogen Species with Membranes

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(in three dimensions): x = (6D.t)1/2

(6)

The diffusion coefficient for both ·NO and O2 in phosphate buffer is 2.2–4.5 × 10−5 cm2 s−1 (Denicola et al., 1996; Zacharia and Deen, 2005), while in the center of EYPC membranes is 3.1 × 10−6 for ·NO and 6.6 × 10−6 cm2 s−1 for O2 (Denicola et al., 1996; Möller et al., 2005), approximately 10 times slower than in water. Note that the Stokes–Einstein model of diffusion, where diffusion is inversely dependent on solvent viscosity, is not applicable to membranes and small molecules such as ·NO and O2 where the assumption of solute size greater than solvent size is violated (Lieb and Stein, 1986; Möller et al., 2005). There is still no definitive theory about diffusion in membranes, but the steep dependence of permeability on molecular volume suggests that cavities (or “free volume”) are again involved (as in solubility) (Lieb and Stein, 1986). The diffusion process would occur through a series of jumps from an occupied to a free cavity after it forms from molecular fluctuations. The diffusion coefficient would then be proportional to the probability of finding a cavity with an equal or greater volume than the molecule that is diffusing (Lieb and Stein, 1986).

III. CHEMICAL EFFECTS: LIPID PEROXIDATION As mentioned above, membranes could represent a barrier to the transport of some RONS, while others can easily permeate the membrane and react with targets inside the cell, depending on the available targets, their reactivity and quantity. However, the membrane itself is not inert towards these RONS and could be chemically modified in the transport process. Next, we will focus on the chemical effects of RONS on lipids of the membrane. The prototypical effect of reactive species on membrane lipids is lipid peroxidation. Lipid peroxidation affects mainly polyunsaturated fatty acids (2 or more double bonds in the fatty acid chain), and leads to the formation of lipid hydroperoxides, lipid alcohols, and aldehydes. It all begins with an oxidant producing a lipid radical (L·, Figure 5, Initiation). The intermediate radical is stabilized by the addition of oxygen to form the lipid-peroxyl radical (LOO·). This lipid-peroxyl radical is an oxidant and will oxidize nearby lipids, which will add O2 and continue the oxidation cycle chain reaction (Figure 5, Propagation). Therefore, O2 contributes to propagate the oxidation damage, leading to the formation of lipid hydroperoxides (LOOH). These LOOH alter the structure of the lipid bilayer, and can also lead to new oxidant species. In the presence of metal ions, the hydroperoxides can be decomposed to the lipid alkoxyl radical (LO·), which is even more oxidizing and keeps on propagating the oxidative damage (Figure 5, Branching). Lipid hydroperoxides are relatively long lived but break down spontaneously to

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FIGURE 5 The different stages in lipid peroxidation. Lipid peroxidation starts after a one electron oxidation of polyunsaturated fatty acid (LH), followed by a propagation phase, where O2 is added and new radicals are formed from neighboring lipids, accumulating lipid hydroperoxides. The chain reaction is stopped in termination reactions, either by reduction by α-tocopherol, or by radical–radical reactions. Additional lipid radicals can be generated from branching reactions involving lipid hydroperoxide reaction with reduced metal ions, or with superoxide.

yield aldehydes, end-products of lipoperoxidation. In biomembranes, lipid peroxidation stops because the main antioxidant in membranes, α-tocopherol, reduces the propagating LOO· to LOOH (Figure 5, Termination). After seeing the general mechanism of lipid peroxidation, we can now evaluate the potential participation of the different reactive species.

A. Nitric Oxide Nitric oxide (·NO) mostly inhibits the propagation of lipid oxidative damage. is a free radical that contains one unpaired electron. Similarly to O2 , ·NO

·NO

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is not very reactive, reacting only with other free radicals or with metal centers (another unpaired electron is a must). Differently to O2 , the reaction with a lipid radical yields a non-oxidizing species and terminates lipid peroxidation. The very fast reaction with lipid peroxyl radicals, close to the diffusion limit (Padmaja and Huie, 1993; Goldstein et al., 2004), added to the high solubility of ·NO within the membrane (Möller et al., 2005), make ·NO an extremely efficient lipid antioxidant, even more effective than vitamin E (Rubbo et al., 2000). The reaction products of ·NO with lipid radicals have been identified as nitrated lipids with potent anti-inflammatory properties currently under investigation, thus suggesting that this process not only acts to terminate oxidative damage but also initiates specific signaling processes (Rubbo et al., 1994; Baker et al., 2004; Cui et al., 2006). In addition to inhibiting propagation, ·NO may also react with the initiator oxidant, making it a very ample and powerful antioxidant.

B. Nitrogen Dioxide Nitrogen dioxide is a common oxidant emerging from different possible routes and can enter lipid membranes (Khairutdinov et al., 2000). In addition to peroxynitrite homolysis, it can be formed from ·NO autoxidation and from H2 O2 dependent nitrite oxidation by hemeperoxidases (Augusto et al., 2002). ·NO2 can perform hydrogen atom abstraction leading to lipid peroxidation or lipid nitration, and also add to lipid double bonds generating a radical nitrolipid intermediate that may follow different paths. Another ·NO2 can add to form the non-radical dinitrolipid, it can add O2 and generate a propagating nitrolipid peroxyl radical (Pryor et al., 1982), or it can detach from the lipid leading to double bond isomerization from cis to trans conformation (Augusto et al., 2002; Jiang et al., 1999). Because of its H atom abstraction and addition to double bonds, it can be considered an initiator of lipid peroxidation. However, it can also react with lipid peroxyl radicals in a termination reaction at diffusion-limited rates (Goldstein et al., 2004), so it may even act as an antioxidant under certain conditions. The roles that RONS play in lipid peroxidation are complex and very dependent on the system under study. For instance, the high solubility of both ·NO and O2 in the membrane, leads to a higher rate of ·NO autoxidation, forming ·NO2 and N2 O3 (Liu et al., 1998; Möller et al., 2007a, 2007b). ·NO2 will act as as oxidant, or even as antioxidant while ·NO will inhibit oxidation. N2 O3 (equivalent to nitrous acid) could react with unsaturated fatty acids to yield nitroso-lipids, or with lipid hydroperoxides to yield nitro-lipids, lipid nitrates and nitrites (O’Donnell et al., 1999). These reagents may have little impact on membrane functionality, but the secondary products formed like nitrated lipids may have large effects on the cell physiology (Cui et al., 2006). Furthermore, ·NO2 and N2 O3 may well react with proteins associated to the membrane, specially with reactive cysteine residues, and

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lead to S-nitrosated or some other thiol oxidation product of the protein, resulting in an altered function and signaling (Möller et al., 2007a).

C. Peroxynitrite and Carbonate Radical It is not peroxynitrite itself but the radicals derived from it that can oxidize membrane lipids (Radi et al., 1991). Peroxynitrous acid can penetrate the membrane and yield ·NO2 and ·OH, which can both initiate lipid peroxidation (Radi et al., 1991). Alternatively, peroxynitrite-derived radicals can also enter the membrane and initiate lipid peroxidation, and this may be the most important route of oxidation (Botti et al., 2005). The effects of these radicals are dealt in separate sections. The addition of CO2 to the system protects the membrane from oxidation, because carbonate radical can not get into the membrane, and reactions in the aqueous phase consume the oxidant species (Khairutdinov et al., 2000; Romero et al., 1999). The lipid oxidation of low density lipoprotein (LDL) by peroxynitrite is also prevented by CO2 , and reactivity is then focused on exposed protein Apo-B, resulting in enhanced tyrosine nitration (Botti et al., 2005), and this will probably also occur in membranes.

D. Superoxide Superoxide has been associated with lipid peroxidation and cell lysis (Lynch and Fridovich, 1978a). However, it is not involved in initiating lipid peroxidation but rather in propagating it. Preformed lipid hydroperoxides (LOOH) are needed, · which are reduced by O·− 2 generating propagating LO (Lynch and Fridovich, 1978a; Aikens and Dix, 1991). In the experiments performed by Lynch and Fridovich, it was observed that external superoxide dismutase could inhibit cell lysis, indicating that internally generated O·− 2 that could cross the membrane was responsible for cell lysis (Lynch and Fridovich, 1978a). Furthermore, singlet oxygen (1 O2 ) scavengers also protected cell from lysis, indicating that it played an important role in this system, forming additional LOOH. The proposed reactions are shown in Eqs. (7)–(9). · − LOOH + O·− 2 → LO + OH + O2

(7)

+ 1 LO· + O·− 2 + H → LOH + O2

(8)

LH + 1 O2 → LOOH

(9)

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E. Hydrogen Peroxide and Hydroxyl Radical Hydrogen peroxide is not directly reactive to membrane lipids, but can generate hydroxyl radical able to react with most molecules, and initiate lipid peroxidation by H atom abstraction or addition to double bonds. Hydroxyl radical is one of the most oxidizing molecules (Buettner, 1993), and it can react with most organic molecules at practically diffusion-limited rates (Halliwell and Gutteridge, 2000).

F. Biophysical Effects of Lipid Peroxidation in Membranes Lipid peroxidation introduces a bulky and polar group (–OOH) to a phospholipid acyl chain (Buettner, 1993; Wong-ekkabut et al., 2007), altering the tight packing of the lipid bilayer. The oxidized acyl chains occupy a higher area per molecule than non-oxidized molecules and they are more hydrophilic, with the hydroxy- and hydroperoxy-groups more exposed to the aqueous phase (Abousalham et al., 2000; Wong-ekkabut et al., 2007). Although a looser membrane structure would be expected, several biophysical determinations indicate that lipid peroxidation results in a more rigid and less fluid membrane (Richter, 1987), with a higher leakiness and general permeability (Mandal and Chatterjee, 1980). It is also observed a diminished electrical resistance and an increased flip-flop of phospholipids (Richter, 1987). The effect of lipoperoxidation on membrane proteins is to decrease their mobility, both by forming lipid–protein adducts and protein–protein covalent aggregates (Richter, 1987; Soszynski and Bartosz, 1996). Even though the direct reaction of RONS with membrane lipids yields an altered membrane structure with higher permeability, the most important biological consequences of membrane lipoperoxidation may be due to the formation of secondary more reactive species (like aldehydes, peroxides, nitrolipids) capable of modulating signal cascades or develop cellular damage. References Abousalham, A., Fotiadu, F., Buono, G., Verger, R. (2000). Surface properties of unsaturated nonoxidized and oxidized free fatty acids spread as monomolecular films at an argon/water interface. Chem. Phys. Lipids 104, 93–99. Aikens, J., Dix, T.A. (1991). Perhydroxyl radical (HOO.) initiated lipid peroxidation. The role of fatty acid hydroperoxides. J. Biol. Chem. 266, 15091–15098. Antunes, F., Cadenas, E. (2000). Estimation of H2 O2 gradients across biomembranes. FEBS Letters 475, 121–126. Augusto, O., Bonini, M.G., Amanso, A.M., Linares, E., Santos, C.C.X., De Menezes, S.L. (2002). Nitrogen dioxide and carbonate radical anion: Two emerging radicals in biology. Free Radic. Biol. Med. 32, 841–859. Baker, P.R.S., Schopfer, F.J., Sweeney, S., Freeman, B.A. (2004). Red cell membrane and plasma linoleic acid nitration products: Synthesis, clinical identification, and quantitation. Proc. Natl. Acad. Sci. USA 10, 11577–11582.

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Rubbo, H., Radi, R., Anselmi, D., Kirk, M., Barnes, S., Butler, J., Eiserich, J.P., Freeman, B.A. (2000). Nitric oxide reaction with lipid peroxyl radicals spares a-tocopherol during lipid peroxidation. J. Biol. Chem. 275, 10812–10818. Shaw, A.W., Vosper, A.J. (1976). Solubility of nitric oxide in aqueous and nonaqueous solvents. J. Chem. Soc. Faraday Trans. 1 8, 1239–1244. Smotkin, E.S., Moy, F.T., Plachy, W.Z. (1991). Dioxygen solubility in aqueous phosphatidylcholine dispersions. Biochim. Biophys. Acta 1061, 33–38. Soszynski, M., Bartosz, G. (1996). Effect of peroxynitrite on erythrocytes. Biochim. Biophys. Acta 1291, 107–114. Stein, W.D. (1986). Transport and Diffusion across Cell Membranes. Academic Press, Orlando, FL. Stuehr, D.J., Santolini, J., Wang, Z.Q., Wei, C.C., Adak, S. (2004). Update on mechanism and catalytic regulation in the NO synthases. J. Biol. Chem. 279, 36167–36170. Subczynski, W.K., Hyde, J.S. (1983). Concentration of oxygen in lipid bilayers using a spin-label method. Biophys. J. 41, 283–286. Subczynski, W.K., Hyde, J.S., Kusumi, A. (1989). Oxygen permeability of phosphatidylchilinecholesterol membranes. Proc. Natl. Acad. Sci. USA 86, 4474–4478. Subczynski, W.K., Hopwood, L.E., Hyde, J.S. (1992). Is the mammalian cell plasma membrane a barrier to oxygen transport? J. Gen. Physiol. 100, 69–87. Subczynski, W.K., Lomnicka, M., Hyde, J.S. (1996). Permeability of nitric oxide through lipid bilayer membranes. Free Rad. Res. 24, 343–349. Sugii, T., Takagi, S., Matsuoka, Y. (2005). A molecular dynamics study of lipid bilayers: Effects of the hydrocarbon chain length on permeability. J. Chem. Phys. 123, 184714. Takahashi, M., Asada, K. (1983). Superoxide anion permeability of phospholipid membranes and chloroplast thylakoids. Arch. Biochem. Biophys. 226, 558–566. Walter, A., Gutknecht, J. (1986). Permeability of small nonelectrolytes through lipid bilayer membranes. J. Memb. Biol. 90, 207–217. Widomska, J., Raguz, M., Subczynski, W.K. (2007). Oxygen permeability of the lipid bilayer membrane made of calf lens lipids. Biochim. Biophys. Acta 1768, 2635–2645. Wong-ekkabut, J., Xu, Z., Triampo, W., Tang, I.M., Tieleman, D.P., Monticelli, L. (2007). Effect of lipid peroxidation on the properties of lipid bilayers: A molecular dynamics study. Biophys. J. 93, 4225–4236. Zacharia, I.G., Deen, W.M. (2005). Diffusivity and solubility of nitric oxide in water and saline. Ann. Biomed. Eng. 33, 214–222. Zhang, H., Joseph, J., Feix, J., Hogg, N., Kalyanaraman, B. (2001). Nitration and oxidation of a hydrophobic tyrosine probe by peroxynitrite in membranes: Comparison with nitration and oxidation of tyrosine by peroxynitrite in aqueous solution. Biochemistry 40, 7675–7686. Zhang, H., Bhargava, K., Keszler, A., Feix, J., Hogg, N., Joseph, J., Kalyanaraman, B. (2003). Transmembrane nitration of hydrophobic tyrosyl probes. J. Biol. Chem. 278, 8969–8978.

CHAPTER 3 Modulation of Lung Epithelial Sodium Channel Function by Nitric Oxide Weifeng Song, Ahmed Lazrak, Shipeng Wei, Phillip McArdle and Sadis Matalon Department of Anesthesiology, School of Medicine, University of Alabama at Birmingham, Birmingham, AL, 35233, USA

I. Introduction A. Molecular Structure of Amiloride-sensitive Epithelial Sodium Channels B. Molecular Properties of Na+ Channels of Lung Distal Epithelial Cells C. Biophysical Properties of Na+ Channels in Lung Epithelial Cells D. Importance of Na+ Transport in Lung Fluid Balance E. Interactions of NO and RONS with Biological Targets F. Reactive Intermediates Decrease Ion Transport Across the Aveolar Epithelium in Vivo G. Reactive Intermediates Decrease Vectorial Na+ Transport Across ATII Cells in Vitro H. Regulation of Lung Epithelial Na+ Channels by cGMP References

Abstract Alveolar epithelial cells of mammalian lungs actively transport sodium (Na+ ) and chloride (Cl− ) ions and these processes are important in fluid homeostasis. Sodium ions enter the apical membranes of both type I and type II alveolar epithelial cells through sodium selective, cation and cyclic nucleotide gated ion channels and are extruded across the basolateral membrane by the ouabain-sensitive Na,K-ATPase. This vectorial transport of Na+ ions (and concomitant movement of Cl− ions to maintain electroneutrality) creates an oncotic force leading to the reabsorption of fluid across both normal and damaged lungs. Nitric oxide and reactive oxygen nitrogen intermediates (formed by the reactions of nitric oxide with partially reduced oxygen species), generated in close proximity of epithelial cell membranes by activated inflammatory cells, modulate the activity of sodium channels via signal transduction mechanisms (such as increasing PKG and PKC) or by post-translational oxidative modifications of sodium channel proteins and their chaperons. Channel activity (and vectorial sodium transport) may be either Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00203-2

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increased or decreased depending on levels of reactive intermediates and length of exposure. Better understanding of these interactions would offer considerable new insights into the mechanisms leading to pulmonary edema in a number of pathological conditions.

I. INTRODUCTION For gas exchange to occur optimally, the alveoli of the adult mammalian lungs must remain open and free from fluid. In utero, gas exchange occurs across the placenta and the fetal lung is filled with fluid which enters the trachea and distal lung epithelia because of the osmotic gradient created by the active secretion of chloride (Cl− ) ions (Olver and Strang, 1974). Although this fluid is essential for lung growth and development (Bland and Nielson, 1992), it must be rapidly removed shortly after birth to allow for gas exchange to occur. Various studies in rats, sheep, guinea pigs and other species established that just prior to birth, the distal alveolar epithelium converts from Cl− secretion to sodium (Na+ ) absorption and this active transport of Na+ ions across distal lung epithelial cells is essential for the clearance of fetal fluid (Brown et al., 1983; Olver et al., 1986; O’Brodovich et al., 1990; O’Brodovich, 1991). Studies showing reabsorption of intratracheally instilled isotonic fluid or plasma from the alveolar spaces of adult anesthetized animals and resected human lungs, and its near complete inhibition by elimination of the Na,K-ATPase or by replacement of Na+ ions in the alveolar epithelial fluid by large cations, indicate that adult alveolar epithelial cells are capable of actively transporting sodium (Na+ ) ions (Matthay et al., 1982, 1985, 2002; Sakuma et al., 1994). Based on the results of a large number of studies on anesthetized and conscious animals, human lungs as well as isolated epithelial cells, we know that the distal lung epithelial cells (Clara, alveolar type I and II cells) actively transport Na+ ions in a vectorial fashion from the alveolar to the interstitial sides (Figure 1). Na+ ions diffuse passively down their electrochemical gradient (created by the action of the basolaterally located Na,K-ATPase) into alveolar epithelial cells through apically located amiloride-sensitive channels [ENaC; (Cheek et al., 1989; Yue et al., 1995; Jain et al., 2001; Johnson et al., 2006)] or cyclic nucleotide gated ion channels (Kemp et al., 2001) and are extruded across the basolateral cell membranes by the ouabain-sensitive Na+ ,K+ -ATPase (Factor et al., 1998; Ridge et al., 2003). Movement of Na+ ions from the alveolar to the interstitial space necessitates the simultaneous movement of an anion (such as Cl− ) to preserve electro-neutrality. A number of in vivo studies suggested the movement of Cl− ion occurs via both trans-cellular Cl− channels (such as CFTR) as well as para-cellular pathways (Nielsen et al., 1998; Fang et al., 2002). More recent studies have provided convincing evidence of the presence of functional CFTR in both fetal (Lazrak et al., 2002) and adult ATII cells (Brochiero et al., 2004) and showed

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FIGURE 1 Model of ion transport across alveolar epithelial cells. Na+ ions enter the apical membranes of alveolar epithelial (both type II or type I cells) via a variety of ENaC and non-ENaC cation channels (see Table 1), down a favorable electrochemical gradient created by the action of the electrogenic ouabain sensitive-ATP Na,K-ATPase. K+ ions, brought into the cells by the pump in exchange of Na+ ions, leave the cells passively through basolarerally located K+ channels. To maintain neutrality, Cl− ions enter cells through CFTR type channels and exit basolaterally via a Na,K, Cl cotransporter. The combined movement of Na+ and Cl− ions creates an oncotic gradient leading to the reabsorption of alveolar fluid.

that functional CFTR was necessary for the increase of vectorial Na+ transport by β2 -agonists (Fang et al., 2002). Thus in summary, although the vectorial movement of Na+ ions across epithelial cells requires the presence of both passive (channels) and energy consuming (Na,K-ATPAse) basolateral transporters, ion channels constitute the rate-limiting step in this process, offering more than 90% of the resistance to trans-cellular Na+ transport.

A. Molecular Structure of Amiloride-sensitive Epithelial Sodium Channels A complementary DNA (cDNA) encoding an amiloride-sensitive Na+ channel was cloned from the colon of salt-deprived rats using functional expression cloning techniques (Canessa et al., 1993). This clone, termed αrENaC (for the α subunit of the rat epithelial Na+ channel) contains a 2094-nt open reading frame corresponding to 698 amino acids with a predicted molecular mass of 79 kDa. Furthermore, it shares significant homology with mec-4 and deg-1, members of a family of Caenorhabditis elegans genes involved in sensory touch expres-

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sion. Channels formed by the expression of αrENaC in Xenopus oocytes, are highly selective for Na+ over K+ ions, have much higher affinity to amiloride (Ki = 0.1 µM) than EIPA (Ki = 10 µM) and high permeability to Li+ ions (PLi /PNa = 1.6). Subsequently Canessa et al. (1994b) identified and cloned two additional subunits of this channel, named βrENaC and γ rENaC, with molecular masses of 72 kDa and 75 kDa, respectively. Co-expression of all three subunits in oocytes generated 100-fold higher amiloride-sensitive currents than αrENaC alone. Each subunit has two putative trans-membrane domains, yielding a protein with a large (∼50 kDa) hydrophilic loop and short hydrophilic NH2 and COOH termini. αrENaC spans the membrane twice with the short terminal ends on the cytoplasmic side and a large hydrophilic loop in the extracellular space (Canessa et al., 1994a). The αrENaC message was identified only in organs known to contain epithelial Na+ channels, such as kidney medulla and cortex, distal colon and lungs. Experiments utilizing point mutations suggest that all three subunits are involved in pore formation (Schild et al., 1997), although the exact stochiometry varies. Five ENaC subunits, namely α, β, γ , δ and ε ENaC have been cloned to date, although ε ENaC is not found in mammalian cells. Combinations of the various subunits form channels with varying unitary conductances, ion selectivities and regulatory properties (Snyder et al., 1998). In general the β and γ subunits regulate the channel activity of the conducting α, δ and ε ENaC subunits when they are heterologously expressed in oocytes and cell lines. As mentioned above, heterologous expression of α, β, γ ENaC in Xenopus oocytes or mammalian cells form Na+ channels with unitary conductances of about 4–5 pS which are highly permeable to Na+ but poorly permeable to K+ (PNa+ /PK+ > 20) and inhibited by nanomolar concentrations of amiloride. Channels formed by heterologous expression of αβ, αγ ENaC in Xenopus oocytes, exhibit variant amiloride sensitivities, conductances, and Na+ permeabilities (McNicholas and Canessa, 1997; Fyfe and Canessa, 1998). Kizer et al. (1997) reported that expression of the α rENaC from osteoblasts into a null cell line resulted in 24 pS cation channel with equal permeability to Na+ over K+ ions. Expression of αcRNA alone into Xenopus oocytes results in small currents with little sensitivity to amiloride (Canessa et al., 1994b; Chen et al., 2004a). Furthermore, the presence of δENaC significantly alters the biophysical properties of these channels. δβγ ENaC channels have a Na+ conductance of 12 pS (vs. 4 pS for αβγ ENaC) and are less sensitive to amiloride (IC50 : 26 µM vs. 0.1 µM for αβγ ENaC) (Ji et al., 2006); δ but not α ENaC channel activity is gated by extraoocyte protons (Ji et al., 2006) and cGMP (Ji et al., 2007). These studies provide the molecular basis of the wide diversity of ion channels found in a number of epithelia including the lung alveolar epithelium, as discussed below.

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B. Molecular Properties of Na+ Channels of Lung Distal Epithelial Cells Voilley et al. (1994) used a fragment of the rat colon Na+ channel cDNA to screen a human lung cDNA library. One of the hybridizing clones contained a 2007-bp open reading frame, encoding a 699-amino acid protein with a mass of 76 kDa. When injected into oocytes, the cloned channel exhibited sensitivity to amiloride. The sequence shares 81% identity with αrENaC, if methionine-27 of the rat colon sequence is aligned with the first methionine of the human sequence, and is identical with αhENaC cloned from a human kidney cDNA library (McDonald et al., 1994). When total RNA is extracted from whole lungs, there are very low levels of αENaC mRNA during early stages of fetal rat (O’Brodovich et al., 1993), mouse (Dagenais et al., 1997), and human (Voilley et al., 1994) development. Although αrENaC mRNA does increases prior to birth (O’Brodovich et al., 1993), β and γ rENaC subunits are differentially regulated with maximal expression shortly after birth (Tchepichev et al., 1995). Expression of αrENaC mRNA in adult rat ATII cells was also demonstrated by Northern blot analysis (Yue et al., 1995), PCR (Farman et al., 1997) and in situ hybridization (Matsushita et al., 1996; Farman et al., 1997). Additional in situ studies indicate that although β and γ mRNAs are detected in large and small airways they are less abundant in ATII cells as compared to αrENaC (Farman et al., 1997). In early studies, antibodies, raised against the α, β and γ subunits, labeled surface epithelial cells on the rat trachea, bronchi and bronchioles but not on normal alveoli (Renard et al., 1995). However, more recent studies have shown the presence of α, β and γ -ENaC in both isolated and cultured ATII (Jain et al., 1999, 2001; Thome et al., 2003; Hardiman et al., 2004), ATI (Johnson et al., 2002, 2006) as well as in situ (Matthay et al., 2002). A recent study also reported the presence of δENaC in human Clara cell like lines (Ji et al., 2006), although its presence in alveolar epithelial cells has not been confirmed as yet. Antibodies raised against a purified epithelial sodium channel protein isolated from kidney papillae immunostained alveolar epithelial cells both in vitro and in vivo (Matalon et al., 1992; Yue et al., 1995). Finally various subunits of the Na,K-ATPase have been identified by immunofluorescent and electron microscopy techniques in the basolateral membranes of both ATI and ATII cells (Schneeberger and McCarthy, 1986; Ridge et al., 1997, 2003; Johnson et al., 2002). In short, it is now clear that alveolar epithelial cells contain the necessary machinery for vectorial Na+ transport.

C. Biophysical Properties of Na+ Channels in Lung Epithelial Cells Patch clamp measurements of either isolated ATII cells, or ATI and ATII cells in situ have shown the existence of a variety of Na+ channels (summarized in

48

W. Song et al. TABLE 1 Biophysical properties of cation channels based on patch clamp measurements AT II(rat)

Days in cult

g (pS) PNa+ /PK+ Amil(IC50 uM) Regulation

1–4 1–4

20.4 27

1.15 NM

<1 NM

Ca2+ Terbut.; hyperoxia

1–4 1–4 1.5–2 1–4 1–4 1–4 2 and Lung slices 1–4 1–4 2 and Lung slices

25 21 25 20.6 20.6 21 21

7 1 2.08 0.97 0.97 1 1

<1 <1 >1 <1 <1 <1 NM

6 6 6

>80 >80 >40

<1 <1 NM

cAMP/PKA steroids EGF cGMP/PKG NM cAMP/PKA Ca2+ cardiogenic pulmonary edema fluid steroids cAMP/PKA Ca2+ cardiogenic pulmonary edema fluid

Reference

(Feng et al., 1993) (Yue et al., 1994, 1995) (Yue et al., 1995) (Jain et al., 2001) (Kemp et al., 1999) (Jain et al., 1998) (Jain et al., 1999) (Chen et al., 2002) (Gandhi et al., 2007) (Jain et al., 2001) (Chen et al., 2002) (Gandhi et al., 2007)

AT I cells (rat) Lung slices 8.2

NM

<1

Dopamine

Lung slices 22

NM

<1

Dopamine

(Johnson et al., 2006) (Johnson et al., 2006)

AT I/AT II 0–1

21

1

>1

NM

0–1

9

4

<1

NM

0–1

4–5

>40

<1

NM

(Johnson et al., 2006; Helms et al., 2006) (Johnson et al., 2006) (Johnson et al., 2006; Helms et al., 2006)

Table 1) with diverse biophysical properties (single channel conductance, relative permeability for Na+ vs. K+ , amiloride sensitivity), and response to a variety of signal transduction and pharmacological agents). As seen in Table 1, a number of different group of channels have been identified: (1) non-selective (i.e. equally permeable to Na+ and K+ ions), Ca2+ -activated cation channels (Feng et al., 1993) with unknown ENaC subunit composition; (2) highly selective Na+ channels composed of α, β, and γ -ENaC subunits (Jain et al., 2001;

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FIGURE 2 Measurements of alveolar fluid clearance (AFC) in anesthetized, ventilated C57BL/6 mice. AFC was calculated from changes in instilled BSA concentration during a 30-min period. Data are means 6 SD (n ∼ 6 animals for each group). In all cases, the instillate contained 5% fat-free BSA in NaCl or N -methyl-D-glucamine chloride (NMDG-Cl) as indicated. The osmolality was adjusted to that of the plasma for the appropriate group. Amiloride (amil; 1.5 mM) and/or ouabain (forsk; 50 µM) were also added to the instillate as indicated. ∗ P , 0.05 compared with the corresponding control (adapted from Figure 1 of Hardiman et al. (2001) with permission).

Johnson et al., 2006), which are inhibited by an increase of intracellular Ca2+ ; (3) moderately selective or non-selective cation channels consisting of either αENaC alone or combinations of α with β or γ -ENaC (Yue et al., 1994, 1995; Jain et al., 1999, 2001); (4) cGMP-activated cation channels inhibited by Zn+2 (Kemp et al., 2001), which may contain δ-ENaC. A number of factors (including the culture substratum, the presence or absence of steroids, the presence of an air–liquid interface etc.), determine the biophysical properties and subunit composition of these channels (Jain et al., 1999, 2001; Lazrak et al., 2000b). So the question is which type of channels are present in the alveolar epithelium in vivo? Measurements of Na+ dependent alveolar fluid clearance across anesthetized animals are consistent with the presence of a variety of both highly selective and non-selective Na+ channels on alveolar cells. For example, as shown in Figure 2 instilled amiloride blocks between 50–80% of Na+ driven AFC while substitution of Na+ with NMDG totally abolishes it (Matthay et al., 1996; Hardiman et al., 2001; Hickman-Davis et al., 2006); the higher than expected K+ concentrations in the alveolar lining fluid of anesthetized rabbits and the lowering of these values following application of amiloride (Nielson and Lewis, 1990), are consistent with K+ efflux through non-selective or poorly selective cation channels in alveolar epithelial cells. This diversity has to be kept in

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mind when one tries to assess the effects of various agents on Na+ -driven alveolar fluid clearance in vivo. D. Importance of Na+ Transport in Lung Fluid Balance A variety of studies have provided unequivocal evidence that active Na+ transport plays an essential role in the reabsorption of fetal fluid and the established of fluid free air spaces. Instillation of amiloride or benzamil into the fluid filled airspaces of newborn guinea pigs prior to their first breath, resulted in decreased clearance of the fetal fluid and the development of respiratory distress and arterial hypoxemia (O’Brodovich et al., 1990). Neonatal mice, in which αENaC gene was inactivated by gene targeting, were unable to clear the fetal lung fluid and died within 40 h of birth (Hummler et al., 1996). Transgenic expression of α-rENaC driven by a cytomegalovirus promoter in αmENaC(−/−) knockout mice rescued the perinatal lethal pulmonary phenotype and partially restored Na+ transport in renal, colonic, and pulmonary epithelia (Hummler et al., 1997). In addition, studies showing that newborn infants with either transient tachypnea (Gowen et al., 1988) or neonatal respiratory distress syndrome (Barker et al., 1997) had more negative nasal trans-epithelial potential differences (PD) that were poorly inhibited by amiloride, suggesting that impairment of Na+ absorption across the respiratory epithelia of very premature infants may be one of the factors contributing to the pathogenesis of RDS. In contrast βENaC (−/−) mice showed normal prenatal development but on low-salt diets developed type 1 pseudohypoaldosteronism (Pradervand et al., 1999). While it remains unclear whether active Na+ transport plays an important role in keeping alveolar spaces free of fluid in the normal lung, a variety of studies have clearly established that active Na+ transport plays an important role in limiting the degree of alveolar edema in adult mammalian lungs following acute or chronic injury to the alveolar epithelium. For example, intratracheal instillation of a Na+ channel blocker in rats exposed to hyperoxia, increased the amount of extravascular lung water (Yue and Matalon, 1997). Conversely, intratracheal instillation of adenoviral vectors expressing the Na+ ,K+ -ATPase genes increased survival of rats exposed to hyperoxia (Factor et al., 2000). Moreover, patients with acute lung injury who are still able to concentrate alveolar protein (as a result of active Na+ reabsorption) have a better prognosis than those that cannot (Matthay and Wiener-Kronish, 1990; Ware and Matthay, 2001). Finally, decreased Na+ reabsorption predisposes mountaineers to pulmonary edema (Sartori et al., 2002). In addition, abnormalities in ENaC channel number and opening have been linked to the pathogenesis of cystic fibrosis (Stutts et al., 1995) and Liddle’s syndrome (Pradervand et al., 1999). Thus, lung Na+ transport is highly relevant to human lung disease, and it is important to understand the factors which decrease alveolar fluid clearance in vivo.

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Because nitric oxide (NO) and reactive oxygen nitrogen intermediates (RONS) are produced by activated inflammatory cells, in close proximity to the alveolar epithelium and epithelial Na+ channels, a number of studies have been conducted to understand how they may damage epithelial Na+ channels and how this injury can be reversed. Below we review briefly the sources of NO and RONS in the alveolar space and current studies documenting NO and RONS modification of Na+ channel structure function relationships both in vivo and in vitro.

E. Interactions of NO and RONS with Biological Targets NO is generated from three enzymes (eNOS, cNOs and iNOS) which catalyze the oxidative deamination of L-arginine. Potential sources of NO in the lung include both rat and human activated alveolar macrophages (Laskin et al., 1996; Hickman-Davis et al., 2002), neutrophils, alveolar type II cells (Punjabi et al., 1994; Weinberger et al., 1999), and airway cells (Asano et al., 1994). Increased iNOS levels have been found in airway cells and human lung tissue obtained from patients with ARDS (Haddad et al., 1994; Kobayashi et al., 1998; Sittipunt et al., 2001) and other inflammatory lung diseases. The biological effects of NO depend on its concentration, the biochemical composition of the target, and the presence of other radicals (Figure 3). NO may bind to the heme group of soluble guanylate cyclase resulting in increased cellular cGMP levels and activation of PKGs (Ignarro, 1992); it may react with superoxide (O2 •− ) at diffusion limited rates (6.7 × 109 M−1 × s−1 ) to produce peroxynitrite (ONOO− ) (Beckman et al., 1990) and higher oxides of nitrogen; or, in the presence of an electron acceptor, it may react with thiols to form nitrosothiols (RS-NO) (Stamler et al., 1992a, 1992b, 1992c). In spite of the numerous beneficial effects of NO (such as vasodilation, inhibition

FIGURE 3 Generation of reactive oxygen nitrogen intermediates.

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of platelet and neutrophil adherence in endothelial cells, termination of selfpropagation reactions), at high concentrations, NO itself has been shown to inactivate critical enzymes by interacting with their iron-sulfur centers (Molina y Vedia et al., 1992), causing DNA strand breaks which result in the activation of the nuclear enzyme, poly-ADP-ribosyl transferase (Molina y Vedia et al., 1992), and inhibiting both DNA and protein synthesis (Curran et al., 1991; Delaney et al., 1993). However, at physiological concentrations, the reactivity and toxicity of NO is mild, and most of its toxicity has been attributed to either ONOO− or higher oxides of nitrogen. Peroxynitrite is a potent oxidizing and nitrating agent which oxidizes thiols at rates at least 1000-fold greater than H2 O2 at pH 7 (Radi et al., 1991a), causes iron-independent lipid peroxidation of lipids and human low density lipoproteins (Radi et al., 1991b; Graham et al., 1993), nitrates phenolics including tyrosine residues in proteins (Beckman et al., 1992; Greis et al., 1996) and oxidizes proteins (Pryor et al., 1994). Because of this diverse reactivity, ONOO− has been shown to damage a wide spectrum of biological targets such as DNA (Inoue and Kawanishi, 1995), the mitochondria electron transport chain (Radi et al., 1994), lung ion channels (Hu et al., 1994; Duvall et al., 1998; Guo et al., 1998) and the pulmonary surfactant system (Haddad et al., 1993, 1996). Furthermore, ONOO− , because of its very high reactivity, will attack biological targets even in the presence of antioxidant substances (van der Vliet et al., 1994). A number of reports (Denicola et al., 1996; Gow et al., 1996) indicate that physiological concentrations of carbon dioxide and bicarbonate enhance the reactivity of ONOO− via the formation of the nitrosoperoxycarbonate anion, which by decomposing to carbonate radicals and nitrogen dioxide (NO2 ), increases nitration of important physiological targets both in vitro (Zhu et al., 2000) and in vivo (Lang et al., 2005). Highly reactive oxygen nitrogen intermediates are also formed by heme per• oxidase catalyzed reaction of NO− 2 and H2 O2 to form nitrogen dioxide ( NO2 ) which oxidizes and nitrates phenolics (tyrosine and tryptophan) and unsaturated fatty acids (van der Vliet et al., 1995, 1997). The physiological relevance of these reactions were first demonstrated in vitro using neutrophils or monocytes as the source of MPO and H2 O2 (Davis et al., 1991). However, the extent to which these species can nitrate, chlorinate, oxidize specific target proteins and inhibit their function in vivo during lung inflammation was not clarified until very recently. MacPherson et al. (2001) identified eosinophils and eosinophil peroxidase (EPO) as a major source of oxidants during asthma. Finally, elegant studies by Brennan et al. (2002) show that the extent to which EPO- and MPO-catalyzed reactions contribute to tyrosine nitration in vivo is very much dependent on the underlying cause of the inflammatory response, and that nitrogen dioxide (• NO2 ), which is the one-electron oxidation product of NO− 2 , may be involved in these nitration reactions.

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FIGURE 4 Contribution of reactive oxygen nitrogen species to M. pulmonis-mediated alterations of AFC in vivo. MPO(+/+) and littermate MPO(−/−) mice were infected with 107 CFU M. pulmonis and AFC measured at 48 hours later. AFC measurements were performed on ventilated mice and percent clearance was calculated from changes in instilled BSA concentration during a 30 min period. Data are means ± SE, n = 5 to 9 animals for each group. In all cases the instillate contained 5% fat-free BSA in NaCl with the osmolality adjusted to that of murine plasma (322 mosmol/kgH2 O). Amiloride (1.5 mM) was added to the instillate as indicated. ∗ Significant difference from uninfected same strain control; † Significant difference from 48 h infected group, p = 0.04. (Adapted from Hickman-Davis et al. (2006) (Figure 3) with permission.)

F. Reactive Intermediates Decrease Ion Transport Across the Aveolar Epithelium in Vivo Several studies have investigated the possible association between RONS and Na+ transport across the alveolar epithelium in both animals with acute lung injury and patients with cardiogenic edema. Pittet et al. (2001) showed that reabsorption of isotonic fluid (secondary to vectorial transport of Na+ ions) was inhibited during prolonged hemorrhagic shock. Instillation of aminoguanidine, an inhibitor of iNOS, restored fluid re-absorption to normal levels. HickmanDavis et al. (2006) showed mycoplasma infection of C57Bl/6 mice resulted in significant decrease of both Na+ -dependent alveolar fluid clearance and inhibition of amiloride sensitive Na+ currents across alveolar type II cells isolated from these mice and patched in the whole cell mode. However, normal levels of AFC were seen when myeloperoxidase deficient mice, were infected with mycoplasmas (Hickman-Davis et al., 2006), suggesting that reactive species generated by MPO catalyzed contributed to the decrease of AFC by damagina apical and basolateral transporters (Figure 4). Zhu et al. (2001) showed that increased levels of nitrate and nitrite (the stable by-products of NO and RONS) in edema fluid samples of patients with acute lung injury were associated with slower rates of AFC across the lungs of patients with acute lung injury (Figure 5). Reactive oxygen–

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FIGURE 5 Increased levels of nitrate/nitrite in the edema fluid of patients with ARDS are associated with lower levels of AFC. Box-plot summary of mean interval edema fluid nitrate and nitrite concentration versus two categories of alveolar fluid clearance in patients with Acute Lung Injury/Adult Respiratory Distress Syndrome. Maximal alveolar fluid clearance is 1%/h. Impaired/Sub-maximal alveolar fluid clearance is <14%/h. Horizontal line represents the median, the box encompasses the 25th to 75th percentile and the error bars encompass the 10th to 90th percentile. ∗ p < 0.05 by Mann Whitney U-test. From Zhu et al. (2001) (Figure 2) with permission.

nitrogen intermediates have also been shown to decrease ATII cell Na,K-ATPase in thrombin and oleic acid injury by promoting endocytosis from the basolateral plasma membrane via a mechanism involving phosphorylation of PKCζ (Vadasz et al., 2005, 2004). On the other hand, iNOS (−/−) mice as well as alveolar epithelial cells treated with iNOS inhibitors lack amiloride sensitive transport and have lower levels of α and γ ENaC proteins (Hardiman et al., 2004). Thus although increase levels of RONS damage ion transport, basal levels of NO are necessary for the proper function of the amiloride sensitive channels. G. Reactive Intermediates Decrease Vectorial Na+ Transport Across ATII Cells in Vitro More definitive conclusions concerning the effects of NO and RONS on ion channels on alveolar epithelial cells were drawn from studies measuring ion transport across alveolar epithelial cells. Hu et al. (1994) showed that micromolar concentrations of ONOO− decreased amiloride-inhibitable 22 Na+ uptake across

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freshly isolated rabbit ATII cells by at least 40%. Compeau et al. (1994) reported that incubation of confluent monolayers of fetal lung epithelial cells with LPS activated alveolar macrophages for 16 h resulted in a 60% reduction in amiloridesensitive short circuit current (Isc ) and 60% decrease in the density of a 25-pS non-selective cation, Ca2+ activated channel present in the apical membrane of these cells. These effects were abrogated by blocking the ability of alveolar macrophages to generate NO and were associated with a decrease in ENaC mRNA levels. Guo et al. (1998) also reported that NO, generated by a variety of NO donor decreased Isc across confluent monolayers of rat ATII cells with an IC50 of 0.4 µM without affecting transepithelial resistance. NO also inhibited ∼60% of the amiloride-sensitive Isc across ATII cell monolayers following permeabilization of the basolateral membranes, providing strong evidence of down-regulation of amiloride sensitive ENaC channels. Incubation of ATII monolayers with a cell-permeable form of cGMP (8-bromo-cGMP; 400 µM) did not decrease Isc , indicating that at least under these conditions, the inhibitory effects of NO were not mediated by increasing cGMP levels. These findings are in agreement with the original report of Goodman and Crandall also showed that Br-cGMP did not alter dome formation (a parameter for active salt and water transport) by ATII cells (Goodman et al., 1984) and suggest that the effects of NO are mediated through cGMP-independent mechanisms, such as post-translational modifications of either ENaC per se, or structural proteins (such as actin and fondrin) which are necessary for proper action of ENaC. Indeed, 3-morpholinosydnonimine (SIN-1), a generator of ONOO− profoundly inhibited the amiloride-sensitive whole-cell conductance in Xenopus oocytes injected with α, β, γ -rENaC (Duvall et al., 1998) (Figure 6). On the other hand, even supraphysiological concentrations of NO, generated by a variety of NO donors, had no effect on the amiloride-sensitive current of Xenopus oocytes injected with α, β and γ ENaC. ONOO− may exert its effects by oxidizing, nitrating or nitrosylation critical amino acid residues in either ENaC or other structural proteins necessary for its proper function. The α-rENaC subunit has two membrane-spanning domains and an extracellular loop that is rich in tyrosine residues. Mutations or deletions within the extracellular loop of α-rENaC (residues 278–283) affect the ability of amiloride to block α-rENaC channels as well as their open probability (Ismailov et al., 1997). Furthermore, selected mutations within this region of αENaC or within the corresponding region of γ ENaC altered the channel’s response to external Ni+2 and to changes in extracellular Na+ (Sheng et al., 2004). Chen et al. (2004a) generated α-rENaC mutant proteins, where tyrosine residues at positions 279 (mutation α Y279A ), and 283 (mutation α Y283A ) were replaced with alanine, and expressed these mutants, either alone or in combination with wild-type β and γ -rENaC in Xenopus oocytes. Exposure of oocytes to SIN-1 (1 mM) for five minutes decreased both total Na+ and amiloride-sensitive currents across wt- and α Y279A but not α Y283A β, γ -rENaC. Furthermore, exposure to SIN-1 increased the Ki for amiloride across wt but not α Y279A β, γ -rENaC injected oocytes. These

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FIGURE 6 Peroxynitrite decreases amiloride-sensitive currents in Xenopus oocytes. Current-voltage (I–V) relationships from Xenopus oocytes expressing α, β, γ -rENaC. The upper panel shows currents while membrane potentials were stepped from the resting potential to −100 and +100 mV in the absence and presence of amiloride (10 µM). Measurements were then repeated after oocytes were incubated with SIN-1 (1 mM) for 120 min. Amiloride sensitive currents were then calculated as the difference currents. The solid circles are values from oocytes expressing rENaC, but not exposed to SIN-1 (X ± SEM; n = 6). The open circles are values from oocytes expressing rENaC and exposed to SIN-1 (1 mM) for 2 h (X ± SEM; n = 7). The conductance (slope of I–V relationship) was significantly different from zero in the control oocytes (P < 0.01), but not in oocytes exposed to SIN-1 (modified from Duvall et al. (1998) with permission).

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data indicate that both tyrosines are important for proper ENaC function and their oxidative modifications contribute to altered ENaC function. Both the extracellular and trans-membrane segments of ENaC subunits contains domains rich in cysteine residues. Cysteine residues not involved in disulfide formation may become oxidized or nitrosylated which may alter ENaC function. Reactive species (such as superoxide, hydrogen peroxide and hydroxyl radicals) produced by xanthine and xanthine oxidase, decreased amiloride sensitive currents across toad ventral skin mounted in Ussing chambers when added in the apical but not in the basolateral sides, indicating that these reactive species damaged epithelial Na+ channels (Matalon et al., 1989). Furthermore, intracellular oxidizing agents reduced amiloride-sensitive currents in Xenopus oocytes injected with α, β and γ ENaC (Kellenberger et al., 2005). RONS may be downregulating ENaC activity via activation of PKC, which has been shown to reduce ENaC activity and modify its subunit composition. Inhibition of PKC rapidly increased Po and appearance of new channels in patches of A6 (Ling and Eaton, 1989), and lung ATII cells (Chen et al., 2004b) as well in Xenopus oocytes heterologously expressing α, β and γ ENaC (Awayda et al., 1996). In contrast, stimulation of PKC inhibited whole-cell currents in Xenopus oocytes (Awayda et al., 1996). Likewise, PKC activation decreased expression of both β and γ ENaC levels in A6 cells, the activity of single channels and transepithelial Na+ reabsorption (Stockand et al., 2000). ENaC channels are constitutively open at the plasma membrane and do not appear to require additional activation (Canessa et al., 1994b). ENaC activity is regulated by two fundamental mechanisms: changes in the open probability (Po ) of the preexisting channels, or changes in the number of channels at the cell plasma membrane. In the long run, factors that influence both ENaC mRNA and protein levels can potentially modulate amiloride-sensitive Na+ transport in the lung. In the short term, the number of channels at the cell membrane is controlled by a number of mechanisms that involve on one hand the synthesis of new proteins and their exocytosis, on the other hand their removal by endocytosis and their degradation in the proteasome. The ubiquitin-proteasome pathway is an important regulator of ENaC function. The half-life of ENaC in mammalian cell membranes is short (less than 1 hour). ENaC is ubiquitinated in vivo on the α and γ , but not in the β subunits (Staub et al., 1997). Inhibition of ubiquitination or the proteasome results in increased channel activity, due to an increase in the number of channels present at the plasma membrane (Malik et al., 2001). Ubiquitination (ATPdependent serial addition of ubiquitin monomers to lysine residues on proteins), which targets proteins for rapid degradation by the proteasome, is catalyzed by the sequential action of ubiquitin-activating, ubiquitin-conjugating, and ubiquitin protein ligase enzymes (Hochstrasser, 1995). Neural precursor cell-expressed developmentally downregulated protein 4 (Nedd4-2) is the ubiquitin-protein ligase required for ubiquitination of ENaC (Staub et al., 2000). Nedd4-2 is homologous to the ubiquitin ligase and contains three WW domains (Staub and Rotin, 1996;

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Staub et al., 1996) that bind to the PPXY regions of β and γ ENaC subunits. It is assumed that during the interaction of WW domains of Nedd4 with the PPXY domains of ENaC subunits, an E6-AP carboxyl terminus in Nedd4-2 act as a ubiquitin ligase and conjugates ubiquitin to the amino termini of α, β, γ ENaC subunits (Zhou et al., 2007). Ubiquitin binding to α and γ ENaC subunits occurs in MDCK cells and Hek293 cells transfected with the three ENaC subunits (Staub et al., 2000). The co- expression of ENaC subunits and Nedd4-2 in Xenopus oocytes leads to the reduction of ENaC half life at the cell surface and simultaneous decrease in the whole cell amiloride sensitive current was observed (Abriel et al., 1999). Nedd4-2 directly regulates basal ENaC activity by modulating channel stability at the cell surface. In the lung, Nedd4 is mainly expressed in the epithelia lining the airways and in the distal respiratory epithelium, a pattern of expression similar to that of ENaC (Hochstrasser, 1995). Interestingly, the interaction between ENaC and Nedd4-2 is disrupted in Liddle Syndrome, a hereditable form of salt-sensitive hypertension (Snyder et al., 1995). Truncations of the COOH-terminus or mutations in conserved COOH-terminal PY motif abolish the interaction between ENaC and Nedd4-2 resulting in a longer ENaC half-life at the plasma membrane and lower levels of internalization and degradation. This leads to increased amiloride-sensitive current at the apical membrane and increased salt absorption (Snyder et al., 2004, 2005; Knight et al., 2006). The link between PKC and the ubiquitin-proteasome pathway has just recently been made clear. In A6 cells, PKC has been shown to activate the mitogenactivated protein (MAP) kinase kinase kinase Raf-1, and the MAP kinase kinases MAPK/ERK (MEK) 1 and 2. Activation of MEK 1 and 2 was shown to enhance phosphorylation of β and γ , but not αENaC (Shimkets et al., 1998). This phosphorylation event facilitates binding of Nedd4-2 to ENaC, which may then promote ENaC internalization and removal from the cell surface (Staub et al., 2000). Reactive oxygen-nitrogen intermediates have also been shown to decrease ATII cell Na,K-ATPase in thrombin and oleic acid injury by promoting endocytosis from the basolateral plasma membrane via a mechanism involving phosphorylation of PKCζ (Vadasz et al., 2004, 2005).

H. Regulation of Lung Epithelial Na+ Channels by cGMP In contrast to the results cited above, other studies clearly show that • NO modulates cation channel activity in both renal and alveolar epithelial cells by increasing cGMP levels. (Light et al., 1990) demonstrated the presence of a 28 pS cation channel in rat renal inner-medullary collecting duct cells, the activity of which was decreased both by cGMP per se and via cGMP kinase (PKG)-induced phosphorylation. • NO, released from bradykinin stimulated endothelial cells, or NO donors decreased net 22 Na+ flux across isolated perfused cortical collecting

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© 1998 American Physiological Society FIGURE 7 GSNO decreases the open probability (Po ) of cation channels in ATII cells. Acute exposure of AT II cells to S-nitrosoglutathione (GSNO) inhibits non-selective cation channel activity in apical cell-attached patches. Channel activity was measured as open probability (Po ) before and 2 min after addition of 100 µM GSNO to bath. Each Po was calculated from at least 3 min of consecutive recording. Individual data and means + SE from 7 cell-attached patch experiments showing reversibility of effect of GSNO after washout. Each symbol represents a different patch; lines connect data points from the same patch. (Reprinted from Jain et al. (1998) with permission.)

ducts (Stoos et al., 1995) and decreased Na+ short-circuit current across a cortical collecting duct cell line while increasing their cGMP content (Stoos et al., 1995). Helms et al. (2005) reported that NO released from PAPANONOate decreased amiloride sensitive Na+ current across confluent monolayers of Xenopus kidney distal nephron A6 and M1 cortical collecting duct cells mounted in Ussing chambers; furthermore when these cells were patched in the cell attached mode, PAPANOate decreased the open probability of the 4 pS ENaC channels without altering their unitary conductance. In vitro studies on the regulation of Na+ transport by cGMP across confluent monolayers of cultured rat type II alveolar cells (AT2) have led to contradictory results. As mentioned above, cGMP did not alter Isc across rat AT2 monolayers (Guo et al., 1998). In contrast, cGMP, as well as NO, increased Isc and 22 Na influx in tracheal and distal lung epithelial cells (Schwiebert et al., 1997; Rafii et al., 2002). Jain et al. reported that cell permeable forms of cGMP and nitrosoglutathione (GSNO) significantly decreased single channel activity in rat AT2 cells (Figure 7) (Jain et al., 1998). However, Kemp et al. showed that Br-

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FIGURE 8 • NO decreases amiloride sensitive currents in A549 cells through cGMP-dependent mechanisms. (A) Time course recording of whole-cell inward (Na+ ) current, evoked by −100 mV voltage pulses every 10 s before, during and after perfusion of an A549 cell with SES containing 100 µM PAPANONOate. The pipette was filled with the SIS. Whole-cell I–V relations before (B) and five min post PAPANONOate perfusion, when the steady state currents were seen (C). The whole-cell current inhibited by • NO (• NO sensitive) was calculated by digitally subtracting the currents at the steady state effect of • NO (as shown in C) from the current before the perfusion with • NO containing SES (as shown in B). Mean I–V relationships for the total and • NO-sensitive currents are shown in panel D. For panel D, values are means ±1 SEM (n = 6). (From Lazrak et al. (2000a) with permission.)

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cGMP increased cation conductance in rat ATII cells which was totally abolished by Zn2+ (Kemp et al., 2001). Different responses of whole-cell Na+ conductance to cGMP and GSNO in A549 cells were reported by three independent groups (Xu et al., 1999; Kamosinska et al., 2000; Lazrak et al., 2000a) (Figure 8). The most likely explanation for these contradictory responses are the existence of multiple families of Na+ channels in ATII cells (most likely with different ENaC subunit composition than the 4 pS ENaC channels), the properties of which may be modified by culture conditions (Matalon and O’Brodovich, 1999; Jain et al., 2001; Matalon et al., 2002). For example, cell permeable forms of cGMP activated a sodium conductance in Xenopus oocytes following heterologous expression of α, β, γ and δ ENaC but not α, β, γ ENaC alone (Ji et al., 2007). Expression of δ ENaC has been documented by both indirect immunofluorescence and RT-PCR in a variety of human lung epithelial cell lines (Ji et al., 2006). In addition to amiloride sensitive channels, lung epithelial cells contain cyclic nucleotide-gated (CNG) cation channels on their apical membranes (Norlin et al., 2001). The biochemical composition of the cGMP-activated, non-αENaC Na+ channels in lung epithelial cells is not known. In a recent study, Kaestle et al. (2007) report that endogenous nitric oxide, generated by eNOS following a transient increase of left atrial pressure, decreased sodium reabsorption by epithelial cells. This effect was mimicked by intratracheal instillation of high doses of Br-cGMP (1 mM) and ameliorated by agents that inhibited either eNOS or soluble guanylate cyclase. Furthermore, eNOS (−/−) mice were protected from this effect. These data clearly support the hypothesis that cGMP decreases alveolar Na+ reapsorption. It must be stressed that in this study, the authors measured bi-directional fluxes across the alveolar epithelium, so their results cannot be attributed to an increase of Cl− secretion, as previously reported (Chen et al., 2006). Based on these data one may conclude that increased levels of NO will increase the amount of fluid in the lungs of patients with cardiogenic edema. Indeed at least one study has suggested that this is the case (Bocchi et al., 1994). Thus, use of inhaled NO or agents that activate cGMP production (in an effort to reduce pulmonary vasoconstriction) may have to be carefully considered since inhibition of alveolar fluid transport has been correlated with worsening clinical outcome in patients with acute lung injury (Matthay and Wiener-Kronish, 1990; Ware and Matthay, 2001). However, it is important to remember that the experiments of Kaestle et al. were conducted in isolated perfused rats (which lack lymph flow) and these results apply only to an acute increase of left atrial pressure. Also, a recent report indicates that cardiogenic edema fluid (but not plasma) increases amiloride sensitive Na+ transport in both adult ATII cells and across the alveolar epithelium (Gandhi et al., 2007). Thus, additional factors may come into play to prevent the NO decrease of alveolar fluid clearance and prevent fluid accumulation into the alveolar spaces.

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In summary, reactive oxygen nitrogen species play important role in the regulation of cation and ENaC type channels by a activating signal transduction pathways (such as PKC, cGMP, MAPK) and via post-translational modifications of a variety of critical aminoacids (such as cysteines and tyrosines). These changes in structure lead to changes in function which may have important implications in lung fluid clearance during inflammation. Acknowledgements This work was supported by grants HL075540, HL31197, HL51173 and 1U01ES015676. The authors acknowledge the expert editorial assistance of Ms. Teri Potter.

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CHAPTER 4 Effects of Nitrogen Oxides on Chloride Channels Benjamin Gaston University of Virginia School of Medicine, Charlottesville, VA 22908, USA

I. Overview II. Introduction III. Regulation by Nitrogen Oxides of Swelling-activated Cl− Channels IV. Regulation of Calcium Activated Cl− Channels by Nitrogen Oxides (NOx ) V. Effects of Nitrogen Oxides on CFTR A. The Effects of NOx on Wild-type (Wt) CFTR Function B. Wild-type CFTR Inhibition C. Wild-type CFTR Degradation D. Effects or Expression and Maturation of F508 CFTR VI. Future Directions References

I. OVERVIEW

Abnormal chloride (Cl− ) channel activity underlies the pathophysiology of several human diseases. Recently, it has been appreciated that nitric oxide synthase (NOS) activation and/or treatment with exogenous nitrogen oxides (NOx ) can modify the expression, maturation and/or function of a variety of Cl− channel proteins, including the cystic fibrosis (CF) transmembrane regulatory protein (CFTR). Mechanisms underlying these effects are both cyclic GMP (cGMP)dependent and cGMP-independent. A more detailed molecular understanding of these mechanisms may enable targeting of Cl− channels in the development of new therapies. Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00204-4

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II. INTRODUCTION Chloride (Cl− ) channels represent a heterogenous group of proteins. The proteins associated with some Cl− transport functions have not been characterized (Jentsch et al., 2002; Do and Civan, 2004; Suzuki et al., 2006). Stratified according to conductance, there are three broad categories of characterized Cl− channels: small (<10 pico siemens [pS]), middle-sized (10–100 pS) and large (>100 pS; or maxi-Cl− ) (Jentsch et al., 2002; Do and Civan, 2004; Suzuki et al., 2006). Functionally, Cl− channels have also been classified by Suzuki and coworkers to include those involved in 1) ligand-gated transmission in post-synaptic membranes; 2) effects on resting membrane potential in muscle; 3) de-polarization of cells, particularly including smooth muscle cells and retinal epithelial cells; 4) cell volume regulation; 5) fluid transport in epithelia; and 6) ion balance in acidified intracellular vesicles. Many of the functional roles of Cl− channel proteins have been identified in knock-out animals (Jentsch et al., 2002; Suzuki et al., 2006). Abnormalities of Cl− channel proteins in humans are associated with a broad spectrum of diseases, including cystic fibrosis (CF), myotonia congenita, Bartter’s syndrome, Dent’s disease, vitelliform macular dystrophy (Best’s disease) and osteopetrosis (Jentsch et al., 2002; Suzuki et al., 2006). There are five general classes of proteins that are established to represent Cl− channels. The largest family is the CLC family, of which there are at least nine members (Jentsch et al., 2002). Other Cl− channel proteins include the CF transmembrane conductance regulator (CFTR), swelling-activated Cl− channels (ICl swell ), calcium activated Cl− channels (CaCC), and the γ -amino buteric acid/glycine (GABA)- and glycine-receptor ligand-gated Cl− channels (Jentsch et al., 2002; Novak, 2003; Do and Civan, 2004; Suzuki et al., 2006). Cl− channel function, in general, is highly regulated by phosphorylation and/or by the intracellular or intravesicular concentrations of a variety of ions, including protons, Cl− itself, calcium and bicarbonate. For example, CaCCs in airway epithelial cells can be activated by Ca2+ flux and following P2 Y2 -purenergic receptor activation stimulated by ATP; and CFTR activation is coupled to A2b purenergic receptors activated by adenosine (Novak, 2003; Tarran et al., 2006a, 2006b). Additionally, recent evidence suggests that products downstream from nitric oxide synthase (NOS) activation also affect Cl− channel expression and function. These recent observations regarding nitrogen oxide (NOx )-mediated regulation are the subject of the current review. Activation of NOS results in the formation of nitric oxide (NO) and Snitrosothiols (Mayer et al., 1998; Gow et al., 2002; Gaston et al., 2006). These compounds cause both cyclic GMP (cGMP)-dependent and cGMP-independent effects, though it is increasingly apparent that these mechanistic classifications are not mutually exclusive (Mayer et al., 1998; Gow et al., 2002; Gaston et al., 2006; Chen et al., 2006; Sayed et al., 2007; Whalen et al., 2007). Classically, NOSderived NOx activate guanylyl cyclase and, through cGMP, downstream protein

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kinases to cause cellular effects. Additional functional protein modifications, including tyrosine nitration and cysteine S-nitrosylation, mediate important effects downstream from NOS activation (Ricciardolo et al., 2004). Tyrosine nitration is a cytotoxic reaction associated with increased coproduction (or administration) of NO and superoxide, resulting in NO+ 2 addition to tyrosine. This tyrosine modification appears to be relatively irreversible in cellular systems. Cysteine Snitrosylation classically occurs through NO+ (nitrosonium) transfer, in which a proton is lost from a cysteine thiolate and replaced with the NO+ . Over 100 proteins have been identified that are post-translationally modified by S-nitrosylation (Hess et al., 2005). It is important to note that the concentration of the NOS product or exogenous NOx is a critical determinant of effect in biological systems (Ricciardolo et al., 2004; Gaston et al., 2006). For example, low µM concentrations of the endogenous compound, S-nitrosoglutathione (GSNO), cause certain effects through S-nitrosylation and/or guanylyl cyclase activation that are relevant to physiology; whereas high µM or mM concentrations can cause opposite—often toxic—effects through NO radical liberation, tyrosine nitration or even through glutathionylation (Jilling et al., 1999; Bebok et al., 2002; Zaman et al., 2004, 2006; Que et al., 2005; Wang et al., 2005; Gaston et al., 2006; Sayed et al., 2007). In this regard, it is often ill-advised to think of S-nitrosothiols as “NO donors”: they are physiological signaling molecules in physiological concentrations, working through NO+ transfer reactions (transnitrosylations); and NO radical “donors” only in high (toxic) concentrations (Ricciardolo et al., 2004; Gaston et al., 2006). A number of enzyme systems have been characterized that are involved in nitrosylation and denitrosylation processes; these likely affect NOS-dependent Cl− channel regulation as well (Lipton et al., 2001; Hess et al., 2005; Que et al., 2005; Gaston et al., 2006; Zaman et al., 2006). Recently, it has also become apparent that phosphorylation and S-nitrosylation can operate in tandem as reversible, physiological signaling mechanisms (Hess et al., 2005; Gaston et al., 2006; Carver et al., 2007; Whalen et al., 2007). Each of these mechanisms downstream from NOS activation—guanylyl cyclase activation, tyrosine nitration and cysteine S-nitrosylation—is relevant to regulation of Cl− channel expression and/or function. Each will be reviewed in the sections that follow.

III. REGULATION BY NITROGEN OXIDES OF SWELLING-ACTIVATED Cl− CHANNELS Swelling-activated Cl− transport may be mediated by small, intracellular proteins that, typically, can migrate from cytosol to the cell membrane during cell swelling induced by extracellular hypotonia challenge (Ellershaw et al., 2000; Jentsch et al., 2002; Fürst et al., 2005). Though controversial, evidence suggests that these proteins can interact with, and be phosphorylated by, cGMP-dependent

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protein kinase type 2 (Fürst et al., 2005) downstream from NOS activation. In one study, the S-nitrosothiol, S-nitroso-N-acetyl penicillamine (SNAP), inhibited swelling-induced increases in Cl− conductance in about half of the rabbit portal vein smooth muscle cells analyzed. In other cells, however, SNAP augmented this conductance; and the effect was recapitulated by the cell permeable cGMP analog, 8-bromo cGMP; further, it was inhibited by the guanylyl cyclase inhibitor, ODQ (Ellershaw et al., 2000). Strikingly, there was also a subpopulation of these rabbit portal vein smooth muscle cells in which SNAP inhibited swelling-induced Cl− transport in a cGMP-independent fashion. Thus, both cGMP dependentand independent-effects may be operative under different conditions in the regulation of swelling-dependent Cl− transport by S-nitrosothiols. Dose, location, cell-specific metabolism and condition appear to be critical determinants of these differential effects. However, a great deal more work needs to be done to understand the precise molecular mechanisms underlying these effects under physiological conditions. IV. REGULATION OF CALCIUM ACTIVATED Cl− CHANNELS BY NITROGEN OXIDES (NOx ) In 1997, Kamonsinska and coworkers showed that apical DIDS-sensitive Cl− transport in human lung epithelial cells is inhibited by NOS inhibition; and that this effect of NOS inhibition could be overcome with 100 µM GSNO (Kamonsinska et al., 1997) (Figure 1). This observation was consistent with subsequent work suggesting that NOS inhibition could modify stimulated Cl− transport in the colonic epithelium (Reddix et al., 2000; Stoner et al., 2000). The effect was proposed to represent an effect on CaCC. However, since these publications, additional Cl− channels have been identified and additional information has been discovered regarding sensitivity to DIDS and other agents of Cl− channels (Jentsch et al., 2002; Do and Civan, 2004; Suzuki et al., 2006); more specific pharmacological identification of CFTR- and non-CFTR-mediated epithelial cell apical Cl− transport is now possible. Calcium activated Cl− channels are widely distributed in nature. Hirakawa et al. (1999) found that high concentrations (mM range) of SNAP could activate these channels in aortic smooth muscle cells by causing an increase in intracellular calcium. Consistent with this observation, Kito and Suzuki showed that sodium nitroprusside could increase CaCC activity by increasing intracellular calcium concentrations in pacemaker cells of the guinea pig gastric antrum (Kito and Suzuki, 2003), and Garcia-Mata et al. (2003) showed that SNAP (10 µM) increases calcium-sensitive Cl− conductance in guard cells of higher plant (Vicia) leaves. Of note, the effect observed by Garcia-Mata proved to be cGMPindependent. Paradoxically, Sakagami et al. (2001) showed that sodium nitroprusside can inhibit CaCC currents in retinal contracting pericytes in a cGMP-

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FIGURE 1 Inhibition of whole-cell Cl− current by N G -monomethyl-L-arginine (L-NMMA) and its reversal by GSNO. A: typical recordings showing effects of 100 µM L-NMMA and 100 µM GSNO on whole-cell current. B: current-voltage relationships for recordings shown in A. C: histogram showing effects of L-NMMA and GSNO on whole-cell current. Data are represented as means SD (n = 5). Control value (100%) is shown as dotted line. ∗ P < 0.05 and ∗∗ P < 0.01 compared with control value. (From Kamonsinska et al., 1997; with permission.)

dependent fashion. Importantly, NOS inhibition in these microvascular cells increased CaCC current. Of note in this regard, Parai and Tabrizchi (2005) have observed that CaCC—studied using the inhibitor, niflumic acid—may be involved in regulation of vascular smooth muscular tone downstream from NOS activation. NOS inhibition (with L-NAME) augmented the increase in vascular resistance associated with Cl− -free buffer perfusion during α agonist stimulation. Classically, there are two Cl− channel proteins on the basolateral side of epithelial cells: CLC2 and bestrophin. Bestrophin, so named because its absence is associated with Best’s disease, is a recently discovered protein, believed to function as a CaCC (Duta et al., 2006; Suzuki et al., 2006). Duta et al. (2006) have shown that iNOS activation and exposure GSNO activate bestrophin on the basolateral side of Calu-3 cells through a cGMP-dependent process, likely mediated by PKG-dependent phosphorylation. They hypothesize that GSNO augments transcellular Cl− transport by activating both apical CFTR (Chen et al., 2006) and basolateral bestrophin. Taken together, these data reveal the complexity of NOx signaling: it is not strictly a cGMP-dependent process, and the effect of varying concentrations of different NOx on different cell types—and at different intracellular locations— can be dramatically different; even opposing. These observations underscore the

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possibility that NOS does not behave simply to generate a gaseous free radical, the bioactivities of which are limited by its diffusion and free radical reactivity. These various data regarding dose, agent, calcium-dependence and cGMP-dependence of NOS and of various NOx on CaCC transport will not likely be comprehensively understood until the specific molecular mechanisms involved in these effects are characterized.

V. EFFECTS OF NITROGEN OXIDES ON CFTR Cystic fibrosis is a common and often fatal disease. The effects of NOS products and other exogenous NOx on the expression and function of CFTR have received substantially more attention than the interaction of NOx with any other Cl− channel. NOS products can effect CFTR expression, maturation, degradation and function; cyclic GMP-dependent signaling, nitration chemistry and Snitrosylation signaling are each involved in various ways in these diverse effects.

A. The Effects of NOx on Wild-type (Wt) CFTR Function Building on evidence that NO increases Cl− secretion from airway epithelial cells in vitro (Duszyk, 2001), elegant studies have now established that low µM concentrations of GSNO increase wt CFTR function in Calu-3 cell monolayers (Chen et al., 2006) (Figure 2). There appear to be both cGMP-dependent and cGMP-independent mechanisms by which this occurs (Seidler et al., 1997; Chen et al., 2006). The cGMP-dependent effect is robust and appears to involve G-protein coupled phosphorylation. The mechanism that underlies the cGMPindependent effect is still unknown. Of note, several “corrector” therapies, including GSNO itself (see below) appear to have the potential to increase cell surface expression of the most common human clinical mutant CFTR form, F508. F508 CFTR expressed on the cell surface has the potential to be functional, but it is not yet known whether GSNO increase Cl− conductance through F508 CFTR that has been “corrected” to cell surface.

B. Wild-type CFTR Inhibition S-Nitrosoglutathione at doses 200 µM applied to the cytosolic side of a apical cell membrane can inhibit wt CFTR function, an effect caused by glutathionylation of cysteine 1344 (Wang et al., 2005). However, this effect is unlikely to be of substantial clinical relevance because GSNO concentrations of 200 µM have not previously been reported in vivo, particularly not in the intracellular environment.

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FIGURE 2 GSNO promotes the maturation and function of F508 CFTR but has no effect on wild type CFTR maturation. mIMCD cells were infected with either F508 or wild type CFTR recombinant adenovirus. Cells were left untreated or were treated with 10 µM GNSO for 1 or 2 h prior to cell lysis. Equal amounts of total protein were separated by SDS-PAGE, and CFTR expression was determined by Western blot. A, F508 CFTR: lane 1, mock-infected parental cells; lane 2, F508 CFTR expressing cells, untreated; lane 3, 1-h treatment of 10 µM GSNO; lane 4, 2-h treatment with 10 µM GSNO. B, wild type CFTR: lane 1, mock-infected parental cells; lane 2, wt CFTR expressing cells, untreated; lane 3, 1-h treatment of 10 µM GSNO; lane 4, 2-h treatment with 10 µM GSNO. C, representative recordings of currents elicited by voltage steps (from −100 mV to +40 mV) in F508 CFTR-infected mIMCD cells after stimulation with 20 µM forskolin. Very small currents were recorded in untreated cells. Treatments with GSNO resulted in forskolin-stimulated time- and voltage-independent currents. D, current-voltage relations of currents recorded in C. Open circles, untreated cells; filled circles, GSNO-treated cells. E, specific conductance (Gm ) for GSNO-treated and untreated F508 CFTR- or mock-infected mIMCD cells. GSNO-treated F508 CFTR-infected cells showed a significantly higher Gm (p = 0.033, factorial ANOVA). (From Howard et al., 2003.)

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C. Wild-type CFTR Degradation In stably transfected HELA cells, as well as Calu-3 and murine epithelial cells, 100 µM DETA NONOate decreases CFTR expression, both intracellularly and on the apical surface of polarized cells in culture (Jilling et al., 1999; Bebok et al., 2002; Matalon et al., 2003). The NONOate generates NO radical, which reacts with superoxide to form peroxynitrous acid, nitrating CFTR tyrosines (Bebok et al., 2002). This CFTR nitration leads to increased targeting of the protein for degradation before it reaches full maturation (Jilling et al., 1999; Bebok et al., 2002; Matalon et al., 2003).

D. Effects or Expression and Maturation of F508 CFTR On the other hand, evidence in the last decade has suggested that cysteine nitrosylation—particularly of ubiquitin ligase enzymes—can prevent targeted degradation of specific proteins (Yao et al., 2004; Palmer et al., 2007). Because CFTR maturation is regulated, in part, by degradation, there has been interest in determining whether GSNO or other nitrosylating agents would augment CFTR maturation. Surprisingly, several groups have now reported evidence that GSNO increases expression, maturation and function of F508 CFTR (Zaman et al., 2001, 2004, 2007; Howard et al., 2003; Servetnyk et al., 2006). For example, Howard and coworkers (2003) have shown that exposure of murine renal medullary collecting cells overexpressing F508 CFTR to 10 µM GSNO results in increased CFTR maturation and increased forskolin-driven Cl− transport (Figure 3). This effect of GSNO has features that suggest that the biochemistry involves transnitrosylation (NO+ transfer) reactions (Zaman et al., 2001, 2004, 2006; Howard et al., 2003). Specifically, 1) it is reversed by dithiothreitol; 2) it is inhibited by oxyhemoglobin, but not by N-ethyl-maleimide-pretreated oxyhemoglobin, suggesting that a reduced cysteine of hemoglobin, depleting NO+ — rather than loss of NO radical to the heme porphyrin group—is responsible for the hemoglobin-mediated inhibition; 3) it is not recapitulated by 8-bromo-cGMP; 4) GSNO is only one of a class of nitrosylating agents that increase wild type and F508 CFTR expression and maturation; and 5) this class compounds ethyl nitrite, which produces little NO (Zaman et al., 2006). Indeed, the potency of nitrosylating agents in increasing the CFTR expression and maturation is not related to the ability of these agents to serve as unstable “NO radical donors” (Zaman et al., 2006). The effects of these agents on CFTR expression and maturation are both transcriptional and post-translational. Low µM levels of GSNO increase CFTR transcription by differential effects on the stabilization of transcription factors Sp1 and Sp3; higher GSNO concentrations (especially those close to 1 mM) actually

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FIGURE 3 Effect of GSNO on Sp3/Sp1–DNA-binding activity. (A) EMSA using a consensus Sp3/Sp1 oligonucleotide as probe and nuclear extracts from control (lane 1) and GSNO-treated cells at concentrations of 0.5–500 mM (lanes 2–6). Physiological GSNO concentrations increased Sp3/Sp1–DNA binding. Supraphysiological GSNO concentrations inhibited Sp3 binding but augmented Sp1 binding. (B) Nuclear protein extracts were prepared from A549 cells treated with 1, 5, 10 and 100 mM GSNO for 6 h and then subjected to immunoblot analysis using anti-Sp1, Sp3 and a-tubulin antibodies. Molecular masses of Sp1 and Sp3 were 112 and 90–112 kDa respectively. Results are representative of three separate experiments. (C) The induction of Sp1/Sp3–DNA-binding activities by 10 mM GSNO was confirmed by supershift analysis. Antibodies recognizing Sp1 and Sp3 were added after the addition of radiolabelled oligonucleotide to the nuclear-binding reaction. Results are representative of three separate experiments; Ab, antibody. (D) EMSA was performed on nuclear extracts from A549 cells, with 5 mg of nuclear extracts from control and GSNO-treated cells (10 mM) in the presence or absence of 10 mM oxyHb, or oxyHb pretreated with 10 mM NEM for 4 h. Results are representative of three experiments. (From Zaman et al., 2004, with permission.)

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FIGURE 4 GSNO increases Isc across Calu-3 cells. Confluent monolayers of Calu-3 cells were mounted in an Ussing chamber. Upper panel, GSNO (10 µM) was added into the apical compartment. Isc (left axis) and Rt (right axis) were measured as described in the text. Once Isc stabilized, three boluses of glibenclamide (0.1 mM each) were added into the apical compartment. Results are shown of typical experiments. See Table 1 for mean values. Middle panel, CFTRinh-172 inhibits the GSNO-induced Isc in Calu-3 cells. Following addition of GSNO (10 µM) into the apical compartment, addition of the specific CFTR inhibitor CFTRinh-172 inhibited Isc in a dose-response fashion. Results are shown for a typical experiment. Lower panel, dose-dependent increase of Isc by GSNO. GSNO (0.05–200 µM) were added sequentially into the apical compartments of Ussing chambers containing Calu-3 cells. The percentage of maximum GSNO-induced current (y-axis) was calculated as follows: % = [(Ix − I0 )/(I200 − I0 )] × 100, where I0 is the current obtained without GSNO, Ix is the steady-state current with a given concentration of GSNO (in µM), and I200 is the current measured during with 200 µM GSNO. ki was calculated by fitting the data with the equation, Y = I200 × (ki hc)/[ki hc + Ix ], where hc is the Hill coefficient using Microcal Origin Version 6. Each point represents the mean ±1 S.E. of measurements obtained in six different monolayers. (From Chen et al., 2006, with permission.)

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FIGURE 5 GSNO exposure results in SNO trafficking through, and altered F508 CFTR association of proteins in, the ER and Golgi. A. SNO-modified proteins were immunoprecipitated with CFTR from the ER and Golgi fractions at various times following exposure to 10 µM GSNO in polarized human bronchial epithelial CFBE41o− cells. SNO concentration was assayed by Cu/cys reduction-chemiluminescence (22,114,138). B. Immunoblots of cysteine string proteins (Csp’s) 1 & 2 from cytosolic, ER and Golgi fractions after immunoprecipitation with anti-CFTR antibody at different timepoints after exposure to 10 µM GSNO. C. Immunoblots showing the effect of 10 µM GSNO on heat shock cognate (Hsc) 70 associated with CFTR in cytosolic, ER and Golgi fractions. 50 µg of protein was loaded per lane (from Ref. 22; Appendix A). (Adapted from Zaman et al., 2006, with permission.)

inhibit CFTR transcription by increasing the expression and DNA-binding of Sp1 (Zaman et al., 2004). Again, differential effects reflect differential chemistry and localization of the targets of GSNO (Figure 4). The post-transcriptional effects observed following a whole-cell pulse of GSNO can be followed by studying nitrosothiol-modified proteins that coimmunoprecipitate with CFTR as it trafficks through the endoplasmic reticulum and Golgi during maturation. The cysteine nitrosylation of these chaperone pro-

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teins in the CFTR interactome (Wang et al., 2006) ultimately favor full maturation of F508 CFTR (Zaman et al., 2006; Figure 5). The molecular targets of GSNO that co-immunoprecipitate with CFTR include both heat shock cognate (Hsc) 70 and heat shock protein 70-heat shock protein 90 organizing protein (Hop). In the case of Hsc 70, it appears that S-nitrosylation of a single cysteine in the ATP binding domain is likely to inhibit the function of Hsc 70 as a chaperone, preventing it from targeting misfolded CFTR for degradation (Zaman et al., 2006). In the case of Hop, S-nitrosylation may alter Hop-CFTR interaction to favor full maturation (Zaman et al., 2007). GSNO appears to have additional effects on CFTR through post-translational modifications that are not fully clarified, including an effect on cysteine string protein expression (Zaman et al., 2006). Importantly, the cellular metabolism of GSNO and other nitrosothiols is tightly regulated at the level of synthesis, degradation and localization (Que et al., 2005; Gaston et al., 2006). The study of nitrosylation signaling in general, and as it relates to CFTR maturation and function in particular, is still in its infancy. There is interest in using inhaled GSNO as a corrector therapy for CF. This is particularly appealing because GSNO is an endogenous compound (Gaston et al., 1993), levels of which are low in the CF airway (Grasemann et al., 1999), that may have the added beneficial affects of increasing ciliary beat frequency (Li et al., 2000), relaxing human airway smooth muscle (Gaston et al., 1993; Gaston et al., 1994), improving ventilation perfusion matching (Moya et al., 2001; Snyder et al., 2002), inhibiting the growth of certain microbial agents that colonize the CF airway (Yoon et al., 2006), and, potentially, inhibiting amiloride-sensitive sodium transport (Jain et al., 1998).

VI. FUTURE DIRECTIONS Several core questions remain to be answered. 1. Which Cl− channels are affected by NOx ? In addition to the Cl− channels discussed in this paper, there may be other channels affected by products of NOS activation and other NOx . In particular, channels that may be nitrosothiol-sensitive include channels that are functionally 1) sensitive to modulation by mecuric chloride—which breaks nitrosothiol bonds; 2) acid-responsive, because acid conditions can favor formation of nitrosothiols; and 3) inhibited by dithiothreitol, which reduces nitrosothiol bonds (Jentsch et al., 2002; Do and Civan, 2004; Suzuki et al., 2006; Gow et al., 2007). 2. Molecular characterization of the effects of NOx on Cl− channel expression and function. The structures of many Cl− channels have not been discovered. Indeed, specific proteins associated with some Cl− channel activities remain to be identified.

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Molecular features of these proteins by which they would be modified by cGMPdependent protein kinases and/or by nitrosylation are incompletely understood. Exploiting NOS-Cl− channel interactions for the purpose of pharmacological modification of Cl− function in medicine will likely require a substantially more detailed molecular understanding of these interactions. This is also true for the interactome of these proteins. For example, in the case of CFTR, the design and optimization of pharmacological agents that increase F508 CFTR maturation and function will require a good deal more information about the CFTR interactome and its modification by nitrosylating agents. 3. Improved assays. In the case of nitrosothiol modifications, assays for specific NOx effects remain cumbersome and subject to substantial error (Hausladen et al., 2007; Gow et al., 2007). In particular, because of the lability of S-nitrosothiol bonds, it is difficult to extract nitrosylated proteins from cells and subcellular fractions without modifying the protein and S-NO bond during the process. It is particularly important that the so-called triple iodide method not be used for S-nitrosothiol assays (Hausladen et al., 2007). Indeed, any S-nitrosothiol assay should be paired, in the context of the protein or low mass nitrosothiol being assayed, to other complementary assays that use a different technology (Lancaster and Gaston, 2004; Gow et al., 2007). Fortunately, assays more sensitive and specific than those currently in use are in development. For example there is a recent report of an extremely sensitive new mass spectrometry instrument for measuring plasma GSNO concentrations (Taubert et al., 2007). These types of systems may make the path forward substantially easier to navigate. Once these technical obstacles begin to be cleared, moving forward to study NOS-dependent Cl− channel modifications will likely be extremely rewarding. Treatment for a substantial number of diseases may be facilitated by this new information. References Bebok, Z., Varga, K., Hicks, J.K., Venglarik, C.J., Kovacs, T., Chen, L., Hardiman, K.M., et al. (2002). Reactive oxygen nitrogen species decrease cystic fibrosis transmembrane conductance regulator expression and cAMP-mediated Cl− secretion in airway epithelia. J. Biol. Chem. 277, 43041– 43049. Carver, D.J., Gaston, B., deRonde, K., Palmer, L.A. (2007). Akt-mediated activation of HIF-1 in pulmonary vascular endothelial cells by S-nitrosoglutathione. Am. J. Respir. Cell Mol. Biol. 37, 255–263. Chen, L., Patel, R.P., Teng, A. (2006). Mechanisms of cystic fibrosis transmembrane conductance regulator activation by S-nitrosoglutathione. J. Biol. Chem. 281, 9190–9199. Do, C.W., Civan, M.M. (2004). Basis of chloride transport in ciliary epithelium. J. Membrane Biol. 200, 1–13. Duszyk, M. (2001). Regulation of anion secretion by nitric oxide in human airway epithelial cells. Am. J. Physiol. 281, L450–L457. Duta, V., Duta, F., Puttagunta, L., Befus, A.D., Duszyk, M. (2006). Regulation of basolateral Cl− channels in airway epithelial cells: The role of nitric oxide. J. Membrane Biol. 213, 165–174.

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Ellershaw, D.C., Greenwood, I.A., Large, W.A. (2000). Dual modulation of swelling-activated chloride current by NO and NO donors in rabbit portal vein myocytes. J. Physiol. 528, 15–24. Fürst, J., Schedlbauer, A., Grandini, R., Garavaglia, M.L., Saino, S., Gschwentner, M., Sarg, B., et al. (2005). Icln159 folds into a plecstrin homology domain-like structure. J. Biol. Chem. 280, 31276–31282. Garcia-Mata, C., Gay, R., Sokolovski, S., Hills, A., Lamattina, L., Blatt, M.R. (2003). Nitric oxide regulates K+ and Cl− channels in guard cells through a subset of abscisic acid-evoked signaling pathways. Proc. Natl. Acad. Sci. USA 100, 1116–1121. Gaston, B., Reilly, J., Drazen, J.M., Fackler, J., Ramdev, P., Arnelle, D., Mullins, M., et al. (1993). Endogenous nitrogen oxides and bronchodilator S-nitrosothiols in human airways. Proc. Natl. Acad. Sci. USA 90, 10957–10961. Gaston, B., Drazen, J.M., Jansen, A., Sugarbaker, D.A., Loscalzo, J., Stamler, J.S. (1994). Relaxation of human bronchial smooth muscle by S-nitrosothiols in vitro. J. Pharmacol. Exp. Ther. 268, 978– 984. Gaston, B., Singel, D., Doctor, A., Stamler, J.S. (2006). S-nitrosothiol signaling in respiratory biology. Am. J. Respir. Crit. Care Med. 173, 1186–1193. Gow, A.J., Chen, Q., Hess, D.T., Day, B.J., Ischiropoulos, H., Stamler, J.S. (2002). Basal and stimulated protein S-nitrosylation in multiple cell types and tissues. J. Biol. Chem 277, 9637–9640. Gow, A., Doctor, A., Mannick, J., Gaston, B. (2007). S-nitrosothiol measurements in biological systems. J. Chromatog. B 851, 140–151. Grasemann, H., Gaston, B., Fang, K., Ratjen, F. (1999). Decreased levels of nitrosothiols in the lower airways of patients with cystic fibrosis and normal pulmonary function. J. Pediatr. 135, 770–772. Hausladen, A., Rafikov, R., Angelo, M., Singel, D.J., Nudler, E., Stamler, J.S. (2007). Assessment of nitric oxide signals by triiodide chemiluminescence. Proc. Natl. Acad. Sci. USA 104, 2157–2162. Hess, D.T., Matsumoto, A., Kim, S.O., Marshall, H.F., Stamler, J.S. (2005). Protein S-nitrosylation: Purview and parameters. Nat. Rev. Mol. Cell Biol. 6, 150–166. Hirakawa, Yl., Gericke, M., Cohen, R.A., Bolotina, V.M. (1999). Ca(2+)-dependent Cl(−) channels in mouse and rabbit aortic smooth muscle cells: Regulation by intracellular Ca(2+) and NO. Am. J. Physiol. 277, 1732–1744. Howard, M., Fischer, H., Roux, J., Santos, B., Gullans, S., Yancey, P., Welch, W. (2003). Mammalian osmolytes and S-nitrosoglutathione promote F508 CFTR protein maturation and function. J. Biol. Chem. 278, 35159–35167. Jain, L., Chen, X.J., Brown, L.A., Eaton, D.C. (1998). Nitric oxide inhibits lung sodium transport through a cGMP-mediated inhibition of epithelial cation channels. Am. J. Physiol. 274, 475–484. Jentsch, T.J., Stein, V., Weinreich, F., Zdebik, A.A. (2002). Molecular structure and physiological function of chloride channels. Physiol. Rev. 82, 503–568. Jilling, T., Haddad, I.Y., Cheng, S.H., Matalon, S. (1999). Nitric oxide inhibits heterologous CFTR expression in polarized epithelial cells. Am. J. Physiol Lung Cell Mol. Physiol. 277, L89–L96. Kamonsinska, B., Radomski, M.W., Duszyk, M., Radomski, A., Man, S.F.P. (1997). Nitric oxide activates chloride currents in human lung epithelial cells. Am. J. Physiol. 272, 1098–1104. Kito, Y., Suzuki, H. (2003). Pacemaker frequency is increased by sodium nitroprusside in the guinea pig gastric antrum. J. Physiol. 546, 191–205. Lancaster, J., Gaston, B. (2004). NO and nitrosothiols: Spatial confinement and free diffusion. Am. J. Physiol. Lung Cell Mol. Physiol. 287, 465–466. Li, D., Shirakami, G., Zhan, X., Johns, R.A. (2000). Regulation of ciliary beat frequency by the nitric oxide-cyclic guanosine monophosphate signaling pathway in rat airway epithelial cells. Am. J. Respir. Cell Mol. Biol. 23, 175–181. Lipton, A., Johnson, M., Macdonald, T., Lieberman, M., Gozal, D., Gaston, B. (2001). S-nitrosothiols signal the ventilatory response to hypoxia. Nature 413, 171–174. Matalon, S., Hardiman, K.M., Jain, L., Eaton, D.C., Kotlikoff, M., Eu, J.P., Sun, J., et al. (2003). Regulation of ion channel structure and function by reactive oxygen–nitrogen species. Am. J. Physiol. Lung Cell Mol. Physiol. 285, 1184–1189.

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CHAPTER 5 A Mitochondria-AOS-Kv Channel Axis in Health and Disease; New Insights and Therapeutic Targets for Vascular Disease and Cancer Gopinath Sutendra and Evangelos D. Michelakis Departments of Medicine, University of Alberta, Edmonton, Canada

I. Introduction II. The Components of the Mitochondria-ROS-Kv Axis A. Mitochondria (Sensor) B. AOS (Mediator) C. Kv Channels (Effector) III. The Mitochondria-AOS-Kv Axis in Hypoxia: HPV IV. The Mitochondria-AOS-Kv Axis, Metabolism and Apoptosis A. The Metabolism of Cancer Cells B. DCA Reverses the Mitochondrial Remodeling, Normalizes the MitochondriaAOS-Kv Axis, Unlocking the Cancer Cells from a State of Apoptosis Resistance C. Pulmonary Arterial Hypertension: Intriguing Parallels with Cancer D. Conclusion References

I. INTRODUCTION Mitochondria, activated oxygen species (AOS) and voltage-dependent potassium channels (Kv) have been in the center of multiple signaling pathways in diverse cell types in both health and disease. In addition to production of energy, mitochondria are now recognized as important sensors of cellular metabolic demand; for example, they are important oxygen sensors and major regulators of cell death (apoptosis). This allows the mitochondria to regulate supply and demand of energy, promoting cell death when the energy production cannot keep up with energy demands. In addition, mitochondria are major producers of AOS, and Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

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thus regulate many redox-sensitive cellular functions. Some of these AOS, like H2 O2 , are relatively stable with long effective radius, reaching “remote” targets, like Kv channels in the plasma membrane. In turn, the regulation of Kv channels results in changes in both intracellular calcium and potassium, which can then regulate mitochondrial function and apoptosis as well. Thus, feedback loops are completed, strengthening and sustaining the effects of an axis involving a sensor (mitochondria), mediator (AOS) and effector (Kv channels). There is now evidence that this axis is involved in many important functions in health and disease. For example, this axis and its coordinated response during hypoxia, can provide the basis for Hypoxic Pulmonary Vasoconstriction (HPV), a mechanism critical for the life in all mammals. Furthermore, the axis appears to be regulating apoptosis and proliferation in an identical manner in a variety of seemingly unrelated diseases, like pulmonary arterial hypertension (PAH) and cancer. Not only does this axis provide new insights in diseases characterized by suppressed apoptosis and increased cellular proliferation, but it provides novel therapeutic targets for these diseases. Normalization of this axis in PAH and cancer leads to reversal of the disease. Here, we describe the components of this axis and its role in health (HPV) and disease (PAH and cancer).

II. THE COMPONENTS OF THE MITOCHONDRIA-ROS-KV AXIS A. Mitochondria (Sensor) The mitochondria are traditionally known as the energy generating centers of the cell (Duchen, 1999). Glucose and free fatty acids (FFA) are the main energy sources of the cell and fuels of mitochondria. Glucose enters the glycolytic pathway in the cytoplasm, where it eventually becomes pyruvate. Pyruvate can remain in the cytoplasm and, in the presence of lactic dehydrogenase (LDH), can become lactic acid and complete glycolysis (Gly). However, in the presence of activated pyruvate dehydrogenase (PDH) on the mitochondrial membranes, pyruvate becomes acetyl-CoA, which in the inner mitochondrial space enters the Krebs’ cycle to complete glucose oxidation (GO). Similarly, FFAs enter the mitochondria to undergo β-oxidation in which acetyl-CoA is produced. The acetyl-CoA from both GO and β-oxidation enters the Krebs’ cycle, where the electron donors NADH and FADH2 are produced. These electron donors can “donate” their electrons to the electron transport chain (ETC). The ETC is comprised of five mega-complexes (I–V) which transfer electrons down a redox gradient. The final acceptor of electrons, at complex IV of the ETC, is molecular oxygen, forming H2 O (respiration). As the electrons flow, H+ are generated and are transferred out of the internal mitochondrial membrane, creating the mitochondrial membrane potential (Ψ m), which is quite negative (more than −200 mV). The ATP synthase (complex V) at the end of the ETC brings H+ back into the inner membrane (thus using the

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stored energy of the Ψ m) and phosphorylates ADP, producing ATP. Therefore respiration is coupled to oxidative phosphorylation. A small percentage of the electrons flowing down the ETC, results in the production of AOS (superoxide), mostly in complexes I and III. In the presence of the mitochondrial manganese superoxide dismutase (MnSOD) superoxide can be dismutated to the more stable H2 O2 , which can diffuse out of the mitochondria relatively easily and regulate remote redox-sensitive targets; thus providing a means of communication between mitochondria and important cellular structures like the cell membrane. Therefore the mitochondria, as sensors of metabolic supply and demand (for example, sensors of hypoxia), via the production of a mediator (AOS, H2 O2 ) can regulate the function of effectors in order to match sensing with a cellular response (Duchen, 1999). An example of an acute response is the modulation of redox-sensitive Kv channels in the membrane and, as discussed below, regulation of vascular tone. This regulation, strategically controlled throughout the body can result in more supply of oxygen in hypoxic tissues by either dilating systemic arteries or constricting pulmonary arteries (PA). Because the production of AOS is proportional to the flow or electrons down the ETC, the signaling ability of the mitochondria is proportional to their overall function, an important requirement for any sensormediator-effector system. Since the efflux of H+ from the inner mitochondrial space is also proportional to the electron flow, the Ψ m correlates with production of AOS and overall mitochondrial function. The fact that Ψ m can easily be quantified and monitored in living cells (using positively charged rhodaminebased fluorescent dyes that are selectively uptaken by the negative mitochondria in proportion to the Ψ m (Duchen, 1999)) has allowed for a more comprehensive understanding of the role of this mitochondria-AOS-Kv channel axis and its role in both health and disease. More sustained chronic responses can also be regulated by the mitochondria, through their “communication” with the nucleus. Mitochondrial activity can also regulate intracellular Ca++ (by changing Ψ m, the buffering of the positively charged cytosolic Ca++ can be regulated (Duchen, 1999)) and again, via the production of AOS, can regulate the activity of Ca++ - and redox-sensitive transcription factors, that can be translocated to the nucleus where they can regulate the expression of a number of genes, potentiating, maintaining or suppressing mitochondrial function. This can achieve long-term control of this sensing system. The vast majority of the proteins required for mitochondrial function are produced by nuclear genes. A mitochondria-regulated transcription factor can therefore regulate both mitochondrial but also membrane (i.e. Kv channels) function allowing for the feedback loops necessary for the plasticity of this important sensing system. As will be discussed below, an example of such a transcription factor is the nuclear factor of activated T cells (NFAT) (Bonnet et al., 2007b; Macian, 2005). Such feedback loops can be activated or inhibited further by therapeutic interventions, which result in the activation or inhibition of the whole

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mitochondria-AOS-Kv axis and might have multiple, but co-ordinated and comprehensive effects in many disease states. Such diseases, PAH and cancer, are examples that we discuss below.

B. AOS (Mediator) The source of AOS production in mitochondria is mostly complex I and III (Duchen, 1999). In some tissues or conditions the primary source of AOS is complex I while in others is complex III (Barrientos and Moraes, 1999; Ide et al., 1999; Chandel et al., 2000; Vanden Hoek et al., 1998). The ETC complexes consist of many subunits. For example, complex I consists of 42 subunits, only 7 of which are encoded by mitochondrial DNA (mtDNA) (Duchen, 1999). Inhibition of the complex (and thus the flow of electrons) proximal to the site of AOS production site would cause a decrease in AOS production. Inhibition of the complex distal to this site would disrupt the flow of electron down the respiratory chain, diverting them to react with O2 and generate AOS. Mutations in several subunits of complex I have been described in many human muscular and neurodegenerative diseases that are associated with increased neuronal AOS production, including Parkinson’s disease. Fibroblast mitochondria from patients with complex I and complex III deficiency show increased superoxide production, but not mitochondria from patients with complex IV deficiency (Pitkanen and Robinson, 1996). Barrientos and Moraes used a genetic model (40% complex I-inhibited human-ape xenomitochondrial hybrids) and a drug-induced model (0–100% complex I-inhibited cells using different concentrations of rotenone) and showed a direct quantitative correlation between the level of complex I function, cell respiration, AOS production and Ψ m (Barrientos and Moraes, 1999). The inhibition of complex I results in collapse of the proton gradient (decreased Ψ m) and therefore a decrease in the mitochondrial activity: decreased complex I activity → decreased respiration + decreased Ψ m+ increased superoxide production (Barrientos and Moraes, 1999). Unlike the other two SODs (the CuSOD and the extracellular SOD), MnSOD undergoes significant regulation. Dynamic induction of MnSOD occurs through a redox-sensitive mediator (Pitkanen and Robinson, 1996). As expected, many patients with complex I (but not complex IV) deficiency have increased MnSOD levels (Pitkanen and Robinson, 1996). This is a defensive mechanism of the mitochondria to the stress induced by the increased superoxide production that the patients display. It is possible therefore that differences in a cell’s redox status might result in different levels of expressed MnSOD. Furthermore, diversity in the function/expression of MnSOD among different tissues is not surprising given the fact that diversity in mitochondria function has been described not only among different organs or tissues (for example pulmonary versus systemic arter-

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ies (Michelakis et al., 2002a)) but even within a single cell (for example, within a cardiomyocyte) (reviewed in McMurtry and Michelakis, 2004). These data further support the importance of approaching the AOS production (and their downstream effects) as part of a comprehensive and fine-tuned sensing and feedback system and directly relating them to overall mitochondrial function and metabolism. It also means that the production of AOS is not a simple or an allor none phenomenon and should always be interpreted in the context of the model, tissue or disease state, in which they are measured. Findings in one tissue or one disease state cannot be extrapolated to others. Unfortunately this has contributed to several conflicts in the literature, some of which are discussed below (Weir and Archer, 2006).

C. Kv Channels (Effector) Potassium channels are transmembrane proteins that selectively conduct potassium and have the ability to alter membrane potential, intracellular ion concentration ([K+ ]i ; [Ca++ ]i ), cell volume and impact multiple functions of the cell such as contractile state, size, proliferation and apoptosis. Kv channels evolved from proteins with two transmembrane domains, such as the Kir channels (Gallin W.J, 2001). Many Kv channels are enriched in cysteine and methionine groups in critical points of the protein and are redox-sensitive making them ideal targets of AOS. Oxidation or reduction of these critical cysteine or methionine groups can alter the conformational structure of the pore of the channel, altering its conduction properties; generally oxidation promotes whereas reduction inhibits the open state of the channel. The intracellular concentration of K+ is much higher than the extracellular (145 mM compared to 4.5 mM) resulting in a spontaneous outward current. The equilibrium potential for K+ (EK ) determined by the Nernst equation is approximately −91 mV and illustrates the prominent contribution of K+ to the cell’s resting membrane potential (EM ), which in smooth muscle cells (SMC) is approximately −60 mV. At resting EM , L-type voltage-sensitive Ca++ channels (a major pathway for Ca++ entry) are closed. Inhibition of Kv channels results in decreased K+ efflux from the cell and propels the cell interior to be more positive. This leads to depolarization of the EM and subsequent activation of L-type Ca++ channels. Therefore, inhibition of Kv channels increases both intracellular K+ and Ca++ levels. Increased [K+ i ] exhibits tonic inhibition on the formation of the apoptosome (Cain et al., 2001) and caspases of several cell types including pulmonary artery smooth muscle cells (PASMC) (Remillard and Yuan, 2004), thereby inhibiting apoptosis. Potassium ions are also influential in regulating many enzymes; one such metabolic enzyme pyruvate dehydrogenase kinase (PDK), which phosphorylates and inhibits the pyruvate dehydrogenase (PDH) has recently been suggested to show dependence on K+ ions (Knoechel et al., 2006).

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Increased [Ca++ i ] not only increases vasoconstriction but also contributes to proliferation (Platoshyn et al., 2000). Not surprisingly, inhibition or downregulation of Kv channel function has been described in HPV, PAH and cancer. Specific redox-sensitive Kv channels, like Kv1.5 and Kv2.1, have not only be shown to directly regulate PASMC EM (Archer et al., 1998) but regulate apoptosis and be directly involved in the mechanisms of HPV, PAH (Bonnet et al., 2007b; McMurtry et al., 2005, 2004; Michelakis et al., 2002b) and cancer (Bonnet et al., 2007a). The production of AOS typically promotes a tonic activation of Kv channels. Hyperpolarized mitochondria are associated with a decreased AOS production and inhibited Kv channels. As it will be discussed below, hyperpolarized mitochondria are also associated with a state of resistance to apoptosis. Therefore, it is not surprising that hyperpolarized mitochondria-decreased AOS—inhibited Kv channels are seen in seemingly unrelated diseases characterized by suppressed apoptosis and increased proliferation, like PAH and cancer (Bonnet et al., 2007a, 2006, 2007b; McMurtry et al., 2005, 2004; Michelakis et al., 2002b). More importantly, drugs that primarily target mitochondria, reversing the mitochondrial hyperpolarization (like the PDK inhibitor Dichloroacetate, DCA), also increase AOS and up-regulate and activate Kv channels, reversing established PAH and cancer in vivo in animal models (Bonnet et al., 2007a, 2006, 2007b; McMurtry et al., 2004; Michelakis et al., 2002b). Once again, these observations underline the importance of a comprehensive approach to this axis in many different conditions and diseases.

III. THE MITOCHONDRIA-AOS-KV AXIS IN HYPOXIA: HPV Anoxic pulmonary vasoconstriction was first described in 1894 in the dog (Bradford and Dean, 1894). Von Euler and Liljestrand published the first detailed studies of HPV in 1946, in the cat (von Euler and Liljestrand, 1946). They described the opposite effects of hypoxia on the systemic and pulmonary circulations and went on to say, “oxygen want and carbon dioxide accumulation. . . call forth a contraction of the lung vessels, which leads to improved conditions for the utilization of the alveolar air.” We now refer to this critical function for the optimization of oxygenation in the body as enhanced ventilation/perfusion (V/Q) matching. In the following year, Motley, Cournand et al observed that breathing 10% oxygen in nitrogen almost doubled pulmonary vascular resistance in man (Motley et al., 1947). After birth, HPV plays an important role in V/Q matching. In the fetus, however, the lungs are not ventilated and the result of the high pulmonary vascular resistance is to direct relatively oxygenated blood returning from the placenta to cross from the right side to the left side of the circulation through the foramen

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ovale and the ductus arteriosus. The observation that an increase in oxygen tension at birth, right after the first few breaths, causes both pulmonary vasodilatation and contraction of the ductus arteriosus (Born et al., 1956) has fascinated physiologists for half a century. The characteristics of HPV have been established. The basic mechanism of HPV exists within the lungs, as it is present in the isolated perfused lung and isolated resistance PAs as well as isolated PASMC, occurs immediately and is reversible upon return to normoxia. In addition, maximal HPV is observed in the small, resistance pulmonary arteries, as opposed to the larger, conduit, proximal pulmonary arteries (Michelakis et al., 2004; Weir et al., 2005). Regarding the O2 sensor in HPV, although somewhat controversial, there is agreement that the list of the redox-based candidate O2 sensors is limited to primarily two, i.e. the PASMC mitochondrial electron transport chain (ETC) versus PASMC NAD(P)H and novel oxidases (NOX) and that the redox mediator from these sensors is either AOS or reducing/oxidizing couples in the cytoplasm (Michelakis et al., 2004; Weir et al., 2005). Although the relative importance of these two candidate oxygen sensors might vary between tissues or disease states, the fact that knockout mice lacking critical components of the NADPH oxidase have intact HPV, suggests that the mitochondria might be the primary oxygen sensor at least in the healthy pulmonary circulation (Archer et al., 1999). The recent interest in the effector of HPV has resulted in more controversy regarding its identity, i.e. the intracellular Ca++ machinery versus PASMC K+ channels, and even further, the molecular identity of the leading K+ channel candidates (Michelakis et al., 2004; Weir et al., 2005). The reason for the significant interest in the identification of the main effector of HPV is also related to the possibility that the PA vasoconstriction that initiates HPV might, if sustained, also initiate the process leading to PA remodeling and PAH. This would mean that the HPV effector might also be the target of pharmacologic or gene therapy approaches to treat PAH (Archer and Rich, 2000). The impressive similarities of the O2 sensing effector mechanisms between several O2 sensitive tissues is supportive of a leading role of O2 sensitive Kv channels as the effectors of HPV. For example, the effector in the response to O2 in the PAs, ductus arteriosus, carotid body, neuroepithelial body and adrenomeddulary cells involves redox mediated inhibition of Kv channels (Michelakis et al., 2004; Weir et al., 2005). At the same time, evidence from both humans and animal models suggests that a selective downregulation of Kv channels that occurs in PAH, might be etiologically associated with the development of PAH (Archer and Rich, 2000). Physiologists have been puzzled by the fact that the resistance PAs constrict to hypoxia while systemic arteries dilate. What is the mechanism by which HPV is intrinsic only in the resistance PASMC? We have shown that the resistance PASMC are different from the systemic arterial SMC in that they have different

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FIGURE 1 Hypoxic pulmonary vasoconstriction (HPV) is intrinsic to the pulmonary circulation: hypoxia causes pulmonary artery constriction and systemic arterial dilatation. This might be due to the differences in the oxygen sensor in these tissues (i.e. the mitochondria). Perfusing pressures of a rat lung and kidney, perfused in series, shows a simultaneous increase in the pulmonary artery pressure and decrease in the renal artery pressure during hypoxia. Freshly isolated pulmonary and renal artery smooth muscle cells (handled and imaged under identical conditions) show that mitochondrial membrane potential (Ψ m, an index of mitochondrial function) is lower in the pulmonary artery compared to the renal artery smooth muscle cell mitochondria.

mitochondria (HPV sensors) and are endowed with more O2 -sensitive Kv channels (HPV effectors) (Michelakis et al., 2002a). Mitochondria are attractive candidate O2 sensors since they consume the majority of the cell’s O2 . They are also the major regulators of the cell’s redox status, since they are the major producers of AOS and form large and diffuse networks throughout the cytoplasm, that are sometimes only obvious when mitochondria are imaged in live cells with confocal microscopy (Figure 1). This allows close proximity to critical regulators of the intracellular Ca++ homeostasis, including the endoplasmic reticulum and the membrane ion channels. In addition, PASMC mitochondria, as will be discussed below, are particularly endowed

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with manganese superoxide dismutase (MnSOD), compared with mitochondria from systemic vascular SMC (Michelakis et al., 2002a). This results in immediate dismutation of the ETC-produced superoxide to H2 O2 . Superoxide is a very unstable molecule, although it has been proposed to play a role in the response to hypoxia; mitochondrial-derived superoxide initiates the stabilization of hypoxiainducible factor 1α (Chandel et al., 2000), which in turn induces the expression of several genes important in hypoxia, such as erythropoietin. In contrast, the more stable mitochondrial-derived H2 O2 (Boveris and Chance, 1973), has a much longer effective diffusion radius than superoxide (Wolin, 2000), can diffuse to the cytoplasm and activate the soluble guanylate cyclase (Burke and Wolin, 1987; Wolin and Burke, 1987) or activate sarcolemmal K+ channels (Barlow and White, 1998; Hayabuchi et al., 1998; Wang et al., 1996) in vascular SMC thus causing vasodilatation. The concept of a “mitochondrial-derived effector” is not new. Mitochondria can also release other mediators, such as glutamate, that can affect sarcolemmal proteins and mediate glucose sensing and insulin secretion in betacells of the pancreas (Maechler and Wollheim, 1999). The mitochondrial ETC was proposed to be the O2 sensor in the carotid body in 1972 (Mills and Jobsis, 1972). Since then multiple studies have supported the role of the mitochondrial ETC as O2 sensor in the carotid body (reviewed in LopezBarneo et al., 2001). The involvement of the PASMC mitochondria in O2 sensing and HPV was proposed in 1993 (Archer et al., 1993b) and since then their role was confirmed in a more definitive manner by the same (Michelakis et al., 2004; Weir et al., 2005) and other groups (Waypa et al., 2001). The role of mitochondria as universal oxygen sensors allows them to match and couple energy (or oxygen) demand with oxygen delivery in the whole body, either by optimizing ventilationperfusion matching (through HPV in the PAs) or oxygen delivery (through the activation of the respiration center in the brain by the carotid body). The evidence suggesting that the mitochondrial ETC could be important in O2 sensing is based in part on the concordant effects of certain ETC inhibitors and hypoxia. In fact, proximal ETC inhibitors are the only class of drugs that can mimic hypoxia in a variety of O2 sensitive tissues. Inhibitors of complex I (rotenone) and complex III (antimycin), but not inhibitors of complex IV (cyanide) mimic hypoxia’s effect in the pulmonary circulation (Archer et al., 1993a; Waypa et al., 2001). Inhibitors of the mitochondrial ETC mimic hypoxia’s effects on the carotid body (e.g. increase sinus nerve activity (Mulligan et al., 1981)) and PA (cause vasoconstriction) (Archer et al., 1993a; Rounds and McMurtry, 1981). Like hypoxia, rotenone and antimycin inhibit Kv current in PASMCs and decrease the production of AOS (Archer et al., 1993b). The mitochondrial uncoupler, carbonyl cyanide p-trifluoromethoxyphenyl-hydrazone (FCCP), also decreases PASMC Kv current (Yuan et al., 1996). Buckler et al made similar observations in the carotid body, noting that both mitochondrial uncouplers, FCCP and 2,4-dinitrophenol (DNP), excite the carotid body by inhibiting an outward K+ current and inducing

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a small inward current. This leads to membrane depolarization of the type 1 cell and voltage-gated Ca++ entry (Buckler and Vaughan-Jones, 1998). Mitochondrial Diversity Explains the Localization of HPV Within the Resistance PAs The critical role of mitochondria in O2 sensing is further supported by the fact that the responses of both hypoxia and proximal ETC inhibitors are opposing in PA versus RA (Michelakis et al., 2002a). Rotenone and antimycin decrease Kv current in isolated PASMC while they increase K+ current in isolated RA SMC. This explains the effects of hypoxia, rotenone and antimycin in endothelium-free isolated arteries and perfused organs (isolated perfused rat lung perfused in series with isolated rat kidney, Figure 1): they all constrict the PAs and dilate the renal arteries. In contrast, both vascular beds constrict to Angiotensin II and 4aminopyridine, a Kv channel blocker (Michelakis et al., 2002a). This suggests that the reason for these opposing effects lies within the vascular SMC and that it involves the sensor-mediator, rather than the effector. This is further supported by the fact that the production of mitochondrial AOS and H2 O2 at baseline and in response to hypoxia and ETC inhibitors is differentially regulated between PA and renal arteries (RA). We measured the baseline levels of AOS production (during normoxia) as well as the effects of hypoxia and ETC inhibitors in isolated, denuded, resistance PA and RA rings using 3 independent methods: Lucigenin-enhanced chemiluminescence, which preferentially detects superoxide anion levels (Choi et al., 1998) and the AmplexRed and DCF (2 ,7 -dichlorofluorescin diacetate) fluorescence assays, which are specific for H2 O2 . The data from all these assays are concordant and they show that the production of H2 O2 is higher in the PA versus the RA at baseline (Michelakis et al., 2002a). Once again, proximal ETC inhibitors (rotenone and antimycin A, 50 µM) mimic hypoxia and they both decease the production of H2 O2 in the PA. However, neither hypoxia nor the proximal ETC blockers alter the production of H2 O2 in the RA. Cyanide 1 µM (a distal ETC, complex IV) blocker does not affect H2 O2 production in either artery (Michelakis et al., 2002a). Evidence that the differences of the AOS production are intrinsic to the mitochondria, comes from experiments with isolated lung versus kidney mitochondria. The production of AOS and H2 O2 from isolated mitochondria was measured with lucigenin-enhanced chemiluminescence and AmplexRed. Once again, the lung mitochondria make more AOS and H2 O2 than the kidney and while hypoxia and proximal ETC inhibitors decrease AOS and H2 O2 in the lung, they do not in the kidney mitochondria (Michelakis et al., 2002a). Since the AOS production is directly linked to mitochondrial respiration, we studied O2 consumption in both lung and kidney mitochondria. Lung mitochondria have significantly lower rates of respiration, compared to kidney mitochondria, and quality control assays show that this is not due to differential damage during isolation (Michelakis et al., 2002a). As discussed above, the AOS produc-

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tion correlates with Ψ m and respiration, in that lower respiration is associated with depolarized Ψ m and higher AOS levels. We used two different Ψ msensitive dyes (JC-1 and TMRM) and showed that mitochondria from RASMC have significantly higher Ψ m than mitochondria from PASMC under identical loading and imaging conditions, when studied in situ (Michelakis et al., 2002a) (Figure 1). When the SMC were superfused with a hypoxic solution, the PASMC mitochondria hyperpolarized whereas the RASMC mitochondria depolarized. Lastly, PA mitochondrial express less complex I and III subunits but more MnSOD compared to the RA mitochondria (Michelakis et al., 2002a). The Model for the Molecular Basis of HPV (Figure 2) The findings discussed support many of the known features of vascular O2 sensing. The high level of H2 O2 production at baseline in the PAs, could explain the relatively vasodilated (low-pressure) state of the pulmonary, compared to the systemic, circulation. Tonically produced H2 O2 from the proximal ETC can diffuse to the cytoplasm and cause Kv channel activation, PASCM hyperpolarization, decreased opening of the voltage-gated Ca++ channels, decreased [Ca++ ]i levels and vasodilatation. During acute hypoxia, the tonic production of this mitochondrial-derived vasodilator (H2 O2 ) decreases, thus promoting Ik inhibition and vasoconstriction, i.e. HPV. In contrast a hypoxia-induced increase in AOS would increase Ik (Gebremedhin et al., 1994) and promote systemic arterial vasodilatation. Controversy remains as to whether hypoxia decreases or increases AOS production. Other groups have indicated that hypoxia increases mitochondrialderived ROS production (Liu et al., 2003; Waypa et al., 2001, 2002). This is more likely due to differences in techniques and preparations used to study AOS. However all groups involved identify that the mitochondria and ROS account for the hemodynamic effects of hypoxia. A recent summary of the controversy on this issue can be found at Weir and Archer (2006). IV. THE MITOCHONDRIA-AOS-KV AXIS, METABOLISM AND APOPTOSIS A shift of the metabolism away from mitochondria (oxidative phosphorylation) and towards the energetically less efficient glycolysis (GO generates 36 moles ATP per mole of glucose whereas Gly generates only 2) has now been shown to be associated with a resistance to apoptosis. As expected, this metabolic shift is associated with a suppression of the mitochondria-AOS-Kv axis (hyperpolarized mitochondria, suppressed AOS, inhibited Kv channels). This characterizes proliferative diseases like PAH and cancer and at first appears paradoxical; why are these proliferative and energy-hungry states associated with the switch to an energetically inefficient metabolism? Is this switch causally related to these diseases? Recent work attempts to explain this fascinating “paradox”, also known in history as the “Warburg effect”.

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FIGURE 2 The mechanism of HPV. Vasodilatation requires the production of H2 O2 (redox mediator) from the mitochondria (sensor), which oxidizes and opens Kv channels, leading to hyperpolarization of the plasma membrane. This inhibits voltage-gated calcium channels, decreasing calcium entry. A hypoxia-induced decrease in the production of AOS in complexes I and III of the mitochondria ETC, results in decreased H2 O2 production, reduction and inhibition of Kv channels; this results in depolarized plasma membrane potential (EM ), opening of the voltage-gated Ca++ channels, influx of Ca++ and pulmonary vasoconstriction, i.e. HPV.

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A. The Metabolism of Cancer Cells Most cancers are characterized by aerobic Gly, i.e. they use glycolysis for energy production, despite the fact that oxygen is present. In 1929, Warburg first observed this and suggested it resulted from mitochondrial dysfunction, preventing the mitochondria-based glucose oxidation (GO) (Warburg, 1930). Because Gly is far less efficient in generating ATP compared to GO, the glycolytic cancer cells up-regulate glucose receptors and significantly increase glucose uptake in an attempt to “catch up”. Positron emission tomography (PET) imaging, the most sensitive way to diagnose cancer, has now confirmed that most solid tumors have significantly increased glucose uptake and metabolism, compared to noncancerous tissues. This bio-energetic difference between cancer and normal cells, might offer a very selective therapeutic target, as glycolysis is not typically seen in normal tissues apart from skeletal muscle during strenuous exercise. However, this area of experimental oncology had remained controversial; the glycolytic profile has traditionally been viewed as a result of cancer progression, not a cause and therefore the interest in targeting tumor metabolism has been low. The glycolytic profile of cancer is difficult to understand, using an evolutionary model of carcinogenesis. First, why would these highly proliferating and energy-demanding cells rely on glycolysis rather than the much more efficient GO? Second, Gly results in significant lactic acidosis, which might cause significant toxicity to the surrounding tissues and the cancer cells themselves. Recent advances have caused a rekindling of the metabolic hypothesis of cancer suggesting that these facts are not as conflicting as they appear at first (Gatenby and Gillies, 2004): Glycolysis Offers an Early Adaptation to the Hypoxic Microenvironment in Carcinogenesis Gatenby and Gillies recently proposed that since early carcinogenesis often occurs in a hypoxic microenvironment, the transformed cells have to rely on anaerobic glycolysis for energy production (Gatenby and Gillies, 2004). Hypoxiainducible factor (HIF) is activated in hypoxic conditions and it has been shown to induce the expression of several glucose transporters and most of the enzymes required for Gly (Robey et al., 2005). For example, HIF induces the expression of pyruvate dehydrogenase kinase (PDK) (Kim et al., 2006), a gate-keeping enzyme that regulates the flux of carbohydrates (pyruvate) into the mitochondria. In the presence of activated PDK, pyruvate dehydrogenase (PDH) is inhibited, limiting the entry of pyruvate into the mitochondria, where GO can take place. In other words, activated PDK promotes completion of Gly in the cytoplasm with metabolism of pyruvate into lactate; inhibited PDK ensures an efficient coupling between Gly and GO. Initially, tumors compensate by increasing glucose uptake into the cells. Furthermore, Gatenby and Gillies list a number of mechanisms through which lactic acidosis facilitates tumor growth: breakdown of extra-cellular matrix al-

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lowing expansion, increased cell mobility/metastatic potential and (along with HIF) activation of angiogenesis (Gatenby and Gillies, 2004). While tumors eventually become vascularized and are not significantly hypoxic anymore (although some tumors remain hypoxic at the core because the quality of the neo-vessel formation is poor) the aerobic glycolytic profile persists This suggests that the (initially adaptive) metabolic remodeling confers a survival advantage to cancer cells. Indeed, recent evidence suggests that transformation to a glycolytic phenotype offers resistance to apoptosis (Plas and Thompson, 2002; Kim and Dang, 2005, 2006). Gly is Associated with Resistance to Apoptosis Several of the enzymes involved in Gly are also important regulators of apoptosis and gene transcription, suggesting that links between metabolic sensors, cell death and gene transcription are established directly through the enzymes that control metabolism (Kim and Dang, 2005). For example, hexokinase activation leads to a significant suppression of apoptosis; activated hexokinase translocates from the cytoplasm to the mitochondrial membranes where it interacts with and suppresses several key components of mitochondria-dependent apoptosis (Kim and Dang, 2005; Pastorino and Hoek, 2003). It is therefore not surprising that hexokinase is upregulated and activated in many cancers (Kim and Dang, 2005; Pastorino and Hoek, 2003). How does this occur? The promoter of hexokinase contains both p53 and HIF response elements and both mutated p53 and activated HIF increase hexokinase expression (Mathupala et al., 1997, 2001). In addition, the oncogenic protein Akt is upregulated in many cancers and induces a glycolytic metabolic profile via a number of mechanisms (Elstrom et al., 2004; Osaki et al., 2004). Akt increases both the expression and activity of hexokinase (Elstrom et al., 2004; Gottlob et al., 2001). The gene that normally antagonizes Akt, PTEN, is mutated (loss of function mutation) in a large number of cancers. Very recent data revealed even more links between p53 and metabolism: p53 regulates the expression of a critical enzyme of Gly via the production of TIGAR and is also directly regulating the expression of a subunit of cytochrome c oxidase, an important element of complex IV of the ETC in mitochondria (reviewed in Pan and Mak, 2007). In other words, the most common molecular abnormality in cancer, i.e. the loss of p53 function, induces metabolic and mitochondrial changes, compatible with the glycolytic phenotype. Similarly, the c-myc transcription factor increases the expression of many enzymes of Gly and can induce this same metabolic phenotype (Kim and Dang, 2005, 2006). In summary, an evolutionary theory of carcinogenesis identifies metabolism and Gly as a critical and early adaptive mechanism of cancer cells against hypoxia, that persist because it offers resistance to apoptosis in cancer cells (Gatenby and Gillies, 2004). The genetic theory on carcinogenesis, also identifies Gly and metabolism as an end-result of activation of many diverse oncogenes, including c-myc, Akt/PTEN and p53 (Pan and Mak, 2007). Therefore, it is possible that

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this metabolic phenotype is centrally involved in the pathogenesis of cancer and is not simply a “by-product” of carcinogenesis. Although it is not clear whether this metabolic phenotype directly induces malignancy, it certainly “facilitates” carcinogenesis (Kim and Dang, 2006). In addition, this metabolic signature is the common denominator of multiple and diverse pathways; which means that if it is therapeutically targeted it might offer selectivity for malignant cells of diverse cellular and molecular origins. Mitochondria, Cancer and Apoptosis Shifting metabolism away from mitochondria (GO) and towards the cytoplasm (Gly), will suppress apoptosis, a form of cell death that is dependent on mitochondrial energy production. Pro-apoptotic mediators, like cytochrome c and apoptosis-inducing factor, are protected inside the mitochondria. When the voltage- and redox-sensitive mitochondrial transition pore (MTP) opens, they are released in the cytoplasm and induce apoptosis. Mitochondrial depolarization and increased AOS are associated with opening of the MTP (Zamzami and Kroemer, 2001). As discussed above, Ψ m and AOS production are dependent on the flux of electrons down the ETC, which in turn are dependent on the production of electron donors (NADH, FADH2 ) from the Krebs’ cycle. Suppressing the entry of pyruvate to the mitochondria and thus the production of acetyl-CoA, will suppress both the Krebs’ cycle and the ETC and thus MTP opening and apoptosis. The decreased flow of electrons will also result in a decrease in AOS and thus an inhibition of Kv channels, depolarizing the cells and eventually resulting in increased intracellular Ca++ and K+ . In turn, these will promote proliferation and further suppress apoptosis. In summary, suppression of mitochondrial function will result in suppression of the mitochondria-AOS-Kv axis and in suppression of apoptosis at two levels: first at the level of mitochondria, where the increased Ψ m is shifted away from the apoptosis threshold and opening of MTP and second at the level of Kv channels, where the increased levels of intracellular K+ will inhibit caspases. Remarkably, all of these abnormalities are reversed, leading to reversal of cancer growth, by a mitochondria-targeting drug, DCA (Bonnet et al., 2007a).

B. DCA Reverses the Mitochondrial Remodeling, Normalizes the Mitochondria-AOS-Kv Axis, Unlocking the Cancer Cells from a State of Apoptosis Resistance We recently showed that several cancer cell lines (non-small cell lung cancer, breast cancer and glioblastoma) had hyperpolarized mitochondria, compared to non-cancer cell lines (Bonnet et al., 2007a), a finding that was first described (but remained unappreciated) by Dr Chen at the Dana Farber Institute in the 80s (Chen, 1988). We also showed that this was associated with suppressed levels of

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FIGURE 3 A glycolytic environment is associated with an anti-apoptotic and pro-proliferative state, characterizing PAH and cancer. Increased entry of pyruvate into the mitochondria by dichloroacetate or LDH inhibition promotes glucose oxidation, increased apoptosis and decreased proliferation (see text for discussion).

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mitochondria-derived AOS and decreased activity and expression of Kv channels (Bonnet et al., 2007a). The Ca++ -sensitive transcription factor NFAT was also active (i.e. nuclear) in the cancer cells (Bonnet et al., 2007a). NFAT is a transcription factor that has been show to increase the levels of the anti-apoptotic mitochondrial bcl-2 and decrease the levels of the Kv channel Kv1.5 (Bonnet et al., 2007b; Macian, 2005). All of these features are compatible with an anti-apoptotic state and could be secondary to suppressed mitochondrial activity (Figure 3): decrease entry of pyruvate would eventually result in decrease flux of electrons in the ETC and therefore decreased AOS production, closing of the existing redox-sensitive Kv channels and increased intracellular Ca++ . The decreased AOS could also contribute to closure of the redox-sensitive MTP and mitochondrial hyperpolarization. The decreased entry of pyruvate into the mitochondria (and therefore the decreased GO) would result in compensatory Gly. Increased hexokinase levels would contribute to the hyperpolarization of the mitochondria; increased hexokinase in a glycolytic environment is known to be translocated to the mitochondrial membrane, inhibiting the voltage-dependent anion channel (a component of the MTP), resulting in hyperpolarization and suppression of apoptosis (Pastorino et al., 2005). By inhibiting PDK (Figure 4), DCA activated the pyruvate dehydrogenase, which resulted in an increased delivery of pyruvate into the mitochondria. As predicted, DCA increased GO and depolarized the mitochondria, returning the Ψ m towards the levels of the non-cancer cells, without affecting the mitochondria of non-cancerous cells. Remarkably, all the above features of the cancer cells were “normalized” following the increase in GO and the mitochondrial depolarization: AOS increased, NFAT was inactivated and function/expression of Kv channels was increased. Most importantly, apoptosis was induced in the cancer cells with both cyt c and apoptosis inducing factor efflux from the mitochondria (Figure 3). This resulted in a decrease in tumor growth both in vitro and in vivo in xenotransplant models. It is important here to clarify that simply inhibiting Gly (particularly at proximal stages), will not promote pyruvate entry into the mitochondria, i.e. will not re-activate mitochondria. It will also be toxic to several non-cancerous tissues that depend on Gly for energy production. Inhibiting Gly (which has previously been tested as a potential treatment for cancer) typically results in necrosis, not apoptosis, since apoptosis is an energy-consuming process, requiring active mitochondria. The “trick” is to enhance the Gly to GO coupling, not just inhibit Gly. One of the ways that this can happen is by activating PDH, or inhibiting LDH, bringing pyruvate into the mitochondria and enhancing GO. This hypothesis is also supported by the recently published work that inhibition of LDH (by siRNA), which promotes the transfer of pyruvate into the mitochondria (in that sense mimicking DCA), also promotes cancer apoptosis and decreases tumor growth in vitro and in mice xenotransplants (Fantin et al., 2006) (Figure 3).

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FIGURE 4 Structure of human pyruvate dehydrogenase kinase 2 (PDK2) bound with dichloroacetate (DCA). (A) Ribbon representation of PDK2 with DCA bound. (B) Surface representation of PDK2 bound with DCA. The carboxylate group of DCA forms a salt bridge (dashed white line) with Arginine 154 of PDK2. (C) Structure of DCA. The structures were generated using visual molecular dynamics (VMD) version 1.8.6 with the coordinates acquired from Protein Data Bank (PDB#2BU8).

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DCA has been used clinically to treat inherited mitochondrial disorders over 30 years (Stacpoole, 1989; Stacpoole et al., 2003). It is a very small molecule (Figure 4), orally available and cheap (it is a generic drug) and the recent findings suggest that it might be a promising selective therapy for cancer. Indeed, clinical phase I and II clinical trials in patients with several forms of cancer have now started at the University of Alberta. Intriguingly, DCA reverses PAH with an identical mechanism, again normalizing the mitochondrial-AOS-Kv axis and promoting apoptosis.

C. Pulmonary Arterial Hypertension: Intriguing Parallels with Cancer Since its first description (Dresdale et al., 1951; Romberg, 1891) PAH has fascinated scientists and, despite the recent progress, remains an intriguing and mysterious disease. PAH pathology spreads through the whole spectrum of vascular disease: there are well-described abnormalities involving the endothelium, the smooth muscle cells, the fibroblasts and the interstitial matrix in the vascular wall as well as abnormalities in platelets and circulating inflammatory cells. These abnormalities result in an obliterative remodeling of the pulmonary circulation, characterized by lumen occlusion in medium and small-sized pulmonary arteries (PA) due to excessive cellular proliferation in the vascular wall (Figure 5); the right ventricular (RV) afterload increases, resulting in RV failure and premature death (Voelkel et al., 2006). Systemic arteries are typically normal in PAH and this represents a major challenge since candidate therapies will have to reverse the vascular remodeling in the PAs without affecting the systemic arteries. Therapies are limited in effectiveness, typically suffer form severe adverse effects and the mortality in PAH remains very high (Archer and Michelakis, 2006). Recent evidence suggests that many features of cancer are also present in PAH. For example, PAs from patients and animals with PAH express cancer markers, like the mitochondrial apoptosis inhibitory protein survivin (McMurtry et al., 2005) and are characterized by a remarkable resistance to apoptosis (Michelakis, 2006). Like in cancer, there is a great need for effective and selective disease-modifying therapies (Archer and Michelakis, 2006). The mitochondria-AOS-Kv axis is an attractive target for novel PAH therapies. As discussed above, the differences in the PA compared to the systemic arterial mitochondria might provide the basis of pulmonary artery-selective therapies. Since the axis is involved in the regulation of both vascular tone and proliferation/apoptosis, its targeting might have several beneficial effects. The mitochondria-AOS-Kv axis is significantly suppressed in PAH, including human and PAH in several animal models. Hyperpolarized mitochondria, decreased AOS and inhibited/downregulated Kv channels have been described in PASMC from patients with clinical PAH and in several rat models of the disease (Bonnet et al., 2006, 2007b; McMurtry et al., 2005, 2004; Michelakis et al.,

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FIGURE 5 PAH pathology: immunohistochemistry in a human sample from a patient with PAH; staining with an antibody against smooth muscle actin shows the impressive proliferation of the PASMC in the media, essentially obliterating the PA lumen. As discussed in text, the PA media in PAH is characterized by suppressed apoptosis and increased proliferation, resembling a neoplastic tissue. Like in cancer, this is associated with a suppressed mitochondria-AOS-Kv channel axis.

2002b). In addition, this axis remains suppressed even in isolated PASMC in culture. This phenotype likely contributes significantly to the resistance in apoptosis that underlines the pathology of PAH. Even more remarkable is the fact that DCA, reverses all animal models of established PAH by the exact and identical mechanism that it inhibits cancer growth (Figure 6 and Table 1) (Bonnet et al., 2007a; McMurtry et al., 2004). Both in vivo and in vitro DCA decreases and normalizes the mitochondrial Ψ m, increases H2 O2 production and acutely activates Kv channels. Like in cancer, PAH is also associated with an activated NFAT, explaining in part the down-regulation of Kv channels and the regulation of several mitochondrial enzymes, in order to produce a mitochondrial remodeling similar to that of cancer (Bonnet et al., 2007b).

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FIGURE 6 The mitochondria acting Dichloroacetate reverses cancer (Bonnet et al., 2007a) and PAH (McMurtry et al., 2004) by an identical mechanism including mitochondrial depolarization, activation of K+ channels, induction of apoptosis, resulting in normalization of hemodynamics in rat PAH and decrease in cancer growth in a xenotransplant model of non-small cell lung cancer in nude rats. Representative pictures of the effects of DCA in PASMC and cancer mitochondrial Ψ m (TMRM staining) and outward K+ current (patch claming) are shown. These result in induction of apoptosis, measured by TUNEL (green: TUNEL, blue: nuclear staining by DAPI) and reversal of the disease: the decreased PA pressure (measured by catheterization of anesthetized rats via the jugular vein) and the decreased in tumor growth (measured by a rat microCT). (For colors see the web version of this book.)

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A Normal PASMC PAH PASMC PAH PASMC + DCA B Normal Epithelial Cell Cancer Cell Cancer Cell + DCA

Mitochondrial Ψ m

AOS Production

K+ Current

↓ ↑ ↓

↑ ↓ ↑

↑ ↓ ↑

Mitochondrial Ψ m

ROS Production

K+ Current

↓ ↑ ↓

↑ ↓ ↑

↑ ↓ ↑

DCA results in reversal of the AOS- and calcium-activated NFAT, leading to the reversal of the Kv channel downregulation as well (McMurtry et al., 2004). Like in cancer, DCA has no effects whatsoever in the non-diseased pulmonary arteries and in the normal systemic arteries (McMurtry et al., 2004). One would expect that the vascular wall in PAH would be characterized by a glycolytic phenotype, like in cancer’s Warburg effect. Although direct measurements of metabolism within the PA wall in PAH are not yet available, it is intriguing that a recent report describes high glucose uptake within the PA wall in patients with PAH, using PET imaging (Xu et al., 2007). This raises the intriguing possibility that the anti-apoptotic glycolytic environment in the remodeled PA wall in PAH is associated with compensatory increased uptake of glucose, exactly like in cancer. D. Conclusion These recent observations on the mitochondria-AOS-Kv axis in PAH and cancer open a new window in our understanding of these diseases, their therapy and even their diagnosis with metabolic-based molecular imaging techniques. These fascinating parallels between a vascular remodeling disease like PAH and cancer suggest that the mitochondria-AOS-Kv axis and metabolism might play an important role in other vascular disorders characterized by a proliferative remodeling (like restenosis or transplant vasculopathy) and that therapies that target this axis might be effective in those diseases as well. Molecular medicine calls for a look into mechanisms of disease without the traditional limits of medical specialties and suggests that comprehensive signaling systems like the mitochondriaAOS-Kv axis should be studied and therapeutically targeted in many seemingly unrelated diseases. References Archer, S., Rich, S. (2000). Primary pulmonary hypertension: A vascular biology and translational research “Work in Progress”. Circulation 102, 2781–2791.

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McMurtry, M.S., Bonnet, S., Wu, X., Dyck, J.R., Haromy, A., Hashimoto, K., Michelakis, E.D. (2004). Dichloroacetate prevents and reverses pulmonary hypertension by inducing pulmonary artery smooth muscle cell apoptosis. Circ Res. 95, 830–840. McMurtry, S., Michelakis, E. (2004). Mitochondrial diversity in the vasculature: Implications for vascular oxygen sensing. In: Yuan, J. (Ed.), Hypoxic Pulmonary Vasoconstriction: Cellular and Molecular Mechanisms. Kluwer, New York (Chapter 17). Michelakis, E.D. (2006). Spatio-temporal diversity of apoptosis within the vascular wall in pulmonary arterial hypertension: Heterogeneous BMP signaling may have therapeutic implications. Circ. Res. 98, 172–175. Michelakis, E.D., Hampl, V., Nsair, A., Wu, X., Harry, G., Haromy, A., Gurtu, R., Archer, S.L. (2002a). Diversity in mitochondrial function explains differences in vascular oxygen sensing. Circ. Res. 90, 1307–1315. Michelakis, E.D., McMurtry, M.S., Wu, X.C., Dyck, J.R., Moudgil, R., Hopkins, T.A., Lopaschuk, G.D., Puttagunta, L., Waite, R., Archer, S.L. (2002b). Dichloroacetate, a metabolic modulator, prevents and reverses chronic hypoxic pulmonary hypertension in rats: Role of increased expression and activity of voltage-gated potassium channels. Circulation 105, 244–250. Michelakis, E.D., Thebaud, B., Weir, E.K., Archer, S.L. (2004). Hypoxic pulmonary vasoconstriction: Redox regulation of O2 -sensitive K+ channels by a mitochondrial O2 -sensor in resistance artery smooth muscle cells. J. Mol. Cell Cardiol. 37, 1119–1136. Mills, E., Jobsis, F.F. (1972). Mitochondrial respiratory chain of carotid body and chemoreceptor response to changes in oxygen tension. J. Neurophsiol. 35, 405–428. Motley, H.L., Cournand, A., Werko, L., Himmelstein, A., Dresdale, D. (1947). Influence of short periods of induced acute anoxia upon pulmonary artery pressure in man. Am. J. Physiol. 150, 315–320. Mulligan, E., Lahiri, S., Storey, B.T. (1981). Carotid body O2 chemoreception and mitochondrial oxidative phosphorylation. J. Appl. Physiol. 51, 438–446. Osaki, M., Oshimura, M., Ito, H. (2004). PI3K-Akt pathway: Its functions and alterations in human cancer. Apoptosis 9, 667–676. Pan, J.G., Mak, T.W. (2007). Metabolic targeting as an anticancer strategy: Dawn of a new era? Sci. STKE 2007, pe14. Pastorino, J.G., Hoek, J.B. (2003). Hexokinase II: The integration of energy metabolism and control of apoptosis. Curr. Med. Chem. 10, 1535–1551. Pastorino, J.G., Hoek, J.B., Shulga, N. (2005). Activation of glycogen synthase kinase 3beta disrupts the binding of hexokinase II to mitochondria by phosphorylating voltage-dependent anion channel and potentiates chemotherapy-induced cytotoxicity. Cancer Res. 65, 10545–10554. Pitkanen, S., Robinson, B.H. (1996). Mitochondrial complex I deficiency leads to increased production of superoxide radicals and induction of superoxide dismutase. J. Clin. Invest. 98, 345–351. Plas, D.R., Thompson, C.B. (2002). Cell metabolism in the regulation of programmed cell death. Trends Endocrinol. Metab. 13, 75–78. Platoshyn, O., Golovina, V.A., Bailey, C.L., Limsuwan, A., Krick, S., Juhaszova, M., Seiden, J.E., Rubin, L.J., Yuan, J.X. (2000). Sustained membrane depolarization and pulmonary artery smooth muscle cell proliferation. Am. J. Physiol. Cell Physiol. 279, C1540–C1549. Remillard, C.V., Yuan, J.X. (2004). Activation of K+ channels: An essential pathway in programmed cell death. Am. J. Physiol. Lung. Cell Mol. Physiol. 286, L49–L67. Robey, I.F., Lien, A.D., Welsh, S.J., Baggett, B.K., Gillies, R.J. (2005). Hypoxia-inducible factor1alpha and the glycolytic phenotype in tumors. Neoplasia 7, 324–330. Romberg, E. (1891). Ueber slerose der lungenarterien. Deutsch Arch. Klin. Med. 48, 197. Rounds, S., McMurtry, I. (1981). Inhibitors of oxidative ATP production cause transient vasoconstriction and block subsequent pressor responses in rat lungs. Circ. Res. 48, 393–400. Stacpoole, P.W. (1989). The pharmacology of dichloroacetate. Metabolism 38, 1124–1144. Stacpoole, P.W., Nagaraja, N.V., Hutson, A.D. (2003). Efficacy of dichloroacetate as a lactate-lowering drug. J. Clin. Pharmacol. 43, 683–691.

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CHAPTER 6 Oxidative Modification of Ca2+ Channels, Ryanodine Receptors, and the Sarco/ Endoplasmic Reticulum Ca2+-ATPase Victor S. Sharov and Christian Schöneich Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, KS 66047, USA

I. Introduction II. Overview of Ca2+ Translocation Membrane Proteins III. Ca2+ Channels A. Voltage-gated Channels B. Ligand-gated Channels IV. SERCA V. PMCA VI. Concluding Remarks References

Abstract This paper reviews our current knowledge on the oxidative modifications of membrane proteins implicated in Ca2+ transport. The high sensitivity of some Ca2+ handling proteins, such as the L-type voltage-gated channel, the ryanodine receptor, and the sarco/endoplasmic reticulum Ca2+ -ATPase (SERCA) to oxidative modifications renders them important targets for oxidative regulation/modulation endowing a crosstalk between redox signaling and calcium transients in the cell. The main focus of this review is to correlate either loss or gain of protein function under conditions of oxidative stress to selective, potentially reversible, modifications of specific amino acids. Among those special emphasis will be placed on S-nitrosation, S-glutathiolation, disulfide formation, and Tyr nitration, as potential mechanism of redox-dependent signaling in the cell. Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00206-8

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I. INTRODUCTION Changes in subcellular, cellular or tissue Ca2+ levels have been reported for various conditions of oxidative stress (reviewed by Annunziato et al., 2002; Camello-Almaraz et al., 2006; Davidson and Duchen, 2006; Mattson, 2007; Beccafico et al., 2007; Hool, 2008). Such modulation of Ca2+ levels can be the result of activation/inactivation and/or post-translational modification of multiple proteins involved in the regulation of Ca2+ metabolism. Generally, conditions of oxidative stress are characterized by increased levels of some but not necessarily by all types of reactive oxygen species (ROS), which may include superoxide anion radical, peroxides, peroxynitrite, hypochlorous acid, and others. Each of these species will show distinct reactivities and selectivities in their reactions with given proteins, and frequently the actual parameters controlling these reactivities and selectivities are not completely understood. Fact is, however, that some of these ROS are involved in very selective reversible modifications of specific proteins, associated with a change in activity, so that these modifications can be considered regulatory in nature. In this paper, we will not provide an overview over all ROS and/or free radicals, which could potentially modify calcium channels and transporters (for reviews, see Stark, 2005; Camello-Almaraz et al., 2006; Hool and Corry, 2007). Rather, we will describe studies demonstrating either loss or gain of protein function when Ca2+ channels and transporters are exposed to specific oxidants and/or biologic conditions. Among these Ca2+ transporting proteins we will focus on the L-type voltagegated channel, the sarco/endoplasmic reticulum Ca2+ -ATPase (SERCA), and the plasma membrane Ca2+ -ATPase (PMCA), where specifically SERCA and PMCA are somehow underrepresented in recent reviews (Hool and Corry, 2007; Hidalgo and Donoso, 2008), which concentrate more on voltage- and ligand-gated Ca2+ channels. II. OVERVIEW OF Ca2+ TRANSLOCATION MEMBRANE PROTEINS To understand the role of redox regulation of Ca2+ channels and transporters (Davidson and Duchen, 2006; Hool and Corry, 2007; Mattson, 2007), we shall briefly classify the proteins involved in Ca2+ translocation through membranes, and their specific functions. Generally, cells maintain low cytoplasmic concentrations of Ca2+ (below 10−7 M) through ATP-dependent translocation of Ca2+ ions across the plasma membrane by PMCA and the Ca2+ /Na+ exchanger, and into intracellular stores by SERCA, the mitochondrial Ca2+ uniporter (MCU) or ion exchangers. Since extracellular Ca2+ concentrations are usually 3 to 4 orders of magnitude higher, the opening of plasma membrane Ca2+ channels can rapidly elevate intracellular calcium levels. Similarly, Ca2+ release channels located in the membranes of intracellular organelles can generate fast Ca2+ fluxes into the

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cytoplasm and into other cellular compartments. Often the influx of extracellular Ca2+ serves as an external signal triggering massive release of Ca2+ from intracellular stores, especially in excitable cells. The mitochondrial membrane harbors the MCU, a channel, which transports Ca2+ into mitochondria using an electrogenic membrane potential (Kirichok et al., 2004). Instead, the Na+ /Ca2+ exchanger releases Ca2+ from mitochondria through exchange with Na+ ions. Inside the cell, Ca2+ serves as a second messenger to regulate the activity of diverse proteins and cellular functions, such as cell signaling, gene expression, cell motility, apoptosis, etc.; in specific tissues, Ca2+ is involved in physiological processes such as muscle contraction, neurotransmitter secretion, neuron excitability, and action potential generation. Hence, the maintenance of Ca2+ homeostasis through the various calcium handling mechanisms is very important, specifically because high intracellular calcium concentrations are toxic and may lead to cell death via apoptotic and necrotic pathways (Waring, 2005). III. Ca2+ CHANNELS A. Voltage-gated Channels Ca2+ channels are classified as voltage-gated channels, associated with the plasma membrane, and ligand-gated channels, which reside in different types of membranes (Catterall et al., 2005; Hool and Corry, 2007). Voltage-gated channels are expressed in muscle, neuronal, endocrine, and receptor cells and further classified based on mechanisms of activation and inactivation as L-type (high voltage activated, long lasting, inhibited by dihydropyridines, phenylalkylamines, and benzothiazepines), and T-type (low voltage and transiently activated with only pacemaking function and unknown antagonists). Additionally, N-type, P/Q-type, and R-type channels have been identified in neurons based on specific functions and sensitivity to toxins, where N-type channels are sensitive to ω-conotoxinGVIA, P/Q-type channels are sensitive to ω-agatoxin IVA, and R-type channels are sensitive to SNX-482. Voltage-gated Ca2+ channels are implicated, among others, in excitation-contraction coupling, action potential generation, hormone and neurotransmitter release, pacemaking (repetitive fire activity), and sensing (Catterall et al., 2005). A large effort was placed on the evaluation of the role of nitric oxide (NO) on Ca2+ channel activity and intracellular Ca2+ levels. Experimentally, this was achieved through the use of NO synthase (NOS) inhibitors, NO scavengers and NO donors specifically in brain and muscle tissues and cultured cells. As both soluble guanylate cyclase (sGC) and S-nitrosation (cGMP-independent) pathways may be involved in the NO-dependent modulation of Ca2+ currents, specific inhibitors of both NO pathways were applied for further analysis. Early observations that NO inhibits an L-type Ca2+ current in carotid body sensory cells via a cGMP-independent mechanism were

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based on experiments with N-ethylmaleimide (NEM), which prevented the Snitrosation of calcium channels (Summers et al., 1999). Channel-type specificity was assessed by the influence of specific inhibitors on the effect of NO on the Ca2+ current. Recent data support a role of S-nitrosation in the inhibitory effect of NO on L-type Ca2+ channels, in addition to an sGC-dependent mechanism in vestibular hair cells (Almanza et al., 2007), cultured rat hippocampal neurons, and rat hippocampal slices in the CA1 region (Jian et al., 2007; Tjong et al., 2007). Chemical evidence for an S-nitrosation mechanism was obtained through the reversal of NO-dependent inhibition by ascorbate, which reduces S-nitrosothiols but not disulfides. Importantly, reactive molecules, such as, e.g., the oxidant hydrogen peroxide or the Michael acceptor 4-hydroxynonenal, modulated L-type Ca2+ channel activity in the opposite direction compared to NO, for example, selectively enhancing the nifedipine-sensitive Ca2+ current in dentate granule cells (Akaishi et al., 2004a, 2004b). Unfortunately, however, these conclusions are based on functional studies only and suffer from the lack of a direct demonstration of S-nitrosation of Ca2+ channels (it could well be that the observed Ca2+ levels are the result of modification of regulatory proteins upstream from the Ca2+ channels). No data are available on the structural and functional modification by either NO or other reactive species of Ca2+ channels other than L-type channels. An indirect mechanism of NO-dependent inhibition has been reported for N- and P/Q-types of Ca2+ channels in rat insulinoma and human neuroblastoma cells, likely mediated through sGC (Grassi et al., 1999; D’Ascenzo et al., 2002). Together, these data would suggest that L-type Ca2+ channels could be selectively targeted by reactive oxygen species and that their activity could be regulated through oxidation. In such case oxidative protein modification appears physiologically important instead of damaging. Interestingly, CA1 hippocampal neurons, which are specifically vulnerable to oxidative stress elicited by ischemiareperfusion, exhibited a selective downregulation of L-type calcium-channel activity after the ischemic insult, whereas non-vulnerable CA3 neurons did not (Li et al., 2007). This result could be interpreted such that physiologic oxidative regulation of a protein requires an environment, which tolerates periodic increased levels of ROS. This is the case for CA3 but not for CA1 neurons. Based on selective effects of hypoxia, thiol-reducing and alkylating reagents, and reactive oxygen and nitrogen species on L-type Ca2+ channels, a typespecific pore-forming α1 subunit of the channel was suggested as a target for oxidative modification and redox regulation implicated in aging and diseases (Annunziato et al., 2002; Waring, 2005; Zima and Blatter, 2006; Mattson, 2007; Hool, 2008). Here, Cys residues are the most probable oxidation sensitive sites on this protein subunit; for example, the human cardiac α1C subunit contains 43 Cys residues, and out of them, three are localized to the specific sequence 1786 RERHVPVCEDLELRRDSGSAGTQAHCLLLRRANPSRCHSR1825 , which is responsible for its sensitivity to hypoxia in cultured HEK cells (Fearon et al.,

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2000). Importantly, other Cys residues in the channel, e.g., those involved in the formation of inter-unit disulfide bridges may also contribute structurally to the overall calcium channel activity (Hool, 2008). In addition, the oxidation of Met residues may directly affect the structure of the Ca2+ channel, or indirectly affect the activity of the channel through modification of regulatory proteins, such as calmodulin (reviewed by Bigelow and Squier, 2005). A sequence-specific analysis of oxidative post-translational modifications in voltage-gated Ca2+ channels is still needed. However, one obstacle pertaining to the correlation of such modifications with structure is the lack of atomic resolution structural information about any Ca2+ channel; to date, most of our knowledge is based on the molecular structure and function of other voltage-gated channels, such as the potassium channel (Hool and Corry, 2007).

B. Ligand-gated Channels Three isoforms of the inositol triphosphate (IP3 ) receptor and the ryanodine receptor (RyR) represent the most common types of a ligand-gated calcium release channel. More recently, two additional members of this class of calcium-release channels have been described: (i) the inner nuclear membrane inositol tetrakisphosphate (IP4 ) receptor, which releases Ca2+ from the nuclear envelope lumen to the nucleoplasm (Mishra and Delivoria-Papadopoulos, 2002; Mishra et al., 2003), and (ii) the calcium release-activated current (CRAC) channel, which is located in the plasma membrane of non-excitable cells, such as T-lymphocytes, and which is dependent on extracellular calcium (Zweifach and Lewis, 1996; Zhang et al., 2005). Different types and isoforms of release channels for storeoperated calcium (SOC), which are activated at different calcium concentrations, are usually co-expressed and a combination of these receptors serves to maintain intracellular Ca2+ levels and regulate Ca2+ responses (Zheng et al., 2005; Hool and Corry, 2007). In addition to calcium, SOC release channels are sensitive to specific protein and low molecular weight ligands. Here, we shall briefly review data on the redox regulation of Ca2+ release channels. The RyR is probably one of the best characterized Ca2+ release channel with regard to redox regulation despite of its dimensions (a homotetrameric RyR contains over 20,000 amino acids with a total mass of ca. 2.3 MDa) and complete lack of atomic resolution structural information. Experimental evidence suggests that oxidative modifications of Cys are involved in the alteration of channel activity. Each monomer of RyR contains ca. 100 Cys residues, and approximately half of them are available for covalent modification (Dulhunty et al., 2000; Xia et al., 2004). The pharmacological effects of different classes of sulfhydryl-modifying reagents and oxidants demonstrate the existence of “hyperreactive” Cys residues on RyRs, which could play a role in the regulation of normal contractile function (for skeletal and cardiac muscle tissues) and rationalize alterations in excitation-contraction

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coupling during pathological conditions (Feng et al., 1999; Anzai et al., 2000; Goldhaber and Qayyum, 2000; Morad and Suzuki, 2000; Salama et al., 2000; Fabisiak et al., 2000; Pessah and Feng, 2000; Bidasee et al., 2003a, 2003b; Xia et al., 2000, 2004; Martinez-Burgos et al., 2006; Mochizuki et al., 2007; Cheong et al., 2005; Hidalgo, 2005). In fact, the term “redox sensor” has been applied to the RyR. This term has multiple meanings ranging from a simple correlation of a number of free Cys residues with Ca2+ permeability of the channel to a more specific susceptibility to reversible S-nitrosation, S-glutathiolation, and disulfide formation of selected Cys residues, that control channel activity depending on oxygen tension and/or overall cellular GSH/GSSG ratio (Xu et al., 1998; Eu et al., 1999; Meissner, 2004; Hidalgo and Donoso, 2008). Albeit physiological effects of thiol modifications may vary depending on cell and RyR type, and reagent concentrations, oxidizing conditions generally favor channel opening, whereas reducing conditions have the opposite effect. In addition to direct effects on the RyR properties, thiol modifications alter the channel response to other modifiers and physiological ligands, such as Ca2+ , Mg2+ , ATP, and cADP ribose, and several proteins, which modulate channel activity, such as protein kinases, CaM, junctophilin, and the immunophilin, FK506-binding protein (FKBP12), an essential component of RyR1 and RyR2 in muscle tissues (Zhang et al., 1999; Hamilton and Reid, 2000; Oba et al., 2002, 2008; Meissner, 2004; Aracena et al., 2005; Zissimopoulos and Lai, 2006; Camello-Almaraz et al., 2006; Bull et al., 2007; Phimister et al., 2007; Zissimopoulos et al., 2007). Again, potential oxidative modification of regulatory proteins, e.g., calmodulin, should be taken into consideration (Boschek et al., 2008). An interesting hypothesis suggests that all non-thiol channel modulators may affect RyR through a single mechanism, changing the reduction potential of the channel according to their electron donor/acceptor characteristics (Marinov et al., 2007). The identification of Cys residues responsible for the redox regulation of RyR was an open question until the recent development of sequence-specific mass spectrometry techniques. Based on limited proteolysis of skeletal muscle RyR1 and N-terminal amino acid sequencing, together with the analysis of the distribution of [3 H]NEM labeling, it was initially demonstrated that Cys3635 is rapidly labeled by NEM and that this labeling is blocked by association with calmodulin (Moore et al., 1999). The authors suggested that Cys3635 is located at an intersubunit contact site that is close to or within a calmodulin binding site. By means of cleavable isotopecoded affinity tag (ICAT) labeling of Cys residues, coupled to MALDI-TOF peptide fingerprinting, an additional 8 Cys sites were identified, which are endogenously modified in RyR1 (i.e., Cys at positions 36, 315, 811, 906, 1591, 2326, 2363, 3193, and 3635) and another three Cys residues, which could be modified only exogenously with redox-active reagents (Cys253, Cys1040, and Cys1303) (Aracena-Parks et al., 2006). Recently, combining fluorescent derivatization of Cys residues for quantitative analysis with tandem MS analysis of tryptic digests, two research groups successfully identified functionally important Cys residues

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(“hyperreactive cysteines”) that can be labeled by thiol-reactive probes under physiological conditions. Seven of them were identified as 7-diethylamino-3(4 -maleimidylphenyl)-4-methylcoumarin adducts (Cys1040, Cys1303, Cys2436, Cys2565, Cys2606, Cys2611, and Cys3635) after in-gel tryptic digestion and reversed-phase HPLC of the resulting peptides followed by MALDI-TOF/TOF and ESI-MS/MS analysis (Voss et al., 2004). The application of another Cys label, monobromobimane, coupled to MALDI-TOF and MALDI-TOF/TOF mass spectrometry analysis yielded only one redox-sensitive Cys residue, Cys2327 (Petrotchenko et al., 2006). These experimental differences may be the results of different analytical procedures and mass spectrometry equipment used in the latter studies. To date, the lack of 3-D atomic resolution structures of RyR does not permit any structure-function relationship analysis of the sequence-specific modification of redox-sensitive Cys residues. IV. SERCA The sarco/endoplasmic reticulum Ca-ATPase (SERCA) constitutes an essential Ca2+ -pump responsible for maintaining and/or restoring cytosolic Ca2+ levels during various physiological processes such as, e.g., muscle relaxation, signal transduction, and neuronal activity. SERCA is responsible for the transport of Ca2+ into the endoplasmic or sarcoplasmic reticulum (ER/SR) thereby maintaining high Ca2+ concentrations in the intracellular store, which is used for triggering muscle contraction, cell signaling and apoptosis (MacLennan et al., 1997; East, 2000; Dremina et al., 2004, 2006). The nomenclature of SERCA is based on three genes (SERCA1, SERCA2, SERCA3) and different isoforms, which are expressed in tissue-specific and developmentally regulated patterns (Bobe et al., 2004; Dode et al., 2002). The fast-twitch muscle isoform SERCA1 is restricted to fast-twitch skeletal muscle, while SERCA2a is expressed in cardiac and slow-twitch skeletal muscle. The isoforms SERCA2b and SERCA3a-3f are expressed ubiquitously in muscle and non-muscle cells. A special isoform of SERCA2, the secretory pathway Ca2+ /Mn2+ -ATPase 2 (SPCA) is a Golgilocalized pump with high affinity for Ca2+ ions presented in various types of secretory cells (Vanoevelen et al., 2005). Importantly, a reduced activity or expression of SERCA appears to correlate with various pathologies, including genetic diseases and cancer (MacLennan et al., 1997; Prasad et al., 2004). SERCA activity is modulated through specific post-translational modifications, such as oxidation and/or S-glutathiolation (Viner et al., 1999b, 2000; Adachi et al., 2004). SERCA isoforms differ in their sensitivity to oxidative modifications in vivo and in vitro. Earlier, we had demonstrated that biological aging leads to the accumulation of significant levels of 3-nitrotyrosine (3-NT) on the SERCA2a isoform, i.e., ca. 4 mol 3-NT/mol SERCA2a in skeletal muscle (Viner et al., 1996, 1999a), and ca. 3 mol 3-NT/mol SERCA2a in the heart (Knyushko et al., 2005). Such high yields of 3-NT on SERCA2a may not come as a surprise considering the

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close location of SERCA and nitric oxide synthase (nNOS) (Kobzik et al., 1994; Xu et al., 1999). What is surprising, though, is the apparent selectivity of nitration for SERCA2a even in fast-twitch skeletal muscle, which contains large quantities of the SERCA1 isoform that is ca. 84% homologous to SERCA2a. Mass spectrometry analysis revealed that in vivo aging leads to nitration of predominantly Tyr residues at positions 294 and 295 in rat skeletal muscle SERCA2a (Viner et al., 1999a). This sequence-specific information was utilized to generate sequence-specific antibodies against SERCA2a containing 3-NT at positions 294 and 295, which were then successfully applied for immunohistochemistry of SERCA2 nitration (Xu et al., 2006). Similar observations of a higher sensitivity of SERCA2 to modification by peroxynitrite in vitro (Grover et al., 2003; Kaplan et al., 2003) and in vivo (Li et al., 2006; Fugere et al., 2006) compared to other isoforms have been reported. SERCA2 activity is regulated in part through the NO-dependent reversible S-glutathiolation of Cys residues at positions 669 and/or 674 (Adachi et al., 2004). These sites were identified through a proteomic method, where cells were incubated with biotinylated glutathione prior to affinity enrichment of S-glutathiolated peptides. The redox state of specific SERCA Cys residues is important for enzymatic function, and modifications of different SERCA Cys residues may result in both inhibition and activation of the protein (Viner et al., 1999b, 2000; Adachi et al., 2004; Li and Camacho, 2004). Earlier studies on the role of Cys modifications in the regulation of SERCA activity were based on either selective labeling or site-directed mutagenesis of Cys residues that resulted in a significant change of protein activity. Though these approaches specifically addressed modifications of selected Cys residues, they are less relevant to oxidative modifications of SERCA in vivo. Besides, the oxidative modification of amino acids can lead to a variety of oxidation products, which may differentially affect protein structure and function. Proteolytic mapping of SERCA and HPLC-MS analysis detected several specific Cys-containing peptides with a mass increase fitting to Cys oxidation, Cys S-nitrosation and S-glutathiolation, and quantified the loss of some Cys residues upon oxidation in vitro using reactive thiol labeling (Viner et al., 2000; Adachi et al., 2004). However, all previous studies suffered from both incomplete Cys labeling in SERCA and the use of peptide masses only (MS1 mode) for the identification of labeled peptides. Moreover, these earlier studies did not use modification-resistant peptides as internal standards for digestion yields of Cys-containing peptides. Recently, we addressed these problems when labeling of SERCA1 within the sarcoplasmic reticulum (SR) preparation with ThioGlo1 (TG1=methyl-10-(2,5-dioxo-2,5-dihydro-1H-pyrrol-1-yl)-9-methoxy3-oxo-3H-benzo[f]chromene-2-carboxylate) in 2% SDS resulted in a virtually complete derivatization of reduced Cys residues of SERCA1 (Sharov et al., 2006). In-gel processing (reduction, alkylation and digestion) of the protein allowed the recovery of most Cys-containing peptides, and of additional peptides, which were used to normalize the yields of the labeled peptides by quantitative HPLC-ESI-

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TABLE 1 Content of ThioGlo1-labeled Cys residues in SERCA1 after the exposure of rat skeletal muscle sarcoplasmic reticulum (SR) to different concentrations of peroxynitrite or tryptophan peroxide in vitro, and in the SR isolated from 34-months-old rats relative to control, untreated 6 months old animals. Total loss of Cys residues per SERCA monomer is calculated as sum of losses of individual Cys residues (Sharov et al., 2006; Dremina et al., 2007) Cys sequence position 525 561 636 471 674,675 498 614 364 12 377 268 417,420 938 774 Total loss

TG1-labeled Cys content in SERCA1 (ratio to control) Peroxynitrite (mM) 0.3

1

3

0.1

DL-Trp peroxide (mM) 0.2

Aging (34 months)

0.66 ± 0.13∗ – – – – – – – – – – – – – 0.34

0.74 ± 0.07∗ – – – – 0.77 ± 0.08∗ – 0.65 ± 0.03∗ – – – 0.55 ± 0.05∗∗ – – 1.29

0.69 ± 0.07∗ – – – 0.63 ± 0.05∗ 0.56 ± 0.04∗ – 0.41 ± 0.07∗∗ – – – 0.49 ± 0.01∗∗ 0.35 ± 0.10∗∗ – 2.87

– – – – 0.92 ± 0.05∗ – – – 0.81 ± 0.15∗ – 0.89 ± 0.10∗ 0.83 ± 0.10∗ 0.66 ± 0.29∗ 0.33 ± 0.14∗∗ 1.56

– – – – 0.73 ± 0.11∗∗ – – – 0.66 ± 0.12∗∗ 0.64 ± 0.14∗∗ 0.60 ± 0.21∗∗ 0.67 ± 0.14∗∗ 0.39 ± 0.15∗∗ 0.29 ± 0.15∗∗ 3.02

0.73 ± 0.12∗ 0.55 ± 0.16∗∗ 0.65 ± 0.10∗∗ 0.60 ± 0.20∗∗ 0.69 ± 0.27∗ 0.54 ± 0.07∗∗ – 0.61 ± 0.11∗∗ – – 0.63 ± 0.21∗∗ 0.70 ± 0.22∗ 3.30

∗ Statistically significant results with P > 0.90. ∗∗ Statistically significant results with P > 0.95.

tandem MS. The quantitative analysis of the TG1-labeled sequences provides a differential display of 16 out of 20 reduced Cys residues in SERCA1 available for oxidative modification. Importantly, quantification of total Cys losses by HPLC-MS analysis of SERCA1 digests is in agreement with the data obtained by fluorescence spectroscopy of the intact TG1-labeled protein, validating the approach and permitting the analysis of a role of specific Cys residues in the control of SERCA activity. In vitro, the incubation of SR with different oxidants led to the inactivation of the ATPase function with concomitant loss of TG1-reactive Cys residues. Peroxynitrite, a biologically relevant oxidant, targeted specific SERCA Cys residues at position 364, 417, 420, 498, 525, 674, 675, and 938 (Table 1). A comparative analysis shows that not all the oxidation-sensitive Cys residues are equally important for SERCA function. Peroxynitrite concentration-dependent loss of individual Cys residues coincided with the loss of SERCA activity only for the residues at positions 674/675 and 938 (Sharov et al., 2006). No other Cys residue shows the characteristic lag phase correlating with protein inactivation. It is likely that some of the earlier targeted Cys residues, i.e., at positions 525, 498,

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417, 420, and 364 actually protect functionally important Cys residues from oxidation and could be considered intramolecular antioxidants similar to the function ascribed to protein methionine residues (Levine et al., 1996). Based on peroxynitrite data, we cannot state whether the oxidation of individual Cys674, Cys675 or Cys938 residues alone serve as a “redox switch” for SERCA1 activity, or the loss of the protein activity is due to a combination of multiple Cys oxidation events and/or additional modifications such as Met oxidation and Tyr nitration (Alvarez and Radi, 2003). Considering the importance of selected Cys residues for the NO-dependent activation of SERCA, the sensitivity of SERCA against amino acid peroxides was investigated (Dremina et al., 2007). Such amino acid peroxides form under conditions of oxidative stress and are ca. 100-fold more reactive towards SERCA compared to hydrogen peroxide. Out of 16 SERCA1 Cys residues quantifiable through TG1-labeling, 9 are reactive towards peroxides derived from Trp (Table 1). Among those Cys residues, Cys774 and Cys938 were most reactive, both located in the transmembrane domain of SERCA. A lower reactivity was observed for Cys residues present at positions 12, 268, 377, 417, 420, 674, and 675. The rather high reactivity of the Trp-derived peroxides towards Cys residues located in the transmembrane domain (Cys268, Cys774 and Cys938) is remarkable. The crystal structures of SERCA1 in the absence and presence of Ca2+ (Toyoshima et al., 2000; Toyoshima and Nomura, 2002) show that Cys938 is more exposed on the surface of the protein compared to Cys774 and Cys268, which rather point towards the interior of the transmembrane domain (Figure 1). Hence, the observed reactivity of Trp-derived peroxides with Cys residues of the transmembrane domain is not consistent with their surface exposure. On the other hand, specifically the more buried Cys residues may participate in hydrogen bonding, resulting in partial negatively charged sulfur, which should be easier to oxidize. Interestingly, Cys938, but not Cys774, represents also a target for the water-soluble anion peroxynitrite (Table 1). With such a manifold of targets it is difficult to correlate the oxidation of one specific Cys residue with SERCA inactivation: it is likely, that oxidation of each of the individual Cys residues is associated with a degree of inactivation, and that the measured SERCA inactivation represents the sum of all the partial inactivations caused by the oxidation of individual Cys residues. In addition, at least 4 Cys residues could not be quantified by TG1-labeling and oxidation of the latter may contribute to the overall SERCA inactivation (Li and Camacho, 2004). Biological aging leads to a ca. 40% loss of SERCA1 activity in rat skeletal muscle in vivo (Sharov et al., 2006), and a molecular rationale for this phenomenon could be the age-dependent oxidation of specific Cys residues by peroxynitrite, amino acid peroxides, and/or other endogenous oxidants. Interestingly, the affected residues in vivo do not completely match those targeted by peroxynitrite and Trp peroxide in vitro (Table 1 and Figure 1). Both peroxynitrite in vitro and aging in vivo target Cys residues at positions 498, 525, 674/675 and 938, whereas

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FIGURE 1 Crystal structure of SERCA1 (PDB: 1su4) showing the location of Cys residues sensitive to oxidation by Trp peroxide and peroxynitrite in vitro and to in vivo aging in rat skeletal muscle SR. Cys residues are represented in the spacefill view using elemental (CPK) color palette, Ca2+ ions attached to the high-affinity binding sites in the transmembrane calcium translocation channel are shown as blue balls, and the phosphorylated aspartic acid residue (Asp351∼P) in the active site is shown as a spacefill residue in magenta. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this book.)

Trp peroxide in vitro and aging in vivo target Cys at positions 377, 674/675, 774, and 938. Hence, if we suggest an existence of specific Cys residue(s), which is(are) required in a reduced state for an active conformation and unrestricted

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ATPase activity of SERCA1, Cys residues at positions 674/675 and 938 represent the most probable candidates. Aging additionally affects Cys at positions 561 and 614, while Trp peroxide additionally targets Cys12, Cys268, Cys344 and Cys349, and peroxynitrite additionally targets Cys364. The Cys oxidation pattern observed as a result of aging likely reflects the occurrence and reaction of additional oxidants in vivo, which target, for example, Cys561 and Cys614. Also, the conformation of SERCA in vitro may not be completely identical with that in vivo, a feature, which may lead to some differential reaction kinetics of Cys residues. Moreover, not only the loss of a specific Cys residue, but also the nature of its specific Cys oxidation product will determine its ultimate effect on protein function. At present, we cannot definitely conclude, which Cys residue is more important for the loss of SERCA1 ATPase activity observed in aging rat skeletal muscle. A notable observation is the oxidative modification of Cys674/675 by different oxidants in vitro and in vivo, given that in smooth muscle SERCA2 (which shares more than 80% homology with SERCA1) Cys674 is reversibly modified through S-glutathiolation induced by NO, resulting in SERCA activation, whereas removal of this Cys residue results in a loss of NO-dependent activation (Adachi et al., 2004). Ligand binding studies do not show significant changes in either the Ca2+ or ATP affinity of SERCA1 in aging SR, suggesting that the effect of Cys oxidation on Ca2+ -ATPase function may not be due to modification of specific ligand binding sites on the protein but, rather, through more general effects on protein structure, potentially uncoupling ATP hydrolysis from Ca2+ translocation. In contrast, the inactivation of SERCA by peroxynitrite in vitro is accompanied by a decrease in Ca2+ affinity (Sharov et al., 2006). These data, again, demonstrate that SERCA Cys modification in vitro does not completely simulate the age-dependent modification.

V. PMCA Since observations of an age-associated decrease in PMCA activity in the rat brain (Qin et al., 1998; Zaidi et al., 1998), attempts have been made to link the elevated Ca2+ levels in aging neurons with oxidative modifications of the plasma membrane Ca2+ -ATPase. The sensitivity of PMCA to oxidation in vitro was tested in synaptic plasma membranes exposed to azo-initiators, hydrogen peroxide, and peroxynitrite. All these reactive species markedly changed the kinetic parameters of PMCA enzymatic activity and caused aggregation of the protein in a concentration-dependent manner, suggesting that this protein may represent a target to oxidative stress in aging brain (Zaidi and Michaelis, 1999). Analysis of oxidative post-translational modifications in purified PMCA exposed to hydrogen peroxide, which caused a ca. 50% inactivation of the enzyme and significant structural alterations (partially reversible by DTT), did not reveal significant changes

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in amino acid composition, except for a loss of ca. 2.5 mol Cys/mol protein (Zaidi et al., 2003). In addition, a protective effect of calmodulin bound to PMCA before the exposure to oxidants was documented. The latter effect can be attributed to either a structural change in PMCA upon calmodulin binding that renders PMCA less oxidation-sensitive (Lushington et al., 2005), or an “antioxidant” effect of calmodulin, which contains highly oxidizible Met residues, particularly the C-terminal Met144 and Met145 (reviewed by Bigelow and Squier, 2005). Conformations of a PMCA-calmodulin complex were probed by single-molecule polarization modulation spectrometry and it was found that oxidative modification of either PMCA or CaM in the complex results in an increased coupling of an autoinhibitory/CaM binding domain to the ATP-binding core of PMCA, thereby stabilizing the enzyme in an inactive conformation (Osborn et al., 2004, 2005). Specific Cys targets for PMCA oxidative modifications are still to be identified.

VI. CONCLUDING REMARKS The high sensitivity of some Ca2+ handling proteins, such as the L-type voltage-gated channel, the ryanodine receptor, and SERCA renders them important targets for oxidative regulation/modulation endowing a crosstalk between redox signaling and calcium transients in the cell. With the development of sensitive mass spectrometry and proteomic methodologies, there is no doubt that specific sequences targeted by oxidation in vivo may soon be identified. Acknowledgements This work was supported by the NIH (AG12993, HL31607).

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CHAPTER 7 Regulation of Na,K-ATPase by Reactive Oxygen Species Guofei Zhou, Laura A. Dada and Jacob I. Sznajder Division of Pulmonary and Critical Care Medicine, Feinberg School of Medicine, Northwestern University, 240 E. Huron, McGaw Pavilion M-326, Chicago, IL 60611, USA

I. Na,K-ATPase II. Na,K-ATPase in Alveolar Fluid Reabsorption III. Role of Reactive Oxygen Species in Signaling A. Mitochondrial ROS B. NADPH Oxidase ROS Production C. ROS as Signaling Molecules IV. Regulation of Na,K-ATPase and Alveolar Fluid Reabsorption by ROS A. Hypoxia B. Hyperoxia C. Other Stimuli V. Dopamine and β-adrenergic Agonists Improve ROS-Mediated Decrease in Alveolar Fluid Reabsorption VI. Summary References

Abstract Pulmonary edema occurs in patients with acute respiratory distress syndrome (ARDS) and congestive heart failure. Effective edema clearance is critical for these patients to survive. The edema clearance is carried out mostly by the coordination of the apical Na+ channels and the basolateral Na,K-ATPase effecting active Na+ transport. Downregulation of Na+ transport occurs frequently in ARDS patients and results in impaired edema clearance. Hypoxia, hyperoxia, particulate matter and thrombin inhibit edema clearance. Reactive oxygen species (ROS) generated from the electron transport chain in the mitochondria and/or NADPH oxidase appears to be critical for the impaired edema clearance under these stress conditions. During hypoxia, ROS activate PKCζ , which in turn phosphorylates Na,K-ATPase α1 subunit at Ser-18, leading to the endocytosis of Na,K-ATPase from the plasma membrane. The endocytosis of Na,K-ATPase results in decreased Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00207-X

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Na,K-ATPase activity and can be prevented by antioxidants. We propose that ROS act as signaling molecules regulating the activity of Na,K-ATPase, which is important for alveolar fluid reabsorption. Further studies into the mechanisms by which Na,K-ATPase is regulated will be of importance for development of novel strategies for the treatment of edema patients.

I. NA,K-ATPase The Na,K-ATPase is a membrane-bound protein which belongs to the P-type ATPase protein family, where a phosphorylated enzymatic intermediate is formed during the catalytic cycle (Kaplan, 2002). Plasma membrane Na,K-ATPase utilizes the energy released from ATP hydrolysis to pump 3 Na+ out of cells and exchange 2 K+ into cells simultaneously, generating the Na+ and K+ gradients across the plasma membrane (Blanco and Mercer, 1998). These gradients are crucial to maintain cell volume, membrane potential and to provide a secondary energy source for nutrient uptake, maintaining intracellular pH and Ca2+ concentration. Na,K-ATPase is a heterodimer consisting of α and β subunits, and both are necessary for its activity. It is believed that α and β subunits are synthesized independently and then assembled into a dimer in the endoplasmic reticulum and delivered to the plasma membrane (Therien and Blostein, 2000; Kaplan, 2002). The α subunit is a transmembrane protein that catalyzes ATP hydrolysis and contains the binding sites for Na+ , K+ and its inhibitor ouabain (Blanco and Mercer, 1998). Four α isoforms, α1 to α4 , have been described (Kaplan, 2002). These isoforms are highly conserved with more than 77% identical primary amino acid sequences between each other. The α1 is the main isoform found in most tissues and other isoforms are tissue-specific (Blanco and Mercer, 1998). The β subunit has four isoforms and contains glycosylation sites. It controls the heterodimer assembly and insertion into the plasma membrane. Mutation of Na,K-ATPase has been linked to human diseases. The loss-offunction mutations of α2 subunit have been identified to be associated with familial hemiplegic migraine type 2 (De Fusco et al., 2003; Vanmolkot et al., 2003; Segall et al., 2004). Missense mutation of α3 subunit causes dystoniaParkinsonism (de Carvalho Aguiar et al., 2004). Mice with deletion of either the α1 or α2 isoform fail to survive, suggesting their fundamental roles in organ development (Moseley et al., 2003; Barcroft et al., 2004). The Na,K-ATPase activity can be regulated by changes in affinity for its substrates. However, more recent reports have suggested that the Na,K-ATPase activity is regulated mostly by its trafficking and insertion into the plasma membrane (Bertorello and Sznajder, 2005). The biological function of Na,K-ATPase has been studied in the brain, heart, kidney and lung (Lichtstein and Rosen, 2001; Schwinger et al., 2003; Bertorello and Sznajder, 2005). In the lung alveolar epithelium, the best studied function of

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the Na,K-ATPase is its role in alveolar fluid reabsorption (Mutlu and Sznajder, 2005).

II. NA,K-ATPase IN ALVEOLAR FLUID REABSORPTION Under physiological conditions there is a very thin layer of fluid lining the alveolar epithelium which allows for maintenance of surface tension, basic host defense properties, and gas exchange (Ware and Matthay, 2001). Patients with acute respiratory distress syndrome (ARDS) or congestive heart failure develop pulmonary edema as a consequence of increased permeability across the alveolarcapillary barrier (Fromm Jr. et al., 1995; Ware and Matthay, 2000). The accumulation of edema fluid causes a life-threatening impairment of gas exchange; therefore, the edema needs to be cleared for these patients to survive (Sznajder, 2001; Ware and Matthay, 2001). The edema is cleared by alveolar fluid reabsorption through active Na+ transport across the alveolar epithelium. Sodium is taken in on the apical surface of alveolar epithelial cells, partly through amiloride-sensitive Na+ channels. Subsequently, Na+ is actively extruded from the basolateral surface into the lung interstitium by the Na,K-ATPase, generating a transepithelial osmotic gradient. Then water follows the osmotic gradient created by the active Na+ transport into the interstitium space and the pulmonary circulation, leading to the reabsorption of alveolar fluid (Figure 1) (Matalon et al., 2002; Sznajder et al., 2002). The alveolar epithelium in the adult human lung consists of two morphologically distinct epithelial cells, type I (ATI) and type II (ATII) cells. Although the numbers of ATI and ATII cells are similar, about 95% of surface area is covered by ATI cells. Both ATI and ATII cells express Na,K-ATPase

FIGURE 1 Schematic representation of alveolar fluid reabsorption.

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and contribute to active Na+ transport (Figure 1) (Borok and Verkman, 2002; Johnson et al., 2002, 2006; Ridge et al., 2003). The importance of the Na,K-ATPase in edema clearance is highlighted by experiments where overexpression of Na,K-ATPase β1 subunit in rat lungs results in a significant increase in alveolar fluid reabsorption (Factor et al., 1998, 1999; Machado-Aranda et al., 2005). Importantly, gene transfer of Na,K-ATPase can improve alveolar fluid reabsorption in lung injury models (Azzam et al., 2002; Factor et al., 2002). These studies suggest the potential therapeutic application of Na,K-ATPase gene transfer in treatment of ARDS patients (Adir et al., 2003; Mutlu et al., 2004).

III. ROLE OF REACTIVE OXYGEN SPECIES IN SIGNALING Reactive oxygen species (ROS) production has been associated with a number of systems, including the mitochondrial electron transport chain, NADPH oxidase, xanthine oxidase, glucose oxidase and lipooxygenase (Finkel, 1998; Thannickal and Fanburg, 2000). Accumulating evidence suggests that the primary ROS sources are the mitochondria and NADPH oxidase (Figure 2) (Hancock et al., 2001; Chandel and Budinger, 2007).

FIGURE 2 The generation of reactive oxygen species in cells by mitochondrial electron transport chain and NADPH oxidase.

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A. Mitochondrial ROS In eukaryotes it has been estimated that 2–3% of the O2 consumed by mitochondria is incompletely reduced, yielding ROS (Chandel and Schumacker, 2000). Capture of an electron by O2 generates superoxide, which is then converted to hydrogen peroxide by superoxide dismutase (SOD) (Chandel and Schumacker, 2000). Superoxide can potentially be generated at complex I, complex III, and other electron transfer proteins (Figure 2). In complex III, superoxide is generated from the radical ubisemiquinone during the Q cycle, at two different sites, the Qi and Qo sites, where superoxide is released into the matrix and the intermembrane space, respectively. ROS do not appear to be generated by cytochrome oxidase itself (Guzy et al., 2005; Guzy and Schumacker, 2006).

B. NADPH Oxidase ROS Production The NADPH oxidase was initially discovered in neutrophils, serving as a host defense system against invading microorganisms (Bokoch and Knaus, 2003). The NADPH oxidase is a multi-protein complex consisting of membrane components and cytosolic components. In resting cells, p22-phox and gp91-phox form a heterodimer termed flavocytochrome b558 at the plasma membrane; upon stimulation, the cytosolic components (p40-phox, p47-phox and p67-phox) translocate to the plasma membrane and assemble the active enzyme with flavocytochrome b558 (Hancock et al., 2001). Flavocytochrome b558 contains an FAD group and two heme groups to enable the transfer of an electron from NADPH to oxygen, producing a superoxide anion, which is converted to hydrogen peroxide (Figure 2) (Herkert et al., 2004). Recent evidence suggests that the NADPH oxidase is ubiquitously expressed in mammalian plasma membrane. A family of gp91-phox-like proteins termed nonphagocytic NADPH oxidase proteins (NOX1 through NOX5) has been identified and p47-phox and p22-phox were also discovered in nonphagocytic mammalian cells (Bokoch and Knaus, 2003; Takeya and Sumimoto, 2006). The fact that nonphagocytic NADPH oxidases produce relatively lower amount of ROS than the phagocytic NADPH oxidase suggests that ROS may play an important role as signaling molecules rather than act as toxic agents (Gabbita et al., 2000; Thannickal, 2003).

C. ROS as Signaling Molecules ROS are increasingly considered as second messengers in mediating signal transduction when produced in low concentration (Thannickal, 2003). In many

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ways, ROS are ideal as signaling molecules. They are small, ubiquitous and diffusive. They are produced in multiple sites and their production is regulated by multiple mechanisms (Gabbita et al., 2000). They can be inactivated by catalases, peroxidases and other mechanisms. A large number of signaling pathways appear to be regulated by ROS, including receptor tyrosine kinases, protein kinase C (PKC), Src kinase, Janus kinase (JAK) and mitogen-activated protein kinases (MAPK) (Thannickal and Fanburg, 2000). Growth factors induce dimerization or oligomerization of their cognate receptors, leading to autophosphorylation in their cytoplasmic kinase domain and activation. This effect occurs in parallel with ROS production upon ligand binding and appears to be mediated by ROS (Thannickal and Fanburg, 2000). Similarly, exogenous H2 O2 has been shown to induce similar effects on phosphorylation and activation of the platelet-derived growth factor (PDGF) and epidermal growth factor receptors (Rhee et al., 2000). H2 O2 induces tyrosine phosphorylation of various isoforms of PKC (α, βI , γ , δ, ε and ζ ), leading to its activation even in the absence of receptor-mediated stimulation of phospholipase C (Konishi et al., 1997; Sun et al., 2000). Activation of the JAK-STAT pathway by PDGF appears to be redox sensitive (Simon et al., 1998). In fibroblast cells, H2 O2 -induced activation of p21Ras requires activation of Src kinase and JAK as well as recruitment of phosphatidylinositol 3 -kinase to the plasma membrane, where it interacts directly with p21Ras (Deora et al., 1998). Also, ROS activate ERK, which participates in Src kinases and p21Ras activation (Muller et al., 1997; Aikawa et al., 1997). ROS activate apoptosis signal-regulating kinase 1 (ASK1), resulting in activation of p38 and JNK (Zhang et al., 2003; Zhou et al., 2004). ROS activate signaling mainly through two pathways. On one hand, ROS alter the intracellular redox state. The cytosol normally maintains its redox state by the buffering capacity of intracellular thiols, primarily glutathione (GSH) and thioredoxin (TRX). Reduced GSH and TRX are maintained by their respective reductases. Both thiols can counteract intracellular oxidative stress by reducing H2 O2 (Thannickal and Fanburg, 2000). On the other hand, ROS mediate signaling via direct modification of cysteine residues in target proteins, altering their structure and function (Rhee et al., 2000).

IV. REGULATION OF NA,K-ATPase AND ALVEOLAR FLUID REABSORPTION BY ROS A. Hypoxia Alveolar hypoxia may occur during acute lung injury as a consequence of alveolar flooding, impairing oxygen transfer from the airspace into the pulmonary circulation (Ware and Matthay, 2000, 2001). Hypoxia has been shown to impair alveolar fluid clearance by inhibiting transepithelial active Na+ transport

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FIGURE 3 Time- and dose-dependent generation of mitochondrial ROS during hypoxia. Reproduced with permission from Chandel et al. (1998). Mitochondrial reactive oxygen species trigger hypoxia-induced transcription. Proc. Natl. Acad. Sci. USA 95 (20), 11715–11720.

(Vivona et al., 2001; Litvan et al., 2006). It has been demonstrated that exposure of rats to moderate hypoxia results in a significant decrease in alveolar fluid clearance, which was associated with a decrease in ENaC activity as well as a decrease in Na,K-ATPase activity due to decreased protein abundance at the basolateral membrane (Tomlinson et al., 1999; Carpenter et al., 2003; Litvan et al., 2006). In cultured alveolar epithelial cells, hypoxia-mediated downregulation of Na,KATPase is time- and O2 concentration-dependent (Planes et al., 1996; Wodopia et al., 2000; Dada et al., 2003). Short term exposure to hypoxia decreases Na,KATPase activity and the amount of plasma membrane Na,K-ATPase α1 subunit without significant change in its total amount, suggesting the endocytosis of Na,K-ATPase during hypoxia (Dada et al., 2003). Exposure of cells and tissues to hypoxia (pO2 = 5–50 Torr) increases the production of mitochondrial ROS in a dose-dependent manner (Figure 3) (Chandel et al., 1998), while cellular respiration does not become limited until the pO2 falls below 5–7 Torr. The current reasoning is that hypoxia does not change the apparent Km of the cytochrome oxidase, but reversibly decreases its Vmax (Chandel and Schumacker, 2000). This decrease in Vmax is not sufficient to limit normal respiration but affects the mitochondrial redox state by increasing the reduction state of cytochrome c (the electron donor to cytochrome oxidase), resulting in the ROS leaking from the mitochondria (Chandel and Schumacker, 2000). Accordingly, alveolar epithelial cells treated with mitochondrial inhibitors such as

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antimycin A, which inhibit sites distal to complex III, increase ROS production during normoxia or hypoxia (Dada et al., 2003). In contrast, inhibitors such as rotenone, diphenyleneiodonium chloride, which inhibit sites more proximal to complex III, inhibit ROS generation by blocking the flux of electrons before complex III (Guzy and Schumacker, 2006). The role of mitochondrial ROS during hypoxia is also supported by experiments carried out in mitochondrial DNA deficient cells (ρ 0 cells) (King and Attardi, 1989). The mammalian mitochondrial DNA encodes 13 polypeptides which are critical catalytic subunits for complex I, complex III, complex IV and the F1F0 ATP synthase (Anderson et al., 1981). These cells lack a competent electron transport chain and are incapable of generating ROS during hypoxia (Chandel and Schumacker, 1999). Treatment of cells with H2 O2 caused the endocytosis of plasma membrane Na,K-ATPase, and thus a decrease in Na,K-ATPase activity (Dada et al., 2003). Interestingly, ROS scavengers prevent the hypoxia-induced decrease in Na,KATPase activity and its protein abundance (Dada et al., 2003). The source of ROS in this process was further assessed in ρ 0 -A549 cells, where hypoxia failed to induce Na,K-ATPase endocytosis and decrease Na,K-ATPase activity. In animal studies overexpression of manganese SOD inhibited mitochondrial ROS production and blocked the decrease of Na,K-ATPase activity and alveolar fluid reabsorption during hypoxia (Litvan et al., 2006). Moreover, H2 O2 induced Na,KATPase endocytosis in ρ 0 -A549 cells, suggesting ROS act as messengers downstream of mitochondrion. Together, these studies suggest a role of mitochondrial ROS in hypoxia-mediated endocytosis of Na,K-ATPase. The endocytosis of Na,K-ATPase during hypoxia is triggered by the phosphorylation of its α1 subunit by PKCζ , which is activated by ROS (Dada et al., 2003). Endocytosis of Na,K-ATPase was reported to be clathrin dependent (Khundmiri et al., 2004). In lung alveolar epithelial cells, hypoxia-induced endocytosis of Na,KATPase requires the binding of adaptor protein AP-2 to the tyrosine-based motif (Tyr-537) located in the Na,K-ATPase α1 subunit, leading to the incorporation of Na,K-ATPase into clathrin vesicles. This process again requires ROS signaling (Figure 4) (Chen et al., 2006). Trafficking of clathrin vesicles requires the actin cytoskeleton in mammalian cells. The activation of Rho proteins leads to rearrangement of actin cytoskeleton, thus regulating endocytic process (Ridley, 2006). Most recently, it has been suggested that during hypoxia, mitochondrial ROS promote formation of actin stress fibers in alveolar epithelial cells by activation of the small GTPase RhoA, contributing to hypoxia-induced endocytosis of Na,K-ATPase (Dada et al., 2004). Prolonged hypoxia decreases the total Na,K-ATPase and plasma membrane Na,K-ATPase (Comellas et al., 2006; Litvan et al., 2006). Treatment of alveolar cells with the EUK-134 (combined SOD/catalase mimetic) or overexpression of glutathione peroxidase prevented the hypoxia-mediated Na,K-ATPase degradation. Furthermore, hypoxia-mediated Na,K-ATPase degradation was prevented in

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FIGURE 4 Schematic representation of endocytosis of plasma membrane Na,K-ATPase.

ρ 0 cells and cells with suppression of Rieske iron sulfur protein (which prevents the generation of ROS from the ubisemiquinone at Qi and Qo sites), suggesting that mitochondrial ROS are critical for Na,K-ATPase degradation (Comellas et al., 2006). Both lysosome and proteosome inhibitors prevented the degradation of Na,K-ATPase (Comellas et al., 2006), yet the exact location of the plasma membrane Na,K-ATPase degradation during hypoxia has not been determined. B. Hyperoxia Adult rats exposed to 100% oxygen develop a lethal lung injury (Factor et al., 2000; He et al., 2005). The alveolar epithelium becomes more permeable to solutes and alveolar edema accumulates (Olivera et al., 1995; Carvalho et al., 1998). Moreover, there is an impairment in active Na+ transport and the ability to clear edema, associated with a parallel decrease in Na,K-ATPase activity in ATII cells isolated from rats exposed to 95% O2 (Olivera et al., 1995). Cells exposed to hyperoxia have a decrease in the plasma membrane Na, K-ATPase abundance (Hawkins et al., 2006), probably related to the increased ROS production. Several studies suggest that hyperoxia can stimulate superoxide production from the mitochondria (Freeman and Crapo, 1981; Li et al., 2004; Xu et al., 2006). However, other groups of investigators have reported NADPH oxidase as the primary source of ROS in hyperoxic environments (Buccellato et al., 2004; Parinandi et al., 2003; Brueckl et al., 2006; Thannickal, 2003). Parinandi et al. reported that kinase activation was required for the assembly of the NADPH oxidase to generate the bulk of ROS during hyperoxia (Parinandi et al., 2003). These

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studies suggest that the mitochondrial ROS may act as a trigger to initiate the signaling to assemble the NADPH oxidase, which then amplifies the ROS signal (Parinandi et al., 2003).

C. Other Stimuli Particulate matter contributes to human diseases, especially pulmonary and cardiovascular diseases (Nel, 2005; Cascio, 2005; Brunekreef and Holgate, 2002). In the lung, long term exposure to particulate matter has been linked to obstructive lung diseases and cancer (Pope 3rd et al., 2002). However, the mechanisms involved are not clear. ROS have been implicated to play a role in the development of these diseases caused by exposure to particulate matter (Li et al., 2003). Recently, Mutlu and coworkers reported that exposure to particulate matter (diameter less than 10 µm) significantly decreases alveolar fluid reabsorption in mice as well as Na,K-ATPase activity and its abundance at the plasma membrane in alveolar epithelial cells (Mutlu et al., 2006). Pretreatment with the antioxidant EUK-134 rescued the decreased Na,K-ATPase due to particulate matter, suggesting a role for ROS (Mutlu et al., 2006). Patients with acute lung injury may have elevated concentrations of thrombin, a coagulation protease, in plasma and bronchoalveolar lavage fluids (Idell, 2003). An increased amount of thrombin is frequently associated with elevated ROS levels in endothelial cells and platelets (Herkert et al., 2004). Intravascular thrombin inhibits active Na+ transport in isolated-perfused rabbit lungs, leading to decreased alveolar fluid clearance (Vadasz et al., 2005). Thrombin downregulates plasma membrane Na,K-ATPase protein abundance and activity without affecting the amount of ENaC α and β subunits. These thrombin-induced effects can be prevented by inhibition of ROS production from NADPH oxidase, but not from mitochondria (Vadasz et al., 2005).

V. DOPAMINE AND β-ADRENERGIC AGONISTS IMPROVE ROS-MEDIATED DECREASE IN ALVEOLAR FLUID REABSORPTION The impaired alveolar fluid reabsorption by hypoxia and hyperoxia can be reversed by treatment with dopamine (DA) and the β-adrenergic agonists (terbutaline and isoproterenol (ISO)) (Vivona et al., 2001; Litvan et al., 2006; Saldias et al., 1999a, 1999b). The improvement of alveolar fluid reabsorption by DA and ISO is associated with parallel increases in Na,K-ATPase protein abundance at the basolateral membrane (Ridge et al., 2002; Litvan et al., 2006). This increase in plasma membrane Na,K-ATPase abundance is mediated by the exocytosis of Na,K-ATPase from late endosomal compartments into the basolateral membrane (Pesce et al., 2000; Ridge et al., 2002; Lecuona et al., 2003).

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VI. SUMMARY Alveolar fluid reabsorption is critical for patients with ARDS or congestive heart failure to have better outcomes. Alveolar fluid reabsorption is effected by Na,K-ATPase and Na+ channels. Hypoxia, hyperoxia, particulate matter and thrombin impair alveolar fluid reabsorption via ROS-mediated downregulation of Na,K-ATPase activity in the alveolar epithelium. More recent studies suggest that mitochondria- and/or NADPH oxidase-derived ROS are the common mediator of Na,K-ATPase downregulation in these conditions. ROS activates PKCζ , which phosphorylates Na,K-ATPase at Ser-18, leading to its endocytosis from the plasma membrane of alveolar epithelial cells. Antioxidants prevent endocytosis of Na,K-ATPase, resulting in restoration of the basal level of alveolar fluid reabsorption. Dopamine and isoproterenol can improve the rate of alveolar fluid reabsorption inhibited by hypoxia or hyperoxia, suggesting their potential as therapeutic drugs. However, further studies into the mechanisms by which Na,K-ATPase is regulated are warranted to provide insights for the design of novel therapeutic agents. Acknowledgements This work was supported in part by grants from the National Institutes of Health HL-071643, HL-48129 and Parker B. Francis Foundation fellowship (to G.Z.).

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CHAPTER 8 Reactive Oxygen Species and Endothelial Permeability Masuko Ushio-Fukai, Randall S. Frey, Tohru Fukai and Asrar B. Malik Department of Pharmacology and the Center for Lung and Vascular Biology, University of Illinois, Chicago, IL 60605, USA

I. II. III. IV. V. VI. VII. VIII. IX. X.

Introduction Generation and Metabolism of ROS ROS Generating System in ECs (NADPH Oxidase) Regulation of Adherens Junctions (AJs) by Phosphorylation and by Rho GTPase ROS-generating Stimulants which Regulate Endothelial Permeability ROS Reducing Factors/Proteins which Block Endothelial Permeability Molecular Targets of ROS Regulating Endothelial Permeability Mediators/Regulators of ROS-dependent Endothelial Permeability Functional Significance of ROS-dependent Endothelial Permeability in Vivo Summary and Conclusions References

Abstract Alterations in endothelial permeability are a defining feature of diverse processes including arteriosclerosis, inflammation, ischemia/reperfusion injury, angiogenesis, pulmonary edema in acute lung injury and adult respiratory distress syndrome. Endothelial monolayer permeability increases as a result of both disruption of endothelial cell–cell contacts and EC contraction. Disruption of endothelial cell–cell junctions occurs concomitantly with the redistribution and tyrosine phosphorylation of the VE-cadherin-containing adherens junction (AJ) protein complexes. Little is known about mechanisms of how endothelial permeability is regulated. Reactive oxygen species (ROS) including superoxide (O− 2 ) and hydrogen peroxide (H2 O2 ) generated by activated polymorphonuclear leukocyte (PMNs) and endothelial cells (ECs) impair endothelial barrier integrity by promoting loss of cell–cell adhesions and reorganization of actin cytoskeleton. These responses are involved in promoting transendothelial migration of PMNs and endothelial permeability. Major source of ROS in PMNs and ECs is NADPH oxidase. Phagocyte Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00208-1

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NADPH oxidase consists of membrane-bound gp91phox and p22hox as well as cytosolic components such as p47phox, p67phox and small GTPase Rac. Recently, several novel homologues of gp91phox (Nox2) of NADPH oxidase (Nox) have been cloned in non-phagocytic cells. In ECs Nox1, Nox2, Nox4 and Nox5 are functionally expressed. NADPH oxidase in ECs is activated by inflammatory cytokines, thrombogenic agents, growth factors, G-protein coupled receptor agonists and shear stress. ROS derived from NADPH oxidase function as signaling molecules to activate various redox signaling pathways through modulating activity of kinases and phosphatases, which may contribute to increase in endothelial permeability. Understanding mechanisms by which ROS regulate endothelial permeability is important for the development of novel therapeutic approaches against various diseases such as inflammation, atherogenesis and acute lung injury.

I. INTRODUCTION The endothelium lining the vasculature forms the size-selective and a semipermeable barrier to circulating cells, plasma albumin, macromolecules and bioactive agents between the blood and interstitial tissue. During an inflammatory or thrombogenic response, endothelial monolayer permeability increases as a result of both disruption of endothelial cell–cell contacts and EC contraction. Disruption of endothelial cell–cell junctions occurs mobilization concomitantly with the redistribution of the adherens junction (AJ) protein complexes at the sites of intercellular gap formation. EC contraction occurs by intracellular calcium, Rho GTPase activation, myosin light chain (MLC) phosphorylation, actin microfilament reorganization, and focal adhesion complex (FA) formation. Maintenance of endothelial cell (EC) barrier integrity is critical for preventing development of inflammatory diseases such as atherosclerosis, ischemia-reperfusion injury, diabetes, sepsis and pulmonary edema in acute lung injury (ALI), and adult respiratory distress syndrome (ARDS). Thus, understanding mechanisms of how endothelial permeability is regulated is critically important. Reactive oxygen species (ROS) including superoxide (O− 2 ), hydrogen peroxide (H2 O2 ) and other metalolites have been shown to play a central role in host defense by killing microbes in phagocytic cells such as neutrophils and macrophages. The major source of ROS in phagocytes is the NADPH oxidase, which catalyzes the NADPH-dependent reduction of molecular oxygen to generate large amounts of O− 2 . Accumulating evidence suggest that non-phagocytic cells including EC also produce ROS. Although excess amount of ROS is cytotoxic and induces cell death, ROS at low levels function as signaling molecules to mediate various biological functions such as cell proliferation, migration, differentiation and gene expression. Of note, ROS generated by activated polymorphonuclear leukocytes (PMN) or ECs at sits of inflammation and injury are

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FIGURE 1 Signaling pathways of ROS-mediated increase in endothelial permeability. ICAM-1-dependent PMN binding to ECs and various inflammatory stimuli such as thrombin cytokine, shear stress, acute hypoxia, angiotensin II, advanced glycation end products (AGEs) and VEGF simulate ROS production via activation of NADPH oxidase in ECs. ROS are involved in activation of Src, Pyk2, FAK, PKC and PAK or in oxidative inactivation of PTPs, thereby phosphorylating VE-cadherin and β-catenin. Furthermore, H2 O2 activates TRPM2 channel to increase intracellular Ca2+ to promote loss of cell–cell adhesions at AJs, which may contribute to endothelial barrier dysfunction and increase in endothelial permeability. Moreover, ROS mediate NF-kB-dependent ICAM-1 expression, and thus creating positive-feedback loop whereby PMN-induced ROS in ECs promote PMN endothelial adhesivity through increasing expression of adhesion molecules.

involved in disruption of cell–cell junctions, adhesion molecule expression and leukocyte transendothelial migration, which contribute to endothelial barrier dysfunction and endothelial permeability. ROS are produced following activation of neutrophils, thrombogenic and edemic agents, cytokines, growth factors, Gprotein coupled receptor agonists and shear stress, and play an important role in the pathophysiology of vascular disorders and lung injury (Figures 1 and 2). Signal transduction activated by ROS, “redox signaling” has been an emerging area of investigation.

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FIGURE 2 Role of ROS in endothelial barrier function and inflammatory response in ALI, diabetes and atherosclerosis. In inflammatory state, LPS binding to TLR4 receptor induces NADPH oxidase activation and production of ROS in PMNs as well as ECs, thereby stimulating phosphorylation of AJs complex proteins and MyD88-dependent NF-κB signaling in ECs. These responses play an important role for loss of endothelial barrier integrity and expression of ICAM-1 in cell surface. Adhesion of PMNs to ECs is mediated by binding of constitutive ICAM-1 to CD18 integrin and provides the appropriate coupling required for PMNs to transmit oxidant signals to ECs. Furthermore, various inflammatory stimulants such as thrombin, TNFα, VEGF also stimulates NADPH oxidase activation and production of ROS in ECs, and thus promoting loss of cell–cell adhesions and initiation of NF-kB signaling in ECs, which contribute to increase in vascular permeability. These inflammatory responses are prevented by endogenous growth factor such as PEDF and anticoagulant serine protease such as APC with antioxidant properties. Finally, increased endothelial permeability and/or ROS itself could promote infiltration of inflammatory cells in the tissue via expression of adhesion molecules such as ICAM-1 in ECs, which may participate in tissue repair and remodeling. Thus, ROS produced by PMN and ECs play an important role in various inflammatory diseases such as ALI, diabetes and atherosclerosis through increase in vascular permeability.

ROS are generated from a number of sources including the mitochondrial electron transport system, xanthine oxidase, the cytochrome p450, the NADPH oxidase, uncoupled NO synthase (NOS) and myeloperoxidase. The NADPH oxidase has emerged as major sources of ROS in ECs. The prototype phagocyte NADPH oxidase is composed of membrane-bound gp91phox and p22hox as well

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as cytosolic components such as p47phox, p67phox and small GTPase Rac. Recently, several human homologs of gp91phox (also termed as Nox2) have been identified in non-phagocytic cells. Pro-inflammatory cytokines induce oxidant production via activation of NADPH oxidase and stable adhesion of PMNs to ECs, which in turn promotes microvascular lung injury and pulmonary edema formation (Malik and Lo, 1996) (Figure 2). The overall objective of this review is to summarize the recent information on role of ROS and oxidant signaling in regulating EC permeability. Understanding molecular mechanisms by which ROS regulate endothelial barrier function may lead to development of novel therapeutic strategies for preventing tissue damage due to inflammation, sepsis or other causes of acute lung injury. II. GENERATION AND METABOLISM OF ROS ROS are formed as intermediates in reduction-oxidation (redox) processes, leading from oxygen to water (Fridovich, 1978). The reduction of oxygen by one electron increases the formation of O− 2 , which can be either dismutated to H2 O2 spontaneously or in a reaction catalyzed by superoxide dismutase (SOD). Further reactions lead to the formation of hydroxyl radicals (· OH), especially in the presence of metal ions through the Fenton or Haber–Weiss reactions (Fridovich, 1978; · Deby and Goutier, 1990). O− 2 reacts with nitric oxide (NO ) at a near diffusion− limited rate to form peroxynitrite (ONOO ), which is a potent oxidant. NO· is protective and thus loss of NO· bioavailability via these reactions contributes to endothelial dysfunction and various pathophysiologies. Mammalian peroxidases such as myeloperoxidase (MPO) are activated by H2 O2 to form a highly reac− · tive radical that can oxidize NO· to NO− 2 and react with NO2 to form NO2 . · NO2 can, in turn, participate in nitrating events, such as the formation of nitrotyrosines. Under homeostatic conditions, antioxidant defenses are critical to modulate the steady state balance. ROS are all produced to varying degrees, and are tightly regulated by anti-oxidants such as SOD, catalase, thioredoxin, glutathione peroxidase and other small molecules. Under normal conditions, the rate of ROS production is balanced by the rate of elimination. However, a mismatch between ROS formation and the ability to defend against them by antioxidants results in increased bioavailability of ROS leading to a state of oxidative stress, a risk factor of various inflammatory diseases (Griendling et al., 2000; Landmesser et al., 2005). III. ROS GENERATING SYSTEM IN ECS (NADPH OXIDASE) There are several other enzymatic sources of ROS in mammalian cells, depending on the tissue and environmental context which include the mitochondrial electron transport chain, xanthine oxidase, cytochrome p450, and uncoupled

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eNOS (Griendling et al., 2000; Li and Shah, 2004). There may be complex intereactions among different sources of ROS and feedback and feedforward regulation of ROS accumulation (Li and Shah, 2004). In ECs a major source of ROS is NADPH oxidase (Babior, 2000) which is activated by growth factors, cytokines, shear stress, hypoxia, and G-protein coupled agonists (Griendling et al., 2000). The prototype phagocytic NADPH oxidase consists of the membrane bound cytochrome b558 comprising the catalytic subunit gp91phox (Nox2) and regulatory subunit p22phox , as well as cytosolic subunits, p40phox , p47phox and p67phox , and the GTPase, Rac (DeLeo and Quinn, 1996). Superoxide production is induced by assembly of the cytosolic and membrane-bound subunits, which is mediated through the phosphorylation of p47phox (Bokoch and Knaus, 2003). The neutrophil NADPH oxidase is normally quiescent in basal state but releases large amounts of O·− 2 in bursts upon activation during phagocytosis via the one electronreduction of oxygen by NADPH, which participate in host defense by killing invading microbes (Babior, 1995; Lambeth, 2004). By contrast, the non-phagocytic NADPH oxidase(s) continuously produce low levels of O·− 2 intracellularly in basal state, yet it can be further stimulated acutely by various agonists and growth factors. Several human homologs of gp91phox (also termed as Nox2) have been identified including Nox1, Nox3, Nox4, Nox5, Dual oxidases (Duox1/2) (Lambeth, 2004; Cheng et al., 2001; Geiszt et al., 2000; Geiszt, 2006). All Nox family members share the common binding sites for FAD, heme, and NADPH and 6 transmembrane domains. Each Nox is encoded by different genes (Lambeth, 2004; Lassegue and Clempus, 2003). Recently, isoforms of p47phox and p67phox were discovered. These were termed as NoxO1 (for Nox organizer 1) and NoxA1 (for Nox activator 1), which substitute for p47phox and p67phox respectively (Banfi et al., 2003; Geiszt et al., 2003). An important difference between p47phox and NoxO1 is that the latter lacks the p47phox domain that is regulated by phosphorylation; therefore NoxO1 may influence oxidase activity quite differently from p47phox . In ECs, all the phagocytic NADPH oxidase subunits including Nox2, p22phox, Rac1, p47phox as well as Nox1, Nox4 and Nox5 are expressed, and various Nox enzymes contribute to ROS production at different degree, location and EC types (Li and Shah, 2004; Babior, 2000). In porcine pulmonary artery ECs, histochemical data showed the presence of Nox2 and p22phox well as p47phox and p67phox (Hohler et al., 2000). In human umbilical vein ECs, RT-PCR demonstrated the presence of mRNA for all the Nox enzymes (Jones et al., 1996). ECs cultured from rat aorta express mRNA for Nox4 in addition to Nox2 and a membrane preparation demonstrated that ROS production is inhibited by Nox4 antisense oligonucleotide (Datla et al., 2007). In addition, Nox5 is expressed in human microvascular ECs (BelAiba et al., 2007). To our knowledge, expression of Nox3, Duox, NoxO1 and NoxA1 in ECs has not been demonstrated.

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IV. REGULATION OF ADHERENS JUNCTIONS (AJS) BY PHOSPHORYLATION AND BY RHO GTPASE Endothelial AJs maintain endothelial barrier integrity by connecting adjacent ECs and their association with the cytoskeleton, allowing the cells to form a tight barrier between the vessel lumen and the stroma. Vascular endothelial cadherin (VE-cadherin) is an essential transmembrane homophilic adhesion molecule in endothelial AJs that control cell–cell adhesion in ECs (Dejana et al., 1995). The cytoplasmic domain of VE-cadherin binds to cytosolic proteins either β-catenin, plakoglobin or p120catenin which couple to the actin cytoskeleton through αcatenin (Bazzoni and Dejana, 2004). Some studies have shown that interaction of α-catenin with β-catenin or with actin is mutually exclusive (Drees et al., 2005; Yamada et al., 2005). The linkage between VE-cadherin-based AJ complex and actin cytoskeleton contributes to the strong adhesion and the endothelial barrier integrity (Figure 1). Phosphorylation states of AJ proteins can alter the interactions between components of AJ proteins. In order to maintain endothelial barrier function, VEcadherin function is tightly regulated through mechanisms that involve protein phosphorylation and cytoskeletal dynamics (Esser et al., 1998; Lampugnani et al., 1997; Lilien and Balsamo, 2005; Alema and Salvatore, 2007). Tyrosine phosphorylation of AJ components which is related to the impairment of cell–cell adhesion is regulated by the state of confluence of the cells. Tyrosine phosphorylation of VE-cadherin, β-catenin and p120 occurs in looser AJs at early cell contacts, while is reduced by the formation of mature and cytoskeleton-connected junctions in tightly confluent cells (Lampugnani et al., 1997). Thus, changes in phospho-tyrosine content parallels to changes in the molecular organization of cell–cell contacts. Of note, ROS production is also dependent on cell density. ROS levels are increased and decreased at low and high confluent cells, respectively (Pani et al., 2000), which is consistent with the idea that ROS are involved in disruption of cell–cell contacts in ECs (Figure 1). Rho-like small GTPases are key molecular switches that control cytoskeletal dynamics and cadherin function. VE-cadherin colocalizes with actin stress fibers at cell–cell junctions in ECs. Inhibition of VE-cadherin-mediated adhesion induces reorganization of the actin cytoskeleton and loss of cell–cell adhesion, which contributes to increased monolayer permeability and neutrophil transendothelial migration. Modulation of the actin cytoskeleton strongly affects VE-cadherin distribution. Cytoskeletal reorganization resulting from the inactivation of p21Rho causes a diffuse localization of VE-cadherin, which is accompanied by the reduced cell–cell adhesion. Thus, monolayer permeability is modulated by VE-cadherin-mediated cell–cell adhesion, which is in turn controlled by the dynamics of the actin cytoskeleton through Rho GTPases (Hordijk et al., 1999).

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V. ROS-GENERATING STIMULANTS WHICH REGULATE ENDOTHELIAL PERMEABILITY Thrombin Thrombin proteolytically cleaves and activates the protease-activated receptors PAR1 that are present on EC surface, leading to disassembly of AJs and reorganization of the actin cytoskeleton, which results in increased endothelial permeability and barrier disruption (Mehta and Malik, 2006). With the use of an intact lung model, thrombin is shown to increase pulmonary vascular resistance in conjunction with a rise in the capillary filtration coefficient and lung water content (Horgan et al., 1987; Malik and Horgan, 1987). Thrombin increases ROS production albeit to a slower kinetics than that of TNFα (Coughlin, 2000; Lum and Malik, 1996; Tiruppathi et al., 2002). Thrombin stimulates ROS production in a Nox5-dependent manner in human ECs (BelAiba et al., 2007) or through upregulating p22phox of NADPH oxidase in human EC line (Djordjevic et al., 2005). The nonreceptor protein-tyrosine phosphatase (PTP) SHP2 co-precipitates with VE-cadherin complexes in confluent, quiescent HUVECs. Thrombin stimulates SHP2 tyrosine phosphorylation and its dissociation from VE-cadherin complexes, thereby regulating the tyrosine phosphorylation of AJs complex proteins, which contributes to increased endothelial permeability (Ukropec et al., 2000). Although it remains unknown how ROS regulate thrombin-induced increase in endothelial permeability, it is possible that ROS are involved in SHP-2 phosphorylation or oxidative inactivation of this PTP, thereby promoting tyrosine phosphorylation of AJs proteins and barrier dysfunction. PAR1 stimulates RhoA through heterotrimeric G proteins Gα12/13 binding to the guanine nucleotide exchange factor p115RhoGEF that facilitates its GEF activity, leading to RhoA activation (Kozasa et al., 1998). Gαq-mediated stimulation of PKCα activates RhoA by phosphorylating Rho guanosine diphosphate (GDP) dissociation inhibitor (Rho GDI) and p115RhoGEF (Mehta et al., 2001; Holinstat et al., 2003; Knezevic et al., 2007). Downstream of RhoA, Rho-associated kinase (ROCK) and LIM kinase induce reorganization of actin and microtubule cytoskeleton, thereby increasing acto-myosin contractility (Yang et al., 1998; Gorovoy et al., 2005; Essler et al., 1998) by phosphorylating MLC phosphatase, MYPT1, which in turn increasing MLC phosphorylation (Birukova et al., 2004d). These responses result in endothelial barrier dysfunction and pulmonary edema associated with ALI. By contrast, H2O2-induced MLC phosphorylation and actin rearrangement are mediated through MLCK-dependent pathway in bovine pulmonary artery ECs (Zhao and Davis, 1998). Independent of Rho activation, activation of atypical PKCzeta is also involved in thrombin-induced endothelial permeability in HUVECs (Li et al., 2004). Of interest, angiopoietin-1 protects endothelial barrier integrity at least in part through inhibition of thrombin-induced activation of both PKCzeta and Rho (Li et al., 2004).

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TNF-α TNF-α is an inflammatory cytokine, and is released from activated monocytes and macrophages sequestered in the microcirculation (Hocking et al., 1990) and is increased in bronchoalveolar lavage fluid from patients with ARDS (Malik et al., 1989; Suter et al., 1992). TNF-a increases ROS production and stimulates lung vascular barrier dysfunction upon engaging TNF-α receptor-1, leading to increased endothelial permeability in cultured EC and in vivo (Lum and Malik, 1996; Hocking et al., 1990; Horvath et al., 1988; Ferro et al., 2000; Partridge et al., 1993). In addition, TNF-α upregulates ICAM-1 and E-selectin expression (Lo et al., 1992; Rahman et al., 1998, 2000), thereby promoting neutrophil adhesion to endothelium and ROS generation (Rahman et al., 1998), which in turn increases endothelial permeability. Mechanically, TNF-α induces reorganization of the actin cytoskeleton and increases formation of intercellular gaps and EC monolayer permeability through promoting loss of cell–cell junctions by activation of Rho, Rac, and Cdc42 (Wojciak-Stothard et al., 1998) or by tyrosine phosphorylation of VE-cadherin (Nwariaku et al., 2002). Nwariaku et al. (2004) reported that TNF-α-induced loss of endothelial junctional integrity is mediated through PAK1-NADPH oxidase-JNK-VE-cadherin phosphorylation pathway. Thus, PAK1 is a potential therapeutic target for oxidant stress-dependent inflammatory diseases which are associated with increased endothelial permeability. Transforming Growth Factor-β1 (TGF-β1) TGF-β1 is a cytokine critically involved in acute lung injury and EC barrier dysfunction. TGF-β1-induced decrease in pulmonary endothelial monolayer integrity is associated with disassembly of the adherens junction. This response may be due to activation of a MLC kinase (MLCK)-dependent signaling cascade (Hurst et al., 1999). Mechanically, TGF-β1-mediated adherens junction disassembly is associated with interaction of TGF-beta type II receptor with Smad3 and Smad4 (Tian and Phillips, 2002). TGF-β1 induces barrier dysfunction, which is linked to increased actin stress fiber formation, MLC phosphorylation, EC contraction, and gap formation, which is abolished by inhibition of Rho and Rhokinase, and by MT stabilization with taxol (Birukova et al., 2005). Thus, MT dynamics plays an important role in the TGF-β1-mediated Rho regulation, EC barrier dysfunction, and actin remodeling. Lu et al. (2006) reported that TGF-β1 induces endothelial barrier dysfunction through Smad2-dependent p38 activation, resulting in RhoA activation. TGF-β itself induces ROS production as part of its signal-transduction pathway (Koli et al., 2008). However, role of TGF-β-induced ROS in EC barrier dysfunction remains unclear. Vascular Endothelial Growth Factor (VEGF) VEGF is one of the potent angiogenesis factor (Ferrara and Davis-Smyth, 1997) and vascular permeabilizing agents, also named vascular permeability fac-

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tor (Dvorak et al., 1995). Lung endothelial damage is a characteristic morphological feature of ischemia-reperfusion injury. Pretreatment with adenovirus encoding soluble VEGF receptor type II (VEGFR2) prevents I/R-mediated increase in lung vascular permeability in rats. Activation of the stress protein response with geldanamycin or pyrrolidine dithiocarbamate prevents the development of pulmonary edema by inhibiting VEGF-induced phosphorylation of VE-cadherin and the formation of actin stress fibers in a model of lung ischemia-reperfusion injury in rats (Godzich et al., 2006). Hence it plays a key role in the pathogenesis of ALI/ARDS (Medford and Millar, 2006). Endothelial AJ is a downstream target of VEGFR2 signaling linked to increase of vascular permeability in cultured EC monolayers (Godzich et al., 2006). VEGF induces tyrosine phosphorylation of AJ components, leading to loss of cell–cell contacts, which contributes to increase in vascular permeability and angiogenesis (Esser et al., 1998). VEGF stimulation promotes association of cSrc with VEGFR2, which is required for Src-dependent tyrosine phosphorylation of VE-cadherin at Tyr685 and loss of VE-cadherin-mediated cell–cell adhesion, thereby promoting vascular permeability and angiogenesis (Chou et al., 2002; Lambeng et al., 2005; Wallez et al., 2007). Furthermore, VEGF activation of redox-sensitive protein kinase C (PKC)βII increases phosphorylation of tight junction protein occluding, thereby increasing endothelial permeability (Harhaj et al., 2006). Thus, regulation of PKC activity by ROS and tight junction protein modification may have therapeutic implications. Formation of FAs involved in EC contraction also regulates EC permeability. VEGF stimulates tyrosine phosphorylation and recruitment of focal adhesion kinases (FAK) and paxillin to new FAs in ECs (Abedi and Zachary, 1997). VEGF-stimulated FA complex assembly in human microvascular ECs is mediated through activation of FAK and Pyk2 (Avraham et al., 2003) which are targets of ROS. Rho-ROCK pathway contributes to VEGF-induced microvascular endothelial hyperpermeability. MLC phosphorylation and actin stress fiber formation occur concomitantly with the increase in permeability upon VEGF stimulation (Sun et al., 2006). VEGF also promotes Src-mediated phosphorylation of FAK on Tyr861, leading to the formation of a FAK/αvβ5 signaling complex involved in increase in vascular permeability, which is significantly reduced in pp60csrc-deficient mice (Eliceiri et al., 2002). Thus, Src-mediated coupling of FAK to integrin αvβ5 is required for VEGF-stimulated vascular permeability. Taken together, cSrc is a potential molecular target regulating VEGF-induced loss of cell–cell contacts and formation of FAs, which contribute to increase in endothelial permeability. VEGF rapidly increases ROS production via activation of Rac1-dependent NADPH oxidase in ECs (Colavitti et al., 2002; Abid et al., 2007; Ushio-Fukai, 2007; Ushio-Fukai et al., 2002). Resveratrol, a polyphenolic compound in grapes inhibit ROS-dependent Src kinase activation and VE-cadherin tyrosine phosphorylation (Lin et al., 2003). Green tea catechins with antioxidant properties inhibit

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VEGF-induced phosphorylation of VE-cadherin and Akt as well as tube formation in human microvascular ECs presumably through inhibition of cSrc (Tang et al., 2003). Thus, cSrc is a downstream target of VEGF-induced ROS and involved in tyrosine phosphorylation of VE-cadherin, thereby promoting disruption of cell–cell contacts in confluent ECs and endothelial permeability (Lin et al., 2003). VEGF-induced increase in endothelial permeability is also mediated through eNOS-dependent mechanism of transcytosis in caveolae. Localization of Flk-1/KDR and eNOS with caveolin-1 suggests that VEGF signaling occurs within the caveolar compartment (Feng et al., 1999). Of note, NADPH oxidase is also localized in Cav1-enriched caveolae fraction in ECs (Yang and Rizzo, 2006), suggesting that peroxinitrite formed by reaction of eNOS-derived NO and NADPH oxidase-derived O− 2 may be involved in this process. VEGF also mediates airway inflammation and remodeling in asthma of lung through regulating MMP-9 expression (Lee et al., 2006). NADPH oxidase-derived ROS stimulate induction of MMP-9 expression, which may be relevant in oxidative stress-dependent inflammatory diseases (Zalba et al., 2007). VEGF also stimulates rapid endocytosis of VE-cadherin through beta-arrestin, thereby disrupting the endothelial barrier function (Gavard and Gutkind, 2006). This process is initiated by activation of Rac1 through the Src-dependent phosphorylation of Vav2, a guanine nucleotide-exchange factor, which in turn, promotes the p21-activated kinase (PAK)-mediated phosphorylation of the intracellular tail of VE-cadherin. This results in the recruitment of beta-arrentin2 to serine-phosphorylated VE-cadherin, thereby promoting its internalization into clathrin-coated vesicles and the consequent disassembly of intercellular junctions. Given that Rac1 is a critical component of NADPH oxidase, and that Src-dependent VE-cadherin phosphorylation is ROS-dependent, it is likely that NADPH oxidase-derived ROS may be involved in this mechanism. Understanding this novel signaling mechanism may help to identify new therapeutic targets for the treatment of many human diseases that are characterized by vascular leakage. Lysophosphatidylcholine (LysoPC) LysoPC, a major component of oxidized low density lipoprotein (oxLDL), evokes diverse biological and pathophysiological responses through ROS production and increases endothelial permeability (Essler et al., 1999; Yan et al., 2005). In human coronary artery ECs, lysoPC increases endothelial monolayer permeability and reduces the expression of tight junction molecules including zonula occludens-1, occludin, claudin-1, and junctional adhesion molecule through increase in oxidative stress and activation of c-Jun N-terminal kinase (JNK) and p38 mitogen-activated protein kinase (p38MAPK). The antioxidant can effectively block LysoPC-induced endothelial permeability, suggesting an involvement of ROS. Another mechanism for oxidized LDL-induced increased endothelial

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permeability includes its effect on lysyl oxidase, a key enzyme involved in extracellular matrix (ECM) synthesis. Hypoxia NADPH oxidases have been shown to serve as oxygen sensors in the lung. Chronic hypoxia induces vascular remodeling with medial hypertrophy leading to the development of pulmonary hypertension. NOX1, NOXA1, NOXO1, p22phox, p47phox, p40phox, p67phox, NOX2, and NOX4 are present in mouse lung tissue. Mice maintained for 21 days under hypoxic (10% O2 ) or normoxic (21% O2 ) conditions show upregulation of NOX4 mRNA by hypoxia in homogenized lung tissue (Mittal et al., 2007). In lungs from patients with idiopathic pulmonary arterial hypertension (IPAH), expression of NOX4 is upregulated mainly in the vessel media (Mittal et al., 2007). EC grown to confluence under hypoxia (5% O2 ) form a tighter monolayer than ECs grown under normoxia (21% O2 ). This tighter barrier in hypoxic cells appears to be due, in part, to the inhibition of RhoA activity (Solodushko et al., 2007). Conversely, severe acute hypoxia disrupts the EC barrier and increases EC permeability (Wojciak-Stothard et al., 2005; Ogawa et al., 1990) due to an increase of ROS (Wojciak-Stothard et al., 2005). Hypoxia/reoxygenation Hypoxia/reoxygenation-induced changes in endothelial permeability are accompanied by endothelial actin cytoskeletal and adherens junction remodeling, which is mediated through coordinated actions of the Rho GTPases Rac1 and RhoA. Rac1 and RhoA are rapidly activated by changes in oxygen tension, and their activity depends on NADPH oxidase- and PI3 kinase-dependent production of ROS. Rac1 acts upstream of RhoA, and its transient inhibition by acute hypoxia leads to activation of RhoA, followed by stress fiber formation, dispersion of adherens junctions, and increased endothelial permeability. Reoxygenation strongly activates Rac1 and restores cortical localization of F-actin and VE-cadherin. This effect is due to Rac1-mediated inhibition of RhoA, which can be prevented by activators of RhoA and lysophosphatidic acid (Wojciak-Stothard et al., 2005). Thus, Rho GTPases act as mediators coupling cellular redox state to endothelial function. Polymorphonuclear Leukocyte (PMN) Lung inflammatory disease is characterized by increased PMN infiltration and vascular permeability. Leukocyte transendothelial migration occurs through intercellular gaps between ECs or transcellular pathway which is dependent on caveolae/caveolin-1. PMN adhesion to activated EC during acute inflammation promotes formation of intercellular gaps between ECs by loss of endothelial cell– cell adhesion, resulting in increased EC permeability and PMN transendothelial migration (Ionescu et al., 2003; Del Maschio et al., 1996). Mechanically, activated neutrophils induce hyperpermeability through tyrosine phosphorylation

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of AJs proteins in coronary venular ECs (Tinsley et al., 1999). ROS produced from primed leukocytes are involved in the pathogenesis of lung injury. p47phox KO mice show greater alveolar neurtrophilic leucocyosis and have a decreased number of macrophages in bronchoalveolar lavage after acid or saline aspiration. NF-kappaB activation and resultant ICAM-1 expression in ECs occur secondary to oxidants generation by the PMN NADPH oxidase complex. The functional relevance of the PMN NADPH oxidase in mediating TNFα-induced ICAM-1dependent endothelial adhesivity is evident by reduced adhesion of p47phox-/PMN in co-culture experiments (Fan et al., 2002). Thus, oxidant signaling by PMN NADPH oxidase complex is an important determinant of TNFalphainduced NF-kB activation and ICAM-1 expression in ECs. Moreover, augmented NF-kB activation and the TLR2 upregulation in ECs are secondary to oxidant signaling generated by PMN NADPH oxidase. LPS induces TLR2 up-regulation through TLR4- and MyD88-dependent signaling. The functional relevance of NADPH oxidase in mediating TLR4-induced TLR2 expression in ECs is evident by markedly elevated and stable ICAM-1 expression as well as augmented PMN migration in response to sequential challenge with LPS and peptidoglycan (Fan et al., 2003). Thus, PMN NADPH oxidase-derived oxidant signaling is an important determinant of the cross-talk between TLR4 and TLR2 and EC activation (Figures 1 and 2). ICAM-1 and VCAM-1 Endothelial activation is an early step during leukocyte/endothelial adhesion and leukocyte transendothelial migration in inflammatory state. Leukocyte adhesion is mediated totally and transendothelial migration partially by heterotypic interactions between the β1 and β2 integrins on the leukocytes and their ligands, Ig-like cell adhesion molecules VCAM-1 and ICAM-1 on the endothelium. Both integrins and Ig-CAMs are known to have signaling capacities. Clustered ICAM-1 and VCAM-1 are in very close proximity on the EC surface in particular following adhesion of leukocytes that use β1 and β2 integrins for transmigration, suggesting that the signaling induced by these molecules may be interconnected. Whether ROS are involved in cross-talk between ICAM-1 and VCAM-1-induced signaling events remain unknown. ICAM-1 Five ICAMs have been discovered thus far and are members of the Ig supergene family and are receptors for the β2 family of integrins on leukocytes (Hopkins et al., 2004; Hubbard and Rothlein, 2000). ICAM-1 is one of the main integrin ligand and specifically involved in the leukocyte migration across the endothelial barrier. ICAM-1 knockout mice show impaired inflammatory and immune responses (Sligh et al., 1993). Antisense oligonucleotides to ICAM-1 decreases leukocyte adhesion and inflammation (Rijcken et al., 2002). ICAM-1 is normally present in low levels on ECs, but its expression is dramatically in-

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creased in response to various proinflammatory stimuli including TNF-α, IL-1β, and LPS (Myers et al., 1992), phorbol myristate acetate (Wertheimer et al., 1992), thrombin (Rahman et al., 2004) and shear stress (Tzima et al., 2002). ICAM-1 is localized at the apical surface of ECs on activation and is organized in microdomains along with VCAM-1 at around adherent leukocytes. The cytoplasmic C-terminus of ICAM-1 associates to the actin-binding ERM proteins (ezrin, radixin, moesin) as well as to α-actinin (Carpen et al., 1992; Etienne-Manneville et al., 2000). ICAM-1 acts as a signal transducer in ECs. ICAM-1 cross-linking activates the p60src kinase to phosphorylate cortactin (Durieu-Trautmann et al., 1994), and releases intracellular calcium and activates the Rho GTPase, which contributes to regulating actin cytoskeleton and EC contractility (Lorenzon et al., 1998). Moreover, ICAM-1 activates p60src via the activation of xanthine oxidase, leading to tyrosine phosphorylation of ezrin and p38 MAPK (Wang et al., 2003). These effects are mediated by the C-terminus of ICAM-1 and are required for increase in endothelial permeability and enhanced lymphocytes and leukocyte transendothelial migration (Greenwood et al., 2002; Yang et al., 2005; Lyck et al., 2003). ICAM-1 activation also induces tyrosine phosphorylation of focal adhesion signaling proteins including FAK, paxillin, and p130Cas in parallel to Rho activation. Tyrosine-phosphorylated Cas associates with the adaptor protein Crk and the GTP exchange factor C3G. In addition, tyrosine phosphorylation of FAK, paxillin, and Cas as well as Crk and JNK activation are blocked by pretreatment of the cells with inhibition of Rho (Etienne et al., 1998). Thus, activation of ICAM-1 may couple Rho to phosphorylation of FAs proteins, thereby promoting leukocyte transmigration. Moreover, ICAM-1 engagement activates tyrosine kinases, Src and proline-rich tyrosine kinase 2 (Pyk2), which induces phosphorylation of VE-cadherin on Tyr658 and Tyr731, respectively, which correspond to the p120-catenin and β-catenin binding sites, respectively (Allingham et al., 2007). Importantly, both Src and Pyk2 are activated by ROS. PMN adhesion to EC is increased in ROS-treated HUVEC (Sellak et al., 1994) which is inhibited by anti-ICAM-1, anti-CD11a, anti-CD11b, and anti-CD18 antibodies (Sellak et al., 1994). EC surface expression of ICAM-1 increases ROS production (Lo et al., 1993). Inhibition of endogenous catalase in ROS-treated EC augments the PMN adhesion, whereas catalase treatment blocks the ROS-induced PMN adhesion, indicating that oxidant-antioxidant balance at the EC interface is a critical factor modulating PMN-EC adhesive interactions (Lo et al., 1993) (Figures 1 and 2). VCAM-1 The main β1-integrin ligand on the endothelium, VCAM-1, is, in contrast to ICAM-1, absent in resting cells but greatly up-regulated by inflammatory stimuli. Similarly to ICAM-1, VCAM-1 not only acts as an adhesion receptor, but also as a signal transducer upon binding of leukocytes. The cytoplasmic domain of VCAM-1 is only 19 amino acids long and comprises a type I PDZ-binding motif.

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However, whether specific interactions are mediated by this motif is unknown; to date, only ezrin and moesin have been shown to associate with the cytoplasmic domain of VCAM-1 (Barreiro et al., 2002). VCAM-1 cross-linking induces the Rac1-dependent ROS production, activation of p38 MAPK and changes in the actin cytoskeleton (i.e. stress fiber formation), which contribute to increased endothelial permeability and leukocyte transendothelial migration (van Wetering et al., 2003). Of note, VCAM1-induced Rac1-mediated ROS production is involved in phosphorylation of Pyk2 that controls endothelial integrity through the phosphorylation of β-catenin (van Buul et al., 2005). ROS may inactivate VEcadherin either by proteolytic breakdown or by reducing its homophilic adhesion through reduction of its link to the actin cytoskeleton. Moreover, VCAM-1 activation of endothelial NADPH oxidase induces transient activation of PKCα via oxidation, which is necessary for VCAM-1-dependent transendothelial cell migration (Abdala-Valencia and Cook-Mills, 2006). Integrins Adhesions of cells to extracellular matrix and adjacent cells are mediated by integrins and VE-cadherin, respectively. Understanding how they are coordinated to regulate endothelial integrity is essential. Integrins transmit signals from “outside in” as well as “inside out”, thus acting as signal transducing molecules as well as mechano-sensors. Integrin engagement by fibronectin-coated beads leads to disruption of the VE-cadherin-containing AJs (Wang et al., 2006), which is accompanied by increases of tyrosine phosphorylation of β-catenin and p120ctn, as well as the dissociation of α-catenin from VE-cadherin. A membrane-targeted Src reporter with the fluorescence resonance energy transfer in live cells shows that the integrin engagement induces activation of Src at adherens junctions, which is subsequently disrupted as well as significant alteration of cortical actin filaments at AJs. Given that integrin engagement activates cSrc through ROS production (Giannoni et al., 2005), ROS may be involved in disruption of VE-cadherin containing AJs via modulation of the actin cytoskeleton networks. Integrin αvβ5 regulates lung vascular permeability and pulmonary endothelial barrier function (Su et al., 2007). A function-blocking antibody against the alphavbeta5 prevents lung vascular permeability in two different models of ALI: ischemia-reperfusion in rats, which is mediated by VEGF, and ventilation-induced lung injury in mice which is mediated at least in part by TGF-β. Mice lacking β3 show enhanced angiogenesis and vascular leak in response to VEGF (Reynolds et al., 2002; Taverna et al., 2004). Mice lacking β5 show less edema in models of ischemic stroke or pulmonary ischemia-reperfusion or ventilator-induced lung injury (Su et al., 2007). Beta1 integrins can also influence the disruption of vascular permeability during inflammation, since β1 integrins can influence the ability of circulating cells to arrest, adhere, and extravasate at sites of injury and vascular leak (Sixt et al., 2006). Thus, several integrins regulate pulmonary endothelial permeability, and seem to be potentially attractive therapeutic targets for ALI.

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Shear Stress Shear stress plays an important role in atherosclerosis development. The disassembly of AJs in response to shear is important for the endothelial permeability barrier and other aspects of endothelial cell function (Noria et al., 1999). ECs exhibit profound changes in cell shape in response to shear stress that may require disassembly/reassembly of AJs complexes that mediate cell–cell adhesion (Seebach et al., 2000). In response to shear in endothelial monolayers, α-catenin that links β-catenin-bound cadherin to the actin cytoskeleton is dissociated from VE-cadherin complexes. This is associated with increased tyrosine phosphorylation of β-catenin bound to VE-cadherin and loss of SHP-2 from VE-cadherin complexes (Ukropec et al., 2002). Thus, the functional interaction of α-catenin with VE-cadherin-bound β-catenin is modulated by the extent of tyrosine phosphorylation of β-catenin that is regulated by SHP-2 associated with VE-cadherin complexes. Acute laminar shear stress induces cortical cytoskeletal remodeling, which is associated with transient peripheral translocation of cortactin, an actin-binding protein involved in the regulation of actin polymerization. This cortactin translocation is mediated through Rac1 and PAK1, but not Src, MLCK or ROCK (Birukov et al., 2002). In contrast, Shikata et al. (2005) reported that shear stress induces peripheral accumulation of FAs, activation of Rac1 without affecting Rho and FAK phosphorylation, thereby increasing transendothelial electrical resistance (TER) in human pulmonary EC, which is further promoted by barrier-protective phospholipid sphingosine 1-phosphate (S1P). Oscillatory shear stress increases ROS levels in ECs, whereas laminar shear stress blocks this response. Recently, Mowbray et al. (2008) reported that laminar shear upregulates mechanosensitive antioxidant, peroxiredoxin-1 in EC. Thus, effects of shear stress on endothelial barrier integrity seem to be dependent on extent and duration of shear as well as cell types. Advanced glycation end products (AGEs) The formation of AGEs is an important biochemical abnormality that accompanies diabetes mellitus and inflammation (Basta et al., 2004). Interaction between AGEs and their receptor (RAGE) evokes generation of ROS in EC, which may contribute to increase in endothelial permeability and vascular inflammation. The increase in permeability is accompanied by alterations of the physical integrity of the endothelium, as shown by the destruction of organized actin structures and alterations of cellular morphology (Esposito et al., 1989; Wautier et al., 1996). sRAGE, the extracellular ligand-binding domain of RAGE, blocks AGEs from binding to RAGE, and suppresses accelerated atherosclerotic lesion formation and decreases vascular hyperpermeability (Wautier et al., 1996; Park et al., 1998; Bucciarelli et al., 2002; Goova et al., 2001). Levels of AGEs and RAGE are increased in streptozotocin-treated (diabetic) apolipoprotein Enull mice that have advanced atherosclerosis (Esser et al., 1998). Blockade of

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RAGE by sRAGE in mice with diabetes reduces atherosclerotic lesion area, supporting the notion that RAGE is a crucial for the development and acceleration of atherosclerosis and that sRAGE is an effective intervention (Park et al., 1998; Bucciarelli et al., 2002). Angiotensin II (Ang II) Vascular inflammation is involved in the initiation and progression of atherosclerosis, hypertension- and diabetes-induced vascular complications. Angiotensin II (Ang II), the key effector of the renin-angiotensin system, plays a central role in the regulation of blood pressure and also capable of inducing inflammatory response in the vascular wall, which includes increase in vascular permeability, infiltration of leukocytes, and tissue remodeling. Ang II increases vascular permeability via AT1 receptors, whereas stimulation of AT2 receptors exerts an opposite effect (Victorino et al., 2002). In both hypertensive and normotensive humans, ACE inhibitors produce pulmonary vasodilation and an increase in capillary volume that are blocked by cyclooxygenase inhibitors (Linz et al., 1995; Swartz and Williams, 1982). In addition, ACE inhibitors improve pulmonary diffusion in chronic heart failure, which is prevented by aspirin, a prostaglandin inhibitor (Guazzi et al., 1997). These suggest an importance of prostaglandin in Ang II-related alveolar-capillary membrane diffusing capacity or capillary permeability. Ang II-induced VEGF may contribute to the pathogenesis via the up-regulation of vascular permeability (Cheng et al., 2005). Importantly, Ang II is a potent stimulator for ROS production via activation of NADPH oxidase in vasculature (Griendling et al., 2000). Given that both Ang II and VEGF stimulate ROS production in the vessel wall, it is tempting to speculate that Ang II-induced increase in vascular permeability is due to an increase in ROS. 4-Hydroxy-2-nonenal (4-HNE) 4-HNE is one of the major biologically active aldehydes formed during inflammation and oxidative stress, and is implicated in a number of cardiovascular and pulmonary disorders involved in vascular endothelial permeability. In bovine lung microvascular ECs, 4-HNE induces ROS generation, thereby increasing EC permeability by modulating cell–cell adhesion through focal adhesion, adherens and tight junction proteins as well as integrin signal transduction (Usatyuk et al., 2006). This mechanism may lead dramatic alteration in endothelial cell barrier dysfunction during heart infarction, brain stroke, and lung diseases. Microparticles Endothelium-derived microparticles (EMPs) are shown as a new marker of EC dysfunction. Increased levels of circulating microparticles have been observed in inflammatory disorders and diabetes mellitus. Endothelium-derived EMPs induce attenuation of endothelium-mediated vasodilation and eNOS-derived NO production, and promote pulmonary edema, thereby contributing to acute lung

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injury (Densmore et al., 2006). Mezentsev et al. (2005) showed that EMPs impair angiogenesis-related responses in vitro, which is partially inhibited by cell-permeable SOD mimetic Mn-TBAP. Thus, EMPs-induced ROS seem to contribute to impairment of endothelial function and edema formation. Thus, EMPs may represent new therapeutic targets. Rac1 Activation of Rac1 stimulates generation of ROS, which are capable of activating Pyk2 (van Wetering et al., 2003; Frank et al., 2000) and Src (Allingham et al., 2007). Rac1-induced ROS disrupt VE-cadherin-mediated cell–cell adhesion through increase in tyrosine phosphorylation of α-catenin, a component the VE-cadherin-catenin complex in HUVECs (van Wetering et al., 2002). Inhibition of VE-cadherin function induced by VE-cadherin-blocking antibodies activates Rac1, thereby increasing production of ROS, which subsequently promotes intercellular gap formation in HUVECs. This loss of cell–cell contacts is due to increase in tyrosine phosphorylation of VE-cadherin-associated beta-catenin by Pyk2 (van Buul et al., 2005) that regulates cell adhesion (Lakkakorpi et al., 1999) and phosphatidylinositol 3-kinase/Akt pathways (Rocic et al., 2001). Thus, Rac-mediated production of ROS regulate VE-cadherin function and may play an important role in the (patho)physiology associated with inflammation and endothelial damage. In other ECs such as human pulmonary artery ECs, disruption of AJs by thrombin-induced signaling pathways is associated with an increase in RhoA activity and inactivation of Rac1, whereas reassembly of AJs is associated with activation of Cdc42 and Rac1 and inhibition of RhoA (Mehta and Malik, 2006; Kouklis et al., 2004). Activation of Rac1 counteracts RhoA activity, providing a mechanism for small GTPase coordination of AJ integrity (Noren et al., 2001; Noren et al., 2003). RhoA inactivation at cell–cell adhesion sites depends on both the activity and cortical localization of the GTPase activating protein (GAP) p190RhoGAP that stimulates GTP hydrolysis of RhoA (Wildenberg et al., 2006). Activation of p190RhoGAP is mediated through tyrosine phosphorylation by FAK (Holinstat et al., 2006) and Src (Noren et al., 2003). Src-induced phosphorylation of p190RhoGAP at Tyr1105 facilitates p190RhoGAP activity toward RhoA (Chang et al., 1995; Roof et al., 1998). Platelet-derived growth factor activates Rac and leads to translocation of p190RhoGAP to AJs, where it couples to the cadherin complex by interacting with p120-catenin (Wildenberg et al., 2006). A p120-catenin plays an indispensable role in the subcellular localization and function of p190RhoGAP. Thus, p120-catenin provides a platform for the physical interaction between p190RhoGAP and RhoA, and linking the cadherin-catenins complex to RhoA inhibition spatially coordinates the cross-talk between Rho GTPase signaling pathways at AJs (Wildenberg et al., 2006).

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VI. ROS REDUCING FACTORS/PROTEINS WHICH BLOCK ENDOTHELIAL PERMEABILITY Pigment Epithelium-Derived Factor (PEDF) PEDF decreases ROS generation in AGE-elicited, VEGF-mediated vascular hyperpermeability by downregulation of mRNA levels of p22phox and gp91phox of NADPH oxidase (Yamagishi et al., 2006). This leads to blockade of the AGEinduced Ras activation and NF-kappaB-dependent VEGF gene induction in ECs. Thus, PEDF inhibits AGE signaling to vascular permeability by suppression of NADPH oxidase-mediated ROS generation and subsequent VEGF expression. Thus, PEDF is an important endogenous growth factor with antioxidant properties and may function to block ROS-mediated microvascular permeability during inflammation. Activated Protein C (APC) APC is an important natural anticoagulant serine protease that is proteolytically generated from protein C by the modulation of thrombin activity in the presence of thrombomodulin on an endothelial surface. Recent studies have demonstrated that, APC has anti-inflammatory and cytoprotective effects and improves sepsis survival through an unknown mechanism. Yamaji et al. (2005) demonstrated that APC has antioxidant properties and inhibits lipid peroxidation and advanced glycation end products formation. Thus, APC may prevent ROS-related chronic disorders including atherosclerosis and diabetes as well as acute shock conditions. Finigan et al. (2005) reported that APC inhibits thrombin-induced TER reductions and increases cortical MLC phosphorylation which is related to EC barrer protection. APC receptor ligation induces sphingosine 1-phosphate receptor transactivation, thereby resulting in EC cytoskeletal rearrangement and barrier protection. Thus, APC may restore vascular integrity after edemagenic agonists and treatment of patients with severe sepsis (Finigan et al., 2005).

VII. MOLECULAR TARGETS OF ROS REGULATING ENDOTHELIAL PERMEABILITY Tyrosine phosphorylation of VE-cadherin and AJs components through ROS is at least due to activation of kinases, Ca2+ channels and oxidative inactivation of phosphatases, which may contribute to ROS-dependent in crease in endothelial permeability. Several molecular targets of ROS involved in regulating EC permeability are listed as shown below (Figure 1). cSrc Src-family tyrosine kinases play a pivotal role in the disorganization of cadherin-dependent cell–cell contacts and have been shown as targets of ROS.

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Src associates with VE-cadherin and induces tyrosine phosphorylation of VEcadherin on Y685 in angiogenic tissues (Wallez et al., 2007). Csk, a negative regulator of Src family kinase, binds to pTyr685 of VE-cadherin, which is involved in cell density-dependent inhibition of cell growth (Baumeister et al., 2005). The Src family kinases, c-Src, Fyn, Yes bind to p120-catenin and modulate the stability of AJs; so does the kinase Fer (Kim and Wong, 1995). Tyrosine phopshorylation of β-catenin at Tyr142 by Fer or Fyn interferes with its binding to α-catenin, which in turn results in loss of AJs (Piedra et al., 2003; Ozawa and Kemler, 1998). Src-dependent phosphorylation of the RacGEF Vav2 is involved in PAK-mediated phosphorylation of VE-cadherin on Ser665, which contributes to β-arrestin-mediated endocytosis of VE-cadherin in response to VEGF (Gavard and Gutkind, 2006). This response is involved in loss of cell–cell contacts and increase in barrier permeability. Resveratrol, a polyphenolic compound found in grapes, prevents VEGF-induced ROS production, thereby inhibiting Src activation and tyrosine phosphorylation of VE-cadherin and β-catenin (Lin et al., 2003). Given that resveratrol promotes retention of VE-cadherin at cell–cell contacts, this may due to inhibition of ROS-dependent increase in endothelial permeability. Pyk2 Pyk2 is redox-sensitive, Ca2+ -dependent protein tyrosine kinase and it is activated and recruited to cell–cell junctions following the loss of VE-cadherin homotypic adhesion. Inhibition of Pyk2 activity by the expression of CRNK (CADTK/CAKbeta-related non-kinase), an N-terminal deletion mutant that acts in a dominant negative fashion, not only abolishes the increase in β-catenin tyrosine phosphorylation but also prevents the loss of endothelial cell–cell contact (van Buul et al., 2005). Loss of VE-cadherin function promotes Rac-mediated production of ROS, which activate Pyk2 which induce tyrosine phosphorylation of β-catenin, thereby reducing cell–cell adhesion in ECs. This Rac-ROS-Pyk2mediated signaling is important for modulation of endothelial integrity. FAK H2 O2 stimulates tyrosine phosphorylation of FAK, paxillin, beta-catenin, and VE-cadherin as well as distribution of cell–cell adhesions, which in turn decreases TER in bovine pulmonary artery ECs (Usatyuk and Natarajan, 2005). These H2 O2 -mediated responses are inhibited by overexpression of FAK-related non-kinase (FRNK), suggesting that FAK is a target of H2 O2 to transfer ROSdependent signal to endothelial barrier function by regulating cell–cell contacts. Furthermore, low dose H2 O2 increases FAK production in pulmonary microvascular ECs, which may contribute to H2 O2 -mediated cytoskeletal reorganization and cell growth/proliferation (Yang et al., 2004). In rat lung microvascular EC, H2 O2 elicits biphasic responses; initial decrease in TER, followed by recovery to baseline, indicating opening and resealing effects of H2 O2 on EC junction.

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FAK mutant that inhibits FAK activity blocks H2 O2 -induced junctional recovery effects without affecting initial junctional separation (Quadri and Bhattacharya, 2007). Thus FAK rather plays a critical role in H2 O2 -induced remodeling of AJs to reseal the barrier in these ECs. Integrin-induced ROS induce oxidative inactivation of the low molecular weight PTPs, thereby increasing phosphorylation of FAK. Accordingly, FAK phosphorylation and other downstream events, including activation of MAPK and Src, focal adhesion formation, and cell spreading, are all attenuated by inhibition of ROS (Chiarugi et al., 2003). Thus, redox circuitry exists whereby cell adhesion induces oxidative inhibition of PTPs to promote phosphorylation of the downstream signaling of FAK, which results in increasing cell matrix adhesion and spreading onto fibronectin. p21-Activated Kinase (PAK) PAK is phosphorylated on Ser141 following Rac1 activation, and p-PAK translocates to endothelial cell–cell junctions in response to serum, VEGF, bFGF, TNFalpha, histamine, and thrombin. PAK activation and translocation increase permeability across the cell monolayer in response to these factors. Permeability correlates with PAK-induced myosin phosphorylation, formation of actin stress fibers, and the appearance of paracellular pores. Inhibition of myosin phosphorylation blocks the increase in permeability (Stockton et al., 2004). These data suggest that PAK is a central regulator of endothelial permeability induced by multiple growth factors and cytokines via an effect on cell contractility. Thus, PAK may be a potential therapeutic target for the treatment of pathophysiologies which are dependent on vascular leak, such as ischemia and inflammation. Given that PAK is downstream of Rac and phosphorylates p47phox (Knaus et al., 1995; Martyn et al., 2005), PAK-mediated responses may be mediated through ROS. c-Jun N-Terminal Kinase (JNK) TNF-alpha induces complex signaling events in ECs, leading to inflammatory gene transcription and junctional permeability increases. TNFα-induced H2 O2 activates JNK by inhibiting IKK (Pantano et al., 2003). Involvement of TRAF4 and p47phox of NADPH oxidase in oxidative activation of JNK in response to TNFα has been also reported in ECs (Xu et al., 2002; Gu et al., 2002; Li et al., 2002). TNF activates RhoA and Rho kinase, which is required for JNKdependent IL-6 production and increase in permeability in human pulmonary microvascular ECs (Mong et al., 2008). Knockdown of the basic-region leucine zipper protein, c-Jun, by a catalytic DNA molecule, Dz13, suppresses vascular permeability and transendothelial emigration of leukocytes in murine models of vascular permeability, inflammation, acute inflammation and rheumatoid arthritis (Fahmy et al., 2006). Mechanistic studies showed that Dz13 blocks cytokineinducible endothelial c-Jun, E-selectin, ICAM-1, VCAM-1 and VE-cadherin expression but has no effect on JAM-1, PECAM-1, p-JNK-1 or c-Fos. These findings implicate c-Jun as a potential target for anti-inflammatory therapies.

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Protein Kinase C (PKC) PKC is activated in response to various inflammatory mediators and contributes significantly to the endothelial barrier breakdown (Lynch et al., 1990). PKC activation can directly increase permeability of macromolecules across the endothelial or epithelial barriers by phosphorylating cytoskeletal proteins including caldesmon, vimentin, talin and vinculin (Stasek Jr. et al., 1992; Yuan et al., 2007), or by regulating expression or activity of various growth factors including VEGF (Yuan et al., 2007). PKC-mediated endothelial barrier regulation is in part dependent on caveolin-1 (Waschke et al., 2006). The Ca2+ -dependent PKC isoform PKCα plays a crucial role in initiating endothelial cell contraction and disassembly of VE-cadherin junctions. PKC phosphorylates caldesmon77 and vimentin, an event which occurs in concert with agonist-mediated EC contraction, which enhances albumin permeability and barrier dysfunction in bovine pulmonary artery ECs (Stasek Jr. et al., 1992). PKC is a target for redox modification by oxidants and antioxidants. Oxidants selectively oxidize N-terminal regulatory domain contains zinc-binding, cysteine-rich motifs and thus stimulate PKC activity. In contrast, antioxidants such as selenocompounds, polyphenolic agents such as curcumin, and vitamin E analogues react with C-terminal catalytic domain that contains several reactive cysteines, thereby inhibiting PKC activity (Gopalakrishna and Jaken, 2000). In ECs H2 O2 increases PKC activity and diacylglycerol formation (Taher et al., 1993), thereby promoting endothelial permeability (Siflinger-Birnboim et al., 1992). Myosin Light Chain Kinase (MLCK) Inflammatory mediators-induced actin reorganization requires activation of both MLCK and PKC. The high molecular weight endothelial MLCK isoform is stably associated with a complex containing p60srcand cortactin, an actin-binding protein and downstream of p60src. Cortactin-EC MLCK interaction plays an important role in endothelial cortical actin-based cytoskeletal rearrangement (Dudek et al., 2002). H2 O2 -induced MLC phosphorylation and actin rearrangement are dependent on MLCK in EC (Zhao and Davis, 1998). Phosphatase Protein tyrosine phosphorylation is controlled by the tightly regulated balance between protein tyrosine kinases and PTPs (Tonks, 2005). PTPs are considered as negative regulators for signaling process initiated by PTK activation. The reversible oxidative inhibition of PTPs by ROS is an important mechanism through which ROS increase tyrosine phosphorylation signaling events (Lee et al., 1998; Finkel, 1999; Meng et al., 2002; Chiarugi and Cirri, 2003). The PTP activity is dependent on the reactive cysteine residues (Cys-SH) with a low pKa at their active site (Zhang and Dixon, 1993; Lohse et al., 1997) that are readily susceptible to reversible oxidation by H2 O2 (Wu et al., 1998). Kinases and phosphatases cooperate at sites of cell–cell adhesion, thereby regulating AJ integrity or disas-

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sembly both in the steady sate and in response to inflammatory agents (Shasby, 2007). They are recruited to the cell surface through interaction with AJs components, which in turn providing spatiotemporal control of cell–cell adhesion dynamics by modulating interactions between cadherin and catenins. A complex interdependence between AJs components and the activities of kinases and phosphatases contributes to the establishment, maintenance, and disassembly of cell–cell contact in the endothelium. The tyrosine phosphatase inhibitor, sodium orthovanadate induces MLC tyrosine phosphorylation, which decreases electrical resistance across bovine EC monolayers and increases albumin permeability (Gilbert-McClain et al., 1998). Diperoxovanadate stimulates tyrosine phosphorylation, phosphorylation of Src and cortactin, actin remodeling, and endothelial barrier dysfunction in BPAECs through increase in cytosolic Ca2+ (Usatyuk et al., 2003). PTP1B interacts with cadherin and dephosphorylates β-catenin at Tyr654, which is involved in stabilizing the cadherin-β-catenin complex (Xu et al., 2004). Phosphorylation of PTP1B on Tyr152 by Fer is required for interaction with cadherin (Xu et al., 2004). Thus, the presence of both PTP1B and Fer in the cadherinbeta-catenin-120-catenin complex increases the integrity of AJs in the basal state. Endothelial receptor-type phosphatases including Vascular endothelial (VE)-PTP, PTPmu and PTPmicro associate with VE-cadherin through extracellular domain, which inhibits tyrosine phosphorylation of VE-cadherin and stabilizes AJs, independently of its enzymatic activity (Nawroth et al., 2002; Hellberg et al., 2002; Sui et al., 2005). SHP2 associates selectively with β-catenin in VE-cadherin complexes (Ukropec et al., 2000). Thrombin promotes SHP2 tyrosine phosphorylation and dissociation from VE-cadherin complexes. Thus, thrombin regulates tyrosine phosphorylation of VE-cadherin-associated β-catenin, and p120-catenin by modulating the quantity of SHP2 associated with VE-cadherin complexes. Such changes in adherens junction complex composition likely underlie thrombininduced increase in endothelial monolayer permeability. Confluent ECs respond poorly to the proliferative signals of VEGF (Grazia Lampugnani et al., 2003), which is due to the expression of VE-cadherin. In sparse cells or in VE-cadherin-null cells, this phenomenon does not occur and the receptor is fully activated by the growth factor. VE-cadherin truncated in beta-catenin but not p120 binding domain is unable to associate with VEGFR2, and beta-catenin-null ECs are not contact inhibited by VE-cadherin and are still responsive to VEGF. Thus, VE-cadherin-beta-catenin complex participates in contact inhibition of VEGF signaling. Upon stimulation with VEGF, VEGFR2 associates with the complex at cell–cell contacts, where VEGFR2 signaling is inactivated by high cell density-enhanced PTP 1 (DEP-1)/CD148. Thus, it is likely that agonist-induced ROS may inhibit PTPs which negatively regulates tyrosine phosphorylation of VE-cadherin-containing AJs complex, thereby promoting junctional separation and increase in EC permeability.

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Ca2+ Channels Inflammatory mediators stimulate endothelial permeability by increasing the intracellular Ca2+ concentration ([Ca2+ ]i). The rise in [Ca2+ ]i activates key signaling pathways that mediate cytoskeletal reorganization (through myosinlight-chain-dependent contraction) and the disassembly of VE-cadherin at the AJs. Recent studies have identified members of Drosophila transient receptor potential (TRP) gene family of channels that encode functional store-operated cation channels (SOCs) in ECs. Canonical TRP channel (TRPC) homologue TRPC1 is the predominant isoform expressed in human vascular ECs. The malfunction and dysregulation of TRPCs (TRPC1, TRPC4, TRPP1, TRPP2, TRPV4) is associated with endothelial barrier dysfunction. Thus, endothelial TRP channels play an important role in signaling agonist-induced increases in endothelial permeability (Tiruppathi et al., 2006; Ahmmed and Malik, 2005; Kwan et al., 2007). PKCa phosphorylation of TRPC1 is important for Ca2+ Ca2+ entry in human ECs and is involved in thrombin-induced increase in EC permeability (Ahmmed et al., 2004). RhoA induces association of IP3 receptor with TRPC1, thereby activating SOCs and resultant increase in endothelial permeability (Mehta et al., 2003). Vanilloid TRPC, TRPV4 is involved in disruption of the alveolar septal barrier and pathogenesis of acute lung injury (Alvarez et al., 2006; Townsley et al., 2006). H2 O2 activates ROS sensitive channel, transient receptor potential melastatin (TRPM)2, an oxidant-activated channel belonging to the TRP family of cation channels, thereby increasing [Ca2+ ]i and endothelial permeability (Hecquet et al., 2007) (Figure 1). Platelet Endothelial Cell Adhesion Molecule-1 (PECAM-1, CD31) PECAM-1 is a critical modulator of neutrophil-EC transmigration. On stimulation by a vasoactive substance or shear stress, PECAM-1 is tyrosine phosphorylated, enabling recruitment of SHP-2 and tyrosine-phosphorylated β-catenin to its cytoplasmic domain, facilitating dephosphorylation of β-catenin to reconstitute the adherens junctions. In addition, PECAM-1 modulates the levels of β-catenin by regulating the activity of GSK-3β, which in turn affects the serine phosphorylation of β-catenin and its proteosomal degradation, regulating the reformation of AJs (Biswas et al., 2006). PECAM-1-null mice exhibit prolonged and increased permeability after inflammatory insults. In PECAM-1-null ECs, β-catenin is tyrosine phosphorylated, coinciding with a sustained increase in permeability (Biswas et al., 2006). SHP-2 association with β-catenin is diminished; β-catenin/GSK-3β association and β-catenin serine phosphorylation levels are increased; and GSK3β serine phosphorylation (inactivation) in response to histamine or shear stress is blunted. These suggest that lack of PECAM-1 prevents the dephosphorylation of AJs component, promoting a sustained increase in permeability. Thus, PECAM-1 serves as a critical dynamic regulator of endothelial barrier permeability. Of note, ROS support the PECAM-1/SHP-2 formation and induce oxidative inactivation of SHP-2, thereby promoting tyrosine phosphorylation of PECAM-1 (Maas et

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al., 2003). In HUVEC and REN cells (a PECAM-1-negative EC-like cell line) stably transfected with PECAM-1, H2 O2 activates a calcium-permeant, nonselective cation current, and a transient rise in [Ca2+ ]i. Thus, PECAM-1 may function as a sensor of oxidative stress during the process of inflammation.

VIII. MEDIATORS/REGULATORS OF ROS-DEPENDENT ENDOTHELIAL PERMEABILITY IQGAP1 IQGAP1, an F-actin cross-linking protein, interacts with GTP-activated Cdc42 and Rac, and stabilizes AJs through its effects on actin polymerization by actin cross-linking. IQGAP1 binds to the amino-terminal domain of β-catenin and competes for binding at this site with α-catenin (Kuroda et al., 1998). Active Cdc42 and Rac compete with β-catenin for binding IQGAP1 and prevent IQGAP1 from displacing α-catenin (Fukata et al., 1999). Since recent reports suggest that α-catenin does not link the cadherin complex to actin (Drees et al., 2005; Yamada et al., 2005), the functional significance of IQGAP1 displacing α-catenin from the cadherin complex remain uncertain. Knockdown of IQGAP1 or Rac1 with siRNAs reduces VE-cadherin-mediated cell–cell adhesion, and the effects of reducing active Rac1 are overcome by overexpressing IQGAP1. These indicate that IQGAP1 supports cadherin adhesion probably through promoting actin polymerization (Noritake et al., 2005). GTP-activated Rac1 and Cdc42 bind IQGAP1 that has anti-GTPase activity, sustaining the active state of Rac1 and Cdc42. Their interaction inhibits cadherin endocytosis by affecting actin polymerization (Izumi et al., 2004; Sakisaka et al., 2004). IQGAP also transiently interacts with cytoplasmic linking proteins CLIP-170 and CLIP-115 and with adenomatous polyposis coli (APC) at the growing ends of microtubules (Gundersen, 2002; Akiyama and Kawasaki, 2006). Thus, the microtubule cytoskeleton might also contribute to maintenance of Rac1 and Cdc42 activity at the AJs through IQGAP1. IQGAP1 binding is also affected by intracellular calcium concentration. At low, physiological intracellular calcium, IQGAP1 binds Cdc42 and Rac and stabilizes actin. High intracellular calcium displaces IQGAP1 from Cdc42 and Rac, and from β-catenin to calmodulin. This repositioning of IQGAP1 would destabilize cortical actin (Jaffer and Chernoff, 2004). In unstimulated HUVEC confluent monolayers, IQGAP1 colocalizes with VEcadherin at the sites of cell–cell contacts and VEGF stimulation reduces the junctional staining of VE-cadherin and IQGAP1 (Yamaoka-Tojo et al., 2006). IQGAP1 siRNA inhibits localization of VE-cadherin at cell–cell junction, suggesting that IQGAP1 is required for establishment of EC integrity in quiescent ECs (Yamaoka-Tojo et al., 2006). VEGF stimulation promotes IQGAP1 binding to the active VEGFR2 which associates with VE-cadherin/α-catenin complex, which in turn stimulates recruitment of Rac1 to the IQGAP1 containing

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junctional complex (Carmeliet et al., 1999). Moreover, IQGAP1 siRNA inhibits VEGF-induced ROS production and VE-cadherin tyrosine phosphorylation, suggesting that IQGAP1 functions as a scaffold protein to link VEGFR2 to the VE-cadherin-containing complex at AJs, thereby promoting Rac/ROS-dependent tyrosine phoshorylation of VE-cadherin. This mechanism may be required for VEGF-stimulated loss of cell–cell contacts and activation of downstream redox signaling events. By contrast, in ECs in which Rac1 functions to stabilize AJs junction, the cadherin-catenins complex may promote IQGAP1-mediated activation of Rac1, which in turn activate p190RhoGAP. Consistent with this idea, depletion of either IQGAP1 or Rac1 destabilizes AJs, whereas overexpression of IQGAP1 restores AJs in Rac1-depleted cells, possibly through a Cdc42-mediated pathway (Noritake et al., 2004). Microtubules (MTs) Barrier dysfunction of pulmonary endothelial monolayer is associated with dramatic cytoskeletal reorganization, activation of actomyosin contractility, and gap formation. MT disassembly by nocodazole initiates specific signaling pathways that cross-talk with microfilament networks, resulting in Rho-mediated EC contractility and barrier dysfunction (Verin et al., 2001; Birukova et al., 2004a, 2004b, 2004c). Rho-mediated activation of Rho-kinase induces phosphorylation of MLC phosphatase (MYPT1) at Thr(696) and Thr(850), resulting in MYPT1 inactivation. Rho-kinase and MYPT1 are major Rho effectors mediating pulmonary EC barrier disruption in response to MT disassembly. This in turn promotes accumulation of diphospho-MLC which induces acto-myosin polymerization, stress fiber formation and gap formation (Birukova et al., 2004d). Thus, Rho plays an important role in cross-talk between the MT and actomyosin cytoskeleton. MT disassembly-induced endothelial barrier dysfunction is attenuated by protein kinase A (Birukova et al., 2004d). GEF-H1, an MT-associated Rho-specific guanosine nucleotide (GDP/GTP) exchange factor, is involved in Rho activation, MLC phosphorylation, and actin remodeling as well as EC barrier dysfunction induced by thrombin or MT disassembly (Birukova et al., 2006). Thus, GEF-H1 is a key molecule involved in cross talk between MT and actin cytoskeleton in agonist-induced Rho-dependent EC barrier regulation. Importantly, thrombin itself induces MT disassembly through G12/13p115RhoGEF-Rho-Rho kinase activation, thereby promoting EC barrier dysfunction (Birukova et al., 2004d). G12/13-mediated Rho activation is also mediated through PKCα- or Tec-mediated phosphorylation of p115RhoGEF (Holinstat et al., 2003; Mehta et al., 2003; Suzuki et al., 2003). Nocodazole decreases ROS production and NADPH oxidase activity (Devillard et al., 2006), suggesting that MTs play an important role in ROS-mediated EC barrier dysfunction. IQGAP1, an actin cross-linking protein and a downstream target of active Rac1 and Cdc42, also transiently interacts with MTs-binding protein CLIP-170 and adenomatous polyposis coli (APC) at the growing ends of microtubules (Gundersen, 2002;

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Akiyama and Kawasaki, 2006). Thus, IQGAP1 captures and stabilizes MTs through the microtubule-binding protein CLIP-170 near the cell cortex, thereby establishing polarized cell morphology and linking MTs and actin cytoskeleton. Caveolin-1 (Cav1) Endothelial barrier function is controlled by transcellular pathway which transports albumin and other macromolecules via endothelial caveolae as well as by paracellular pathway. Caveolin-1 (Cav1), the primary structural protein required for the formation of caveolae, is important for vesicular trafficking and functions as a negative regulator for NO-mediated cell signaling events (Minshall and Malik, 2006). Lung derived from Cav1-/- mice show marked attenuation of LPS-induced neutrophil sequestration and inhibition of microvascular barrier breakdown and edema formation, which is due to enhanced eNOS-derived NO production and subsequent inhibition of NF-kB (Garrean et al., 2006). LPS also induces NF-kB-dependent Cav-1 expression that results in increased caveolae number, which contributes to increased transendothelial albumin permeability in human lung microvascular ECs (Tiruppathi et al., 2007). Study using Cav1-/PMN reveals that Cav1 expression in PMNs plays a role in mediating PMN activation, adhesion, and transendothelial migration and in PMN activation-induced lung injury (Hu et al., 2007). Furthermore, Cav1 interacts with TRPC1 and this interaction is required for thrombin-induced Ca2+ influx via store-operated Ca2+ channels in human pulmonary artery ECs (Kwiatek et al., 2006). Since Cav1 is required for activation of NADPH oxidase in vascular cells (Zuo et al., 2005), it is temping to speculate that Cav1-mediated endothelial barrier dysfunction may be mediated through NADPH oxidase-derived ROS.

IX. FUNCTIONAL SIGNIFICANCE OF ROS-DEPENDENT ENDOTHELIAL PERMEABILITY IN VIVO Acute Lung Injury (ALI) and Acute Respiratory Distress Syndrome (ARDS) Lung edema due to increased vascular permeability is a hallmark of ALI and ARDS. ALI is characterized by the influx of protein-rich edematous fluid into the airspaces due to increased permeability of the alveolar-capillary barrier. Inflammatory mediators play a critical role in the pathogenesis of this disorder. ARDS has been defined as a severe form of ALI, featuring pulmonary inflammation and increased capillary leak (Ware and Matthay, 2000). A well-described pathophysiologic model of ARDS is one form of acute lung inflammation mediated by inflammatory cells and oxidative stress (Demling et al., 1995). Imbalance in the oxidant–antioxidant system has been recognized as one of the first events that ultimately lead to inflammatory reactions in the lung (Crapo et al., 2003). The generation of oxidants induced by lipopolysaccharide (LPS) has an important signaling function in ECs (Fan et al., 2002; Lo et al., 1993; Rahman et al., 1999).

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Oxidants mediate stable ICAM-1 expression-dependent endothelial adhesivity resulting in the arrest of PMNs (Issekutz et al., 1999). ICAM-1-dependent PMN binding to ECs and EC activation are critical in microbial killing, but they can also mediate lung injury (Horgan et al., 1991) and tissue edema (Lo et al., 1992; Horgan et al., 1991), the hallmarks of ALI associated with severe sepsis. Sepsis is a major factor contributing to ALI resulting from ischemia-reperfusion and or infection. The bacterial overload seen with sepsis is mainly due to gramnegative bacteria that carry LPS. LPS can induce septic conditions as well as induce ROS production. Gram-negative sepsis-induced increase in lung microvascular permeability is inhibited while lung PMN sequestration and transalveolar PMN migration are increased in p47phox-/- and gp91phox-/- mice (Gao et al., 2002). These results suggest that ROS derived from NADPH oxidase is required for sepsis-induced lung microvascular injury, impairment of chemokine generation and lung tissue PMN infiltration. DeLeo et al. (1998) demonstrated that LPS rendered neutrophils more responsive to other stimuli as a result of increased translocation of Rac2, p47phox, and p67phox (i.e., “priming”). Sanlioglu et al. (2001) also reported that LPS induced Rac1-dependent ROS production and TNFα secretion in macrophages. Experiments using PMN from p47phox-/- and gp91phox-/- mice demonstrate that oxidant signaling by the PMN NADPH oxidase complex is an important determinant of TNFα-induced NF-kappaB activation and ICAM-1 expression in ECs (Fan et al., 2002). ROS generated by the PMN NADPH oxidase up-regulate the cell surface expression of the pathogen-associated molecular pattern recognition receptor, Toll-like receptor 2 (TLR2), in ECs (Fan et al., 2003). PMN accumulation depends on TLR4 expression by ECs rather than neutrophils since PMN binding is reduced in EC TLR4-/- mice (Andonegui et al., 2003). TLR4 signaling augments chemokine-induced neutrophil migration by modulating cell surface expression of chemokine receptors (Fan and Malik, 2003). TLR4 signaling also induces TLR2 expression in ECs via PMN NADPH oxidase (Fan et al., 2003). TLR4 expression in ECs is increased under inflammatory conditions. Moreover, in coronary atherosclerotic plaques, TLR4 colocalizes with the p65 subunit of NF-kB (Xu et al., 2001). Thus, expression of pattern recognition receptors such as TLR2 and TLR4 by ECs is regulated by ROS generated by PMN NADPH oxidase, which may contribute to ALI. Most recently, Zhao et al. (2006) reported that the forkhead box M1 (FoxM1) transcription factor induces endothelial regeneration, thereby restores endothelial barrier function after ALI. These findings suggest that the promotion of endothelial regeneration may be a novel therapeutic strategy for ALI (Figure 2). Diabetes mellitus Microvascular barrier injury has been implicated in the initiation and progress of end organ complications of diabetic mellitus. Plasma leakage and fluid retention are seen in various tissues of diabetic patients or animals at the early

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stages of the disease before structural microangiopathy can be detected. Studies have shown that hyperglycemia, often accompanied with insulin deficiency or insulin resistance, can alter the glycocalyx structure and increase permeability in microvessels. Multiple molecular pathways have been identified as contributors to the altered fluid homeostasis, including increased polyol flux that promotes oxidative stress, advanced glycation that leads to carbonyl stress, and excessive glucose metabolism that results in PKC activation. These abnormal metabolic activities are associated with the production of pro-inflammatory cytokines and growth factors, which can stimulate various signaling reactions and structural changes at the endothelial barrier and ultimately cause microvascular leakage (Yuan et al., 2007). Interventions that manipulate these metabolic and inflammatory pathways delay the progress of diabetic microvascular complications. Understanding the molecular basis of diabetes-induced endothelial barrier dysfunction will provide a framework for the development of new therapeutic targets to treat this chronic and debilitating disease process. Given that AGEs which play an important role in diabetes produce ROS (Yan et al., 1994; Schmidt et al., 1994) and induce proinflammatory cytokines (Goh and Cooper, 2008) and that PKC is activated by ROS (Gopalakrishna and Jaken, 2000; Taher et al., 1993), it is likely that ROS are involved in endothelial permeability and dysfunction in diabetes mellitus (Figure 2).

X. SUMMARY AND CONCLUSIONS ROS generated by activated PMN and ECs via activation of NADPH oxidase at site of inflammation and injury in response to inflammatory cytokines, growth factors, G-protein coupled receptor agonists, hypoxia/reoxygenation, shear stress and AGE play an important role in loss of cell–cell junction and leukocyte transendothelial migration. These responses contribute to various disease processes associated with endothelial barrier dysfunction such as ALI/ARDS, diabetes mellitus and atherosclerosis. Mechanically, ROS derived from NADPH oxidase activate various redox signaling pathways through regulating kinases and phosphatases as well as Ca2+ channels, which in turn modulate phosphorylation of AJs complex proteins and actin cytoskeleton. Understanding these mechanisms may provide insights into the ROS, NADPH oxidase and oxidant signaling components as potential therapeutic targets for the treatment of many human diseases that are characterized by vascular leakage. The future will rely on multiple experimental approaches, including cell biology, biochemistry, imaging, structural analysis, and molecular approaches, both in vitro and in vivo. Acknowledgements This work as supported by NIH ROI HL077524 (to M.U.-F), AHA Grant-in-Aid 0555308B (to M.U.-F) and 0755805Z (to M.U.-F), and NIH HL60678 (to A.B.M), NIH ROI HL45638 (to

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A.B.M), NIH POI HL077806 (to A.B.M) T32 HL 07829 (to A.B.M), T32GM070388 (to A.B.M), ROI HL090152 (to A.B.M).

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CHAPTER 9 Cell Signaling by Oxidants: Pathways Leading to Activation of Mitogen-activated Protein Kinases (MAPK) and Activator Protein-1 (AP-1) Arti Shukla and Brooke T. Mossman University of Vermont College of Medicine, Department of Pathology, 89 Beaumont Avenue, Burlington, VT 05405, USA

I. Introduction II. MAPK Signaling A. General Principles of MAPK Classification B. MAPK Signaling by H2 O2 and Generating Systems of ROS/RNS C. MAPK Signaling by Particles and Fibers Inducing Oxidative Stress III. Mitogen-activated Protein Kinase Phosphatases (MKPs) A. Effects of ROS/RNS on MKPs B. MKP Activation/Inactivation by Particles and Fibers IV. Relationships between MAPK and Activator Protein-1 (AP-1) A. General Background B. AP-1 Regulation by Asbestos and other ROS/RNS Generating Particles V. Conclusions References

Abstract Agents inducing oxidative stress activate one or more pathways of the MitogenActivated Protein Kinases (MAPK), a family of signaling proteins leading to activation of the Fos/Jun protooncogene family that comprise the Activator Protein-1 (AP-1) transcription factor. Redox sensitive molecular targets usually contain highly conserved cysteine residues, and their oxidation, nitration and formation of disulfide links are crucial events in oxidant/redox signaling. Oxidation of sulfide groups in signaling proteins causes structural modifications, resulting in the exposure of active sites and protein activation. Thiol proteins such as intracellular glutathione (GSH) and thioredoxin (Trx) are of central importance in the regulated Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00209-3

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control of redox signaling pathways by reducing disulfide bridges or oxidized cysteine residues. We review here how oxidants interact with membrane receptors, other proteins, and Mitogen-Activated Protein Kinase Phosphatases (MKPs) to activate MAPK and AP-1 regulated gene expression critical to cell differentiation, proliferation, death/apoptosis, and disease. These pathways have been studied extensively in cells in culture after exposure to H2 O2 , generating systems of reactive oxygen or nitrogen species (ROS/NOS), and a number of naturally occurring oxidant stresses such as chemotherapeutic drugs or airborne pollutants. The epithelial cell of the respiratory tract and inhalation models of lung injury, inflammation, carcinogenesis, and fibrogenesis have been employed by us and others to study MAPK and AP-1 activation by soluble oxidants and insoluble particulates (asbestos, silica, particulate matter or PM) inducing oxidative stress. Responses of cells to these oxidant stresses are critical to injury or repair, inflammation, and critical outcomes of disease.

I. INTRODUCTION Free radicals are defined as molecules containing one or more unpaired electrons. Radicals derived from oxygen are known as reactive oxygen species (ROS) and those derived from nitrogen are called reactive nitrogen species (RNS). These radicals are generated from both endogenous and exogenous sources such as: (i) irradiation by UV light, X-rays and gamma-rays; (ii) by-products of metal catalyzed reactions; (iii) drugs and drug metabolites; and (iv) pollutants including oxidant-generating gases and particulates in the atmosphere. Endogenous sources of ROS/RNS include mitochondria, cytochrome P450 metabolism, peroxisomes and inflammatory cell activation (Inoue et al., 2003). The H2 O2 molecule does not contain an unpaired electron and thus is not a radical species per se, but it is studied because of its high reactivity. Although H2 O2 was initially considered as toxic to cells, it is clear that it now plays an important role in redox signaling involved in both physiological and pathological processes such as proliferation, differentiation, migration, angiogenesis, aging and cancer. H2 O2 can be generated directly in cells by some oxidoreductases, such as glucose oxidase (Massey et al., 1969), and the recently described DuOXs (Lambeth, 2002), which are isoforms of the NADPH oxidases. Most H2 O2 production, however, results from the dismutation of superoxide (O·2− ) produced by NADPH oxidases (Lambeth, 2002), leakage from the mitochondrial electron transport chain (Forman and Kennedy, 1974; Loschen et al., 1974), and redox cycling of xenobiotic quinones (McCord and Fridovich, 1970) and flavoproteins (Inoue et al., 2003). H2 O2 is a small and non-charged molecule that is readily produced and removed following physiological stimuli and can thus cross membranes, exhibiting properties that are ideal for signal transduction (Rhee, 2006).

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In nonphagocytic cells stimulated with growth factors and cytokines like plateletderived growth factor (PDGF), epidermal growth factor (EGF), insulin, tumor necrosis factor-α (TNFα), and interleukin-1 (IL-1), a major source of H2 O2 is gp91 Phox and its homologs (Lambeth, 2004). H2 O2 exerts its critical functions in physiological signaling events by causing reversible amino-acid oxidations. A variety of ROS/RNS are known to play dual roles in biological systems, since they can be either beneficial or harmful to living systems. Beneficial effects of ROS involve physiological roles like defense against infectious agents and normal signaling events. In contrast at high concentrations, ROS can be important mediators of damage to membranes, lipids, proteins and nucleic acids. Despite the presence of antioxidant defense systems to counteract oxidative damage from ROS, oxidative damage accumulates during aging, and radical related damage to DNA, proteins and lipids plays a key role in the development of age-dependent diseases such as cancer, arteriosclerosis, arthritis, neurodegenerative disorders, and other pathologies. While ROS/RNS are predominantly implicated in cell damage, they also play a major physiological role in several aspects of intracellular signaling and regulation. Some highly reactive oxidant species like peroxynitrite (ONOO. ), nitrogen dioxide (NO2 ), ozone (O3 ), hypochlorous acid (HOCl), and the hydroxyl radical OH. ), oxidize biomolecules without much preference. These reactive species are capable of inducing protein modifications that lead to activation of signal transduction cascades. The redox status of a cell may evoke concentration dependent responses. For example, low concentrations of O−· 2 and H2 O2 stimulate proliferation and enhanced survival in a wide variety of cell types whereas at higher concentrations they cause growth arrest or cell death. Inhaled airborne particulates also generate oxidative stress by a number of mechanisms that have been linked to their pathogenic potential. The most extensively studied of these particulates are asbestos fibers (defined as having a greater than 3:1 length to diameter ratio), crystalline silica, and ambient particulate matter (PM). Asbestos and silica have been classified as carcinogens based on epidemiological studies in occupational settings, particularly in smokers, whereas PM has only recently been implicated with an increased risk of lung cancer in the general population (Pope et al., 2002). Asbestos is a group of naturally occurring crystalline mineral fibers that are associated with the development of malignant (mesothelioma, lung cancers) and nonmalignant diseases in lung and pleura (asbestosis and pleural fibrosis) (Mossman et al., 1990; Mossman and Gee, 1989). A main factor in determining the biological activity and pathogenicity of various types of asbestos fibers is their ability to participate in redox reactions and diminish expression of antioxidant enzymes or intracellular GSH pools [reviewed in Shukla et al. (2003)]. Asbestos fibers generate ROS and RNS via direct interaction with the membranes of epithelial cells and macrophages after inhalation. Longer fibers that have proven to be more toxic, carcinogenic and fibrogenic in vitro and in vivo induce frustrated

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phagocytosis accompanied by a prolonged oxidative burst. During the process of incomplete phagocytosis of long fibers, O−· 2 is generated, and its dismutation results in the production of H2 O2 . In the presence of transition metal ions, such as ferrous iron or cuprous ions, H2 O2 is converted to the potent oxidizing radical, OH· through a Fenton like reaction. Pathogenic asbestos fibers, particularly high iron containing amphiboles such as crocidolite and amosite, also release ROS by iron catalyzed reactions on the fiber surface (reviewed in Shukla et al., 2003). This process augments the phagocytic generation of ROS and results in further generation of oxidants from extracellular and intracellular fibers. Free radicals generated from asbestos fibers are linked to altered cell signaling, inflammation, and a plethora of other mutational and functional responses associated with the pathogenesis of asbestos-associated diseases. In addition to ROS, asbestos signaling may be triggered by RNS like NO· or ONOO− . For example, crocidolite asbestos is able to induce nitric oxide synthase (iNOS) in rat alveolar macrophages after inhalation (Iguchi et al., 1996; Quinlan et al., 1998) and in rat pleural mesothelial cells in vitro (Choe et al., 1998; Iguchi et al., 1996). Moreover, immunoreactivity for nitrotryosine is increased in lung epithelium and mesothelium after inhalation of crocidolite by rats (Tanaka et al., 1998). Generation of oxidants by crystalline silica after grinding (Donaldson and Tran, 2002) or after interaction with macrophages (Castranova et al., 1991) and epithelial cells (Deshpande et al., 2002) also has been characterized. As reviewed elsewhere, OH· may be generated by reactive silanol groups as well as by traces of transition metals present in these particles (Fubini and Hubbard, 2003). Oxidants generated from silica particles also can cause activation of cell signaling pathways, cell and lung injury, increased expression of inflammatory cytokines and activation of specific transcription factors, including AP-1 as detailed below. Free radicals and oxidative stress also are extensively implicated in the inflammatory effects of PM (Donaldson and Tran, 2002) via metal-generated free radicals (Frampton et al., 1999; Gilmour et al., 1996), ultrafine particle surfaces (Brown et al., 2001), and organic components (Squadrito et al., 2001). Synergistic interactions between transition metals and ultrafine particles (defined as particles <100 nm in diameter) in causing oxidative stress and lung inflammation have been reported (Wilson et al., 2002). Diesel exhaust particles (DEP) also generate ROS after metabolism of organic compounds on their surfaces. A correlation between the presence of polycyclic aromatic hydrocarbons (PAH), induction of metabolic pathways by DEP, and radical generation has been established (Baulig et al., 2003; Marano et al., 2002). In this chapter, we initially present a review of the principles and regulation of MAPK cell signaling by oxidants. Next, we describe how H2 O2 and generating systems of ROS/RNS activate MAPK and AP-1 activity. Finally, we discuss

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FIGURE 1 General schema showing activation of MAPK cascades. Note that stimuli can act through growth factor receptors of G-proteins such as Ras, rendering Ras in an active GTP-bound state that can bind to downstream effectors including Raf (middle panel). G-proteins involved in activation of the p38 and JNK cascades include Rac, Cdc42 and Rho which are not depicted in the diagram.

how inhaled particulates stimulate these pathways and their relevance to fiber and particle toxicology.

II. MAPK SIGNALING A. General Principles of MAPK Classification The MAPKs are serine threonine kinases that are activated through receptor tyrosine kinases, i.e., the epidermal growth factor receptor (EGFR), G proteins, and receptor independent mechanisms. These proteins are phosphorylated via a series of upstream kinases (MEKKK, MEKK and MEK) and dephosphorylated via specific phosphatases (see Figure 1 and Section III below). There are three major MAPK families: extracellular signal-regulated kinases (ERKs), including big MAPK or ERK5 and at least 8 reported other ERKs, c-Jun-NH2 -terminal kinases (JNKs 1-3), and p38 (alpha, beta, gamma and delta) kinases. The most commonly studied and most abundant ERKs in mammals are ERKs 1 and 2, most generally studied together because of the lack of specificity of available antibodies that do not differentiate between the two. JNKs 1 and 2 isoforms are ubiquitous, whereas JNK 3 is tissue specific, commonly found in brain tissue. Recent studies support the concept that most JNK inducers significantly activate JNK1, but not JNK 2, to induce biological effects (Liu et al., 2004).

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Their binding to scaffolding accessory protein, kinetics and duration of activation and inactivation, substrate availability, and subcellular localization may dictate distinct responses to MAPK cascades in individual cell types or after exposure to various types and concentrations of oxidants. Moreover, it is clear that there are integrating signals from receptor tyrosine kinases (RTKs), especially through the Raf, MEK and ERK cascades (McKay and Morrison, 2007) that influence cell outcomes as well as cross-talk between various MAPK cascades and other transcription factors such as Nuclear-Factor-κB (NF-κB) (Liu and Lin, 2007; Raman et al., 2007).

B. MAPK Signaling by H2 O2 and Generating Systems of ROS/RNS Products of NOX1 activity, O·− 2 , and H2 O2 , can activate the MAPK cascade at the level of MEK and ERK1/2. Experimental studies on the upregulation of all three MAPK pathways by H2 O2 have shown that the activation of each signaling pathway is type and stimulus specific. Redox reactions catalyzed by H2 O2 involve the oxidation of cysteine residues on proteins, which consequently affect protein function. H2 O2 also appears to promote tyrosine phosphorylation by activating protein tyrosine kinases. For example, upon cell attachment to extracellular matrix and associated generation of H2 O2 , the tyrosine kinase, Src, becomes oxidized at two cysteine residues and thus becomes activated (Giannoni et al., 2005). Transactivation of EGFR, an upstream regulator of ERK1/2 activity in some cell types, requires basal intracellular levels of H2 O2 in prostate cancer cells, suggesting an endogenous role of oxidants in RTK activation of MAPKs (Zhou, Q. et al., 2006). Endogenous H2 O2 production by the respiratory burst response has been shown to induce ERK1/2 but not p38 kinase activity in alveolar macrophages (Iles and Forman, 2002). In contrast, exogenous administration of H2 O2 activates p38 kinase, but not ERK1/2 (Torres and Forman, 1999). The ERK1/2 pathway has most commonly been associated with growth factor/RTK stimulation and the regulation of cell proliferation; however, the balance between ERK and JNK activation is the key factor for cell survival. For example, in cortical neurons and astrocytes, exposure to H2 O2 triggers activation of the ERK1/2 pathway (along with intracellular Ca2+ accumulation and other MAPK activation), resulting in apoptotic like cell death (Numakawa et al., 2007; Shinozaki et al., 2006). A recent study illustrates the role of JNK1 in H2 O2 induced nonapoptotic cell death via sustained poly (ADP-ribose) polymerase activation (Zhang et al., 2007). However, activation of p38 in lung fibroblasts by repetitive low grade H2 O2 leads to prosurvival effects through its command over down-stream survival elements such as Akt (Protein kinase B) and NF-κB (Sen et al., 2007). A recent study reports activation of ERK1/2 leading to expression of cyclin D1 and cell proliferation by NOX1 in mouse lung epithelial cells (Ranjan et al., 2006). ROS/H2 O2 also have been shown to alter vascular tone in

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hypertensive rat thoracic vena cavas through modulation of p38 and the ERK1/2 pathway (Thakali et al., 2007). The MAPKKK, ASK1 (apoptosis signaling kinase), activates protein kinases upstream of JNK and p38 MAPK (Wang et al., 1996). Exogenous addition of H2 O2 or generation of H2 O2 by a respiratory burst can cause ASK1 activation through Trx dissociation from ASK1 and phosphorylation of ASK1 at Thr845 (Liu et al., 2006). The only known mechanism for dissociation of ASK1 from Trx is the oxidation of Trx by H2 O2 (Ichijo et al., 1997). In contrast, there are reports indicating redox- independent regulation of ASK1 by Trx (Liu and Min, 2002). Oxidants can also activate JNK by oligomerization and dissociation of glutathione S-transferase Pi (GSTp) which has emerged as a critical regulator of the JNK pathway (Adler et al., 1999). In a similar manner, H2 O2 induced p21 protein upregulation is mediated via JNK1 by inhibition of the ubiquitination of p21. This model provides new insight for analyzing the regulatory effect of JNK after oxidative stress (Fan et al., 2007).

C. MAPK Signaling by Particles and Fibers Inducing Oxidative Stress Interaction of pathogenic asbestos fibers (see below) and silica particles (reviewed in Fubini and Hubbard, 2003) with cells trigger numerous signaling cascades including MAPKs and nuclear factor kappa B (NF-κB) in vitro and after inhalation (Cummins et al., 2003; Ding et al., 1999; Haegens et al., 2007; Robledo et al., 2000). In lung epithelial and mesothelial cells, the ERK1/2 pathway is induced by pathogenic fibers such as asbestos and erionite through a mechanism involving phosphorylation of the EGFR (Zanella et al., 1996, 1999) and is accompanied by increased biosynthesis of the EGFR receptor. In addition, asbestos fibers cause dimerization of the EGFR in human mesothelial (MET5A) cells, a process that can also activate ERK1/2 (Pache et al., 1998). Inhibition of EGFR phosphorylation using a small molecule inhibitor (AG1478) abolishes fiber induced ERK1/2 activation, c-fos levels, and apoptosis, a finding in line with more recent studies demonstrating that long duration of ERK1/2 phosphorylation contributes to cell death by asbestos in lung epithelial cells (Yuan et al., 2004) and mesothelial cells which can be inhibited by addition of catalase (Jimenez et al., 1997). In contrast, a shorter duration of EGF-induced ERK1/2 signaling in these cell types causes cell proliferation (Buder-Hoffmann et al., 2001). Subsequent work by our group suggests that ERK1/2 and ERK5 pathways cooperatively contribute to epithelial cell proliferation by asbestos; however, initiation of ERK1/2 is via a Src/EGFR pathway, whereas activation of ERK5 is Src-dependent, but EGFR-independent (Scapoli et al., 2004). The JNK pathway is also activated by asbestos (Buder-Hoffmann et al., manuscript in preparation), silica (Shukla et al., 2001) and PM (Timblin et al., 2002) in rodent lung epithelial cells as is phosphorylation of c-jun in these cell types. We have linked transcriptional activation of

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c-jun and its overexpression to cell proliferation by asbestos or low concentrations of H2 O2 (Timblin et al., 1995), but activation of the JNK pathway may also cause cell death at higher concentrations of oxidants.

III. MITOGEN-ACTIVATED PROTEIN KINASE PHOSPHATASES (MKPS) A. Effects of ROS/RNS on MKPs Protein tyrosine phosphatases (PTPs) are the best characterized direct targets of ROS. The reversible phosphorylation of tyrosyl residues in proteins plays a key role in the various signaling pathways induced by number of stimuli that regulate biological responses including growth, proliferation, metabolism and differentiation. The effects of ROS occur through targeting the cysteine containing residues of the active sites of tyrosine phosphatases (Salmeen and Barford, 2005). Oxidation of the essential cysteine abolishes phosphatase activity and can be reversed by cellular thiols. MKPs play critical roles in mediating the feedback control of MAPK cascades in a variety of cellular processes, including proliferation and stress responsiveness. The MKP family is comprised of 10 dual specificity MKPs, all of which share a common structure comprising an N-terminal non-catalytic domain and a C-terminal catalytic domain (reviewed in Owens and Keyse, 2007). MKP-1 is a dual-specificity protein phosphatase encoded by an immediate-early gene responsive to growth factors and stress. The MKP-1 protein selectively inactivates MAPK in vitro by dephosphorylation of the regulatory Thr and Tyr residues. MKPs also have a reactive cysteine required for catalytic activity, and oxidation of the cysteine in MKPs result in inactivation of the phosphatase activity, leading to enhanced activation of MAPKs, including JNKs and ERKs (Kamata et al., 2005; Seth and Rudolph, 2006). Cysteine residues are most susceptible to oxidative damage by H2 O2 and other oxidants, producing sulfenic acid intermediates which can further react with thiols to form catalytically inactive disulfides. In contrast, underlying and continual oxidative stress in portal hypertensive rats leads to overexpression of MKP-1, and as a result, ERK2 activation is impaired (Kawanaka et al., 2001). Furthermore, studies show MKP-2 terminates H2 O2 -induced JNK activation which is associated with the nucleus, but not by the activation of JNK by TNFα found within the cytoplasm (Robinson et al., 2001). A novel MKP designated MKP-7 has recently been identified. It possesses a long C-terminal stretch containing both a nuclear export signal and a nuclear localization signal and the dual-specificity phosphatase catalytic domain (Masuda et al., 2001). MKP-7 suppresses activation of MAPKs in COS-7 cells in the order of selectivity, JNKs > p38s > ERKs. MKP-1 is an oxidative stress inducible gene, and studies have reported that induction of MKP-1 by H2 O2 is mediated by a MAPK/AP-1 pathway involved

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in cellular defense against oxidative stress induced apoptosis of mesangial cells (Xu et al., 2004). In support of these findings, MKP-1 is induced by oxidative stress, and the activation of MKP-1 is a survival mechanism against oxidative damage (Zhou, J.Y. et al., 2006). However, in neuronal cell lines (SH-SY5Y), MKP-1 plays a proapoptotic role in oxidative stress-induced cell death (Kim et al., 2005). Like ROS, nitric oxide (NO· ) also induces MKP-1 in human monocytes as measured by microarray analyses (Turpaev et al., 2005). NO· also mediates modification of the cysteine residue at the active site of dual-specificity phosphatases (MKP-1, MKP-3) and inactivates them, thereby activating MAPKs under hypoxic conditions (Mishra and Delivoria-Papadopoulos, 2004). Conversely, one report (Mishra and Delivoria-Papadopoulos, 2004) suggests that expression of MKP-1 by NO· leads to dephosphorylation of ERK1/2, an initial and essential event that commits breast cancer cells to the apoptotic pathway in breast cancer cells. Further support that NO· mediates MKP-1 induction and consequent inactivation of MAPKs comes from work using NO/cGMP pathway inhibitors to study insulinmediated inhibition of vascular smooth muscle cell migration (Jacob et al., 2002). NO· induced anti-apoptotic effects in endothelial cells also rely on the down- regulation of MKP-3 mRNA by destabilization and a parallel decrease in MKP-3 protein levels (Rossig et al., 2000).

B. MKP Activation/Inactivation by Particles and Fibers Few studies have addressed the role of MKP induction or inactivation in cells exposed to pathogenic particles or fibers. However, inhibition of tyrosine phosphatases by metals and oxidants has been shown to be important in injury to cells by PM and has been reviewed in a recent volume (Samet and Ghio, 2007). We recently reported that crocidolite asbestos mediated Ca2+ /cAMP-response element binding protein (CREB) is regulated by protein kinase A and ERK1/2 in lung epithelial cells and is accompanied by a sustained increase in MKP-1 mRNA and protein which is cAMP-response element- (CRE) regulated (Barlow et al., 2007). Since the regulation of MKPs by ROS/RNS plays an important role in modulation of MAPK signaling pathways in other cell types, the function of these phosphatases in lung cells after exposure to particulates, is an area ripe for study.

IV. RELATIONSHIPS BETWEEN MAPK AND ACTIVATOR PROTEIN-1 (AP-1) A. General Background The transcription factor AP-1 was one of the first mammalian transcription factors to be identified and is involved in cellular proliferation, transformation

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FIGURE 2 A simplistic schema showing a sequence of interactions after increases in mRNA expression of fos/jun early response genes which then encode proteins that form Fos/Jun family member heterodimers or Jun/Jun family member homodimers that comprise the AP-1 transcription factor. The AP-1 transcription factor binds to genes with an AP-1 site on their promoter to increase or decrease gene transcription leading to DNA replication, cell death, and a number of other outcomes of oxidant exposures.

and death [reviewed in Shaulian and Karin (2002)]. Its relevance to toxicity and carcinogenicity in the respiratory tract by oxidant stresses, including pathogenic particulates and tobacco smoke, has been reviewed (Reddy and Mossman, 2002). In brief, AP-1 is a collection of dimeric basic region-leucine zipper (bZIP) proteins that belong to the Jun (c-Jun, JunB, JunD), Fos (c-Fos, FosB, Fra-1, Fra-2) and ATF subfamilies, all of which can bind the tumor promoter (TPA) or CRE. AP-1 activity is induced by plethora of physiological stimuli and environmental insults, including growth factors, cytokines, neurotransmitters, polypeptide hormones, cell-matrix interactions, bacterial and viral infections, and variety of physical and chemical stresses. Once activated, AP-1 regulates a wide range of cellular processes, including cell proliferation, death, survival, and differentiation. AP-1 activity is regulated by MAPK cascades that enhance AP-1 activity through the phosphorylation of distinct substrates. Activation of members of the MAPK family also leads to the transactivation of other transcription factors such as c-Jun, Activator Transcription Factor-2 (ATF-2), CBP, and Elk-1. AP-1 is a redox sensitive transcription factor and ROS and cellular redox status, particularly intracellular thiol status, can be directly involved in the activation of AP-1 and signal transduction (Lo et al., 1996; Meyer et al., 1993). Thus, alterations in regulation of MAPKs by ROS/RNS (discussed above) can lead to altered AP-1 activity. This eventually culminates in chromatin remodeling and increased ex-

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pression of genes with AP-1 sites in their promoter regions (Figure 2). These gene products are typically involved in inflammation, apoptosis, proliferation, transformation and differentiation. Transcription factors per se can also be direct targets of redox control (Na and Surh, 2006). An example of a transcription factor that is inhibited following cysteine oxidation includes the c-Jun member of the AP-1 family, and both S-Nitrosylation and S-Glutathionylation of c-Jun interferes with its ability to bind DNA (Klatt et al., 1999). The in vitro transcriptional activity of AP-1 is regulated by the redox state of cysteine 64 located at the interface between the two c-Jun subunits, highlighting the importance of redox status on gene transcription. However, recent in vivo experiments demonstrate that cysteine 64/65 is not required for redox regulation of AP-1 to DNA binding in vivo (Whitmarsh and Davis, 1996). Direct oxidative modification of transcription factors may not be the only way to alter their activities as chromatin remodeling factors, such as histone acyl transferases (HAT), or histone deacetylases (HDAC) also can sense redox changes, thereby leading to changes in transcription. Oxidant generating systems and pro-inflammatory mediators influence histone acetylation/phosphorylation via a mechanism dependent on the activation of the MAPK pathway (Bohm et al., 1997; Tikoo et al., 2001). Oxidative stress can also cause overexpression of one or more of the Fos/Jun family members of AP-1, thereby increasing complex formation and binding. For example, oxidative stress causes overexpression of JunD which activates transcription of the human ferritin H gene through the antioxidant response element (ARE) (Tsuji, 2005). A new redox signal transduction pathway in activation of AP-1 has been proposed whereby oxidizing equivalents flow from hepatopoietin (HPO) to Trx and then to substrate protein by dimerization of HPO which then interacts with Trx to activate AP-1 (Li et al., 2005). Several examples from the recent literature further support the role of oxidants in AP-1 regulation in different cell types. For example, increased levels of H2 O2 are observed in zinc deficiency which triggers the activation of JNK, p38 and AP-1 in human IMR-32 cells (Zago et al., 2005). H2 O2 also is important in stimulation of both proliferation and migration of human prostate cancer cells through activation of AP-1 and upregulation of the heparin affin regulatory peptide gene (Polytarchou et al., 2005). Some metals also generate ROS and RNS and activation of signaling pathways are related to activation of AP-1 (Valko et al., 2005). In cardiomyocytes, H2 O2 -induced upregulation of heme oxygenase-1 involved JNKs/p38 MAPK and AP-1 activation (Aggeli et al., 2006). In pancreatic stellate cells, H2 O2 activated AP-1 via the three classes of MAPKs, ERK, JNK and p38 (Kikuta et al., 2006). Activation of one transcription factor by oxidants can also lead to activation of other transcription factors. For example, H2 O2 activates Nuclear Factor of Activated T cells-3 (NFAT3) through an AP-1 transcription factor dependent mechanism (Tu et al., 2007). A definitive role of H2 O2 -induced AP-1 in cell death has

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also been reported (Zhou et al., 2007). In vitro studies using lung epithelial cells and generating systems of toxic concentrations of NO· and ONOO· showed that NO· was able to induce c-fos and c-jun expression and AP-1 binding whereas ONOO· failed to induce these effects (Janssen et al., 1997).

B. AP-1 Regulation by Asbestos and other ROS/RNS Generating Particles Several studies show that asbestos fibers trigger increased expression of early response protooncogenes (fos/jun), Fos/Jun proteins, and AP-1 transactivation. In contrast to a variety of other nonpathogenic fibers and particles, asbestos and erionite fibers cause increased and protracted mRNA levels of fra-1, c-fos, and cjun, and increased AP-1 binding to DNA in lung epithelial and pleural mesothelial cells in vitro (Heintz et al., 1993; Janssen et al., 1994, 1995; Shukla et al., 2001) and after inhalation (Manning et al., 2002). In the latter studies, responses were altered in mice expressing a mutant EGFR that could not be phosphorylated. Studies support a role of oxidants in asbestos-induced AP-1 activation. For example, it was initially suggested that asbestos-induced alterations in cellular thiol status caused subsequent AP-1 activation in rat pleural mesothelial cells (Janssen et al., 1995). These data were supported by work showing ROS generation during the interaction of silica and asbestos with epidermal cells of AP-1 luciferase reporter mice (Ding et al., 1999). Addition of superoxide dismutase and catalase inhibited this activation. Recently, our laboratory has provided evidence that regulation of asbestos-induced EGFR activation, fra-1 transactivation, expression of AP-1 family members, and AP-1 to DNA binding in lung epithelial cells is linked to intracellular levels of glutathione and γ -glutamylcysteine synthetase levels (Shukla et al., 2004), confirming several mechanisms of redox signaling by asbestos-induced oxidative stress. Exposure of tracheal epithelial and mesothelial cells or alveolar macrophages to asbestos causes persistent increases in redox factor-1 (Ref-1 or APE) as well as AP-1 to DNA binding activity that is mediated by oxidative stress (Flaherty et al., 2002; Fung et al., 1998) Treatment of the rat lung fibroblast cell line (RFL-6) with crocidolite asbestos in the presence and absence of the membrane antioxidant, vitamin E, decreases levels of crocidolite-induced AP-1, indicating an involvement of lipid peroxidation (Faux and Howden, 1997). p38 activity, an oxidative stresssensitive effect, is also important for crocidolite-induced AP-1 to DNA binding in pleural mesothelial cells (Swain et al., 2004). Using microarray analysis, our lab has determined time-dependent expression of many signal transduction and oxidative stress genes in murine lungs after inhalation of chrysotile asbestos over a 40 day period (Sabo-Attwood et al., 2005). These include the AP-1 regulated gene, heme oxygenase and matrix metalloproteases (MMPs) (Shukla et al., 2006). There are also reports indicating silica-induced upregulation of the AP-1 transcription factor after intratracheal instillation into mice. For example, increased

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luciferase activity was noted three days after instilling silica into AP-1 luciferase reporter transgenic mice (Ding et al., 1999). Using this same model, we demonstrated AP-1 driven gene expression in lung macrophages and bronchiolar epithelial cells (Hubbard et al., 2001). As described above, PM increases JNK activity in alveolar epithelial cells that is accompanied by increases in phosphorylated c-Jun and transcriptional activation of AP-1 (Timblin et al., 1998). In addition, others have shown stimulation of AP-1 by PM in alveolar epithelial cells (Gilmour et al., 2001). Work from our laboratory demonstrated that PM and ultrafine carbon black at low concentrations upregulated mRNA levels of the early response protooncogenes c-jun, junB, fos related antigen-1 (fra-1) and fra-2 in C10 cells that were associated with cell proliferation (Timblin et al., 2002). Diesel exhaust particles (DEP) also induce JNK activation in mouse lung epithelial cells, resulting in upregulation of fra1 mRNA coding for AP-1 proteins (Zhang et al., 2004). DEP activate fra-1 but not fra-2 mRNA as well as protein expression and activity levels in a time- and dose-dependent manner. Lastly, oxidants in cigarette smoke (CS) also activate AP-1 family member protooncogenes, AP-1 binding, and transactivation resulting in upregulation of AP-dependent genes (reviewed in Mossman et al., 2006). Synthesis of the predominant airway mucin, MUC5AC, was transcriptionally upregulated by CS and was mediated by an AP-1-containing response element binding JunD and Fra-1 (Gensch et al., 2004). Use of ROS scavengers confirmed the involvement of ROS in CS-induced responses. In a recent study, the effects of CS on Jun and Fos family member expression and regulation was demonstrated in the 1HAEo nonmalignant human bronchial epithelial cell line (Zhang et al., 2005). Detailed studies to understand the mechanisms behind CS induced fra-1 showed that CS stimulated fra-1 induction at the transcriptional level, and this effect was suppressed with the use of the EGFR phosphorylation inhibitor, AG1478.

V. CONCLUSIONS Dissecting the MAPK responses by oxidants and how they result in increased AP-1 activity as well as increases or decreases in expression of other transcription factors is important to deciphering how individual cells respond to oxidant stress and embark upon a program of gene expression related to repair from or causation of diseases. The fact that MAPK responses by some oxidants, especially particulates, is mediated at the plasma membrane by receptor interactions or modifications is exciting in terms of both prevention and therapy for a number of cancers and other fibroproliferative diseases. Clearly, membrane-active antioxidants targeting RTKs or other receptors necessary for MAPK activation is a goal for future therapeutic approaches.

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Acknowledgements We thank Jennifer Díaz for preparation of this manuscript, and Maximilian MacPherson for illustrations. Work by our group is supported by P01 HL67004 from the NHLBI, P01 CA11407 from the NCI, and a grant from the Mesothelioma Applied Research Foundation.

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CHAPTER 10 The Interaction of Mitochondrial Membranes with Reactive Oxygen and Nitrogen Species Paul S. Brookes∗ , Andrew P. Wojtovich† , Lindsay S. Burwell‡ , David L. Hoffman‡ and Sergiy M. Nadtochiy∗ ∗ Department of Anesthesiology, University of Rochester Medical Center † Department of Pharmacology & Physiology, University of Rochester Medical Center ‡ Department of Biochemistry & Biophysics, University of Rochester Medical Center

I. Mitochondria as a Source of Reactive Species A. Regulation of Mitochondrial ROS Generation by Trans-membrane Potential (ψm ) B. Mitochondrial NO Synthase C. Mitochondrial ROS & O2 Tension II. Effects of ROS and RNS on Mitochondrial Respiration A. ROS & Respiration B. RNS & Respiration III. Mitochondrial Membrane Lipids A. ROS & Membrane Lipids B. RNS & Membrane Lipids IV. ROS, RNS & Mitochondrial Ion Transport A. Overview B. Ca2+ Uniporter C. Ryanodine Receptor D. K+ ATP Channel E. Permeability Transition (PT) Pore F. ANT G. UCPs V. Complex Interactions & Concluding Remarks References

Abstract Within the context of the overall effects of free radicals (ROS and RNS) on biological membranes, the mitochondrion offers a unique biochemical environment which leads to a number of organelle-specific reactions, This includes a unique Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

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complement of membrane lipids and proteins, an alkaline pH in the matrix, specialized systems for the generation and quenching of ROS and RNS, and a tightly controlled redox status. All of these parameters come together in a highly orchestrated system that is essential to the proper functioning of the mitochondrion. This chapter will examine some of the mitochondria-specific aspects of ROS and RNS reactions with membranes, within the context of the many disease processes that originate at the organelle level.

I. MITOCHONDRIA AS A SOURCE OF REACTIVE SPECIES A full discussion of the molecular mechanisms underlying mitochondrial reactive species generation is beyond the scope of this chapter, and readers are directed to recent reviews on the subject (Adam-Vizi and Chinopoulos, 2006; Brookes et al., 2004). Nevertheless, it is worthwhile to mention some recent developments and controversial debates in the field.

A. Regulation of Mitochondrial ROS Generation by Trans-membrane Potential (ψm ) The precise nature of the relationship between mitochondrial ψ and ROS generation is the subject of some controversy, and there are some nuances associated with the two main ROS generation sites, complexes I and III, which should be considered. In complex III, the essential parameter that regulates ROS generation is the dwell-time of the ubisemiquinone radical (QH• ) in the Q0 site of the enzyme (Cape et al., 2007). It is apparent that uncoupling of mitochondria has a different effect on this parameter, depending on the prevailing conditions (Figure 1) (Boveris et al., 1972; Boveris and Chance, 1973; Brookes et al., 2004). When mitochondria are in state 4 respiration (i.e. not phosphorylating ADP to ATP), ψm is high and the electron-driven H+ pumps (complexes) of the respiratory chain are feedback inhibited by a large H+ gradient. This gradient increases the lifetime of QH• by preventing the species from donating a proton to the cytosolic face of the membrane. Thus, a decrease in the H+ gradient (e.g. due to uncoupling) serves to dissipate such feedback inhibition, thereby allowing QH• to donate its H+ to the cytosolic face, and simultaneously its electrons to the b-heme system, and therefore decreases the ability of QH• to donate electrons to make O•− 2 . This concept is illustrated in panels 2 and 3 of Figure 1. Conversely, if the enzyme is inhibited, for example by antimycin A, the QH• dwell-time is enhanced because now QH• cannot pass electrons onto the bhemes of the enzyme. Thus, control over the steady-state concentration of QH• now lies not in QH• within the consuming reaction (which is blocked), but in the

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FIGURE 1 Differential effects of mitochondrial uncoupling on ROS generation by the Q cycle, dependent on inhibition status. Panel 1 shows the intact Q cycle at complex III of the respiratory chain. “Q” represents ubiquinol (oxidized), “QH2 ” ubiquinone (reduced), and “QH• ” the ubisemiquinone radical. The positions of cytochromes c, c1 , bH and bL are indicated. In the subsequent panels, magnitudes of fluxes are indicated by thickness of arrows. Panel 2 shows the generation of O•− 2 from the Q0 site, and the feed-back inhibition of this process by a large H+ gradient. When uncoupler is added (panel 3) this gradient is dissipated and O•− 2 generation accelerates. This is because control lies within the UQ• generating reaction. In contrast, when the enzyme is inhibited (e.g. by antimycin A), as shown in panel 4, control lies within the UQ• consuming reaction, and therefore anything that accelerates this reaction (such as uncoupling, panel 5) serves to accelerate O•− 2 generation. See text for full explanation.

QH• producing reaction. Since the reaction producing QH• itself requires the donation of a H+ to the cytosolic face (from QH2 ), then dissipating the H+ gradient by uncoupling allows this reaction to proceed more freely, and thus uncoupling enhances the generation of ROS. This is illustrated in panels 4 and 5 of Figure 1.

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Thus, uncoupling appears to both increase and decrease the generation of ROS from complex III, depending on whether the enzyme is inhibited or not. The exact mechanism of ROS generation by complex I is not completely understood, with debate surrounding the relative importance of the upstream FMN site vs. the downstream Q binding site (see Brookes et al., 2004 for review). Nevertheless, it is also apparent that the generation of ROS by complex I is highly dependent on the trans-membrane pH (Lambert and Brand, 2004b; Starkov and Fiskum, 2003). Together, the observations on complexes I and III have led to a consensus that dissipation of ψ (e.g. by uncoupling) leads to a decrease in ROS generation (reviewed in Brookes, 2005). Given the agreed role of excessive ROS generation in a number of disease pathologies, it has been consistently shown that a number of mild uncoupling strategies are protective in disease models. For example in heart tissue, mitochondrial uncouplers (e.g. FCCP, DNP) and overexperession of uncoupling proteins, are both protective against oxidative stress conditions such as ischemia-reperfusion injury (Brennan et al., 2006b, 2006a; Hoerter et al., 2004; McLeod et al., 2005; Minners et al., 2001; Nadtochiy et al., 2006; Teshima et al., 2003). However, this simple idea may not be universally applicable, and appears to be tissue-specific; recent reports suggest no benefit from uncoupling in models of neuronal pathology (Johnson-Cadwell et al., 2007; Tretter and Adam-Vizi, 2007). The differential make-up of mitochondria at the protein level between different tissues, which has recently been elucidated (Johnson et al., 2007a, 2007b) may provide some clues as to the specific proteins that regulate ROS in these different tissues.

B. Mitochondrial NO Synthase A prolonged and heated debate has surrounded the existence of a mitochondrial isoform of nitric oxide synthase (termed “mtNOS”) since its discovery over a decade ago (Bates et al., 1995, 1996; Kobzik et al., 1995). A full discussion of the controversy surrounding this enzyme is beyond the current chapter, but it may suffice to mention the following key facts: • There are numerous papers from prominent labs, all calling into question the key findings in the field of mtNOS (Brookes, 2004; French et al., 2001; Lacza et al., 2006b, 2003, 2004, 2006a; Tay et al., 2004). In these papers, much of methodology used to measure mtNOS has been shown to be fraught with artifacts. Antibodies against NOS recognize multitude off-target proteins. The urea cycle in mitochondria has significant enzymatic and inhibitor pharmacological overlap with NOS. The amounts of NO• claimed to be produced by mtNOS are incredulously high (>1 µM). No gene for mtNOS exists. None of the known NOS genes contain a mitochondrial target sequence. The kM for O2 of mtNOS precludes NO• generation at physiological O2 tension.

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• Even within the relatively small field of mtNOS (4 competing laboratories), there is considerable disagreement on the identity of the enzyme, which has been variously claimed to be an isoform of eNOS, iNOS, nNOS, or an unrelated protein (Bates et al., 1995, 1996; Ghafourifar, 2002; Ghafourifar and Richter, 1997; Giulivi et al., 1998; Giulivi, 2003; Kanai et al., 2001; Kobzik et al., 1995; Lopez-Figueroa et al., 2000; Riobo et al., 2002; Tatoyan and Giulivi, 1998). • A plant NOS in Arabidopsis thaliana (termed atNOS) was shown to contain a mitochondrial target sequence (Guo et al., 2003; Guo and Crawford, 2005), and a mammalian ortholog of this protein was lauded as a candidate for mtNOS (Zemojtel et al., 2006b). In-fact, atNOS is not a NOS, but a GTPase, and work on this enzyme has been retracted (Zemojtel et al., 2006a) (see also erratum to Ref. Zemojtel et al., 2006b). Such events in the plant literature are rarely publicized within the animal research field, and publications on mtNOS since this event have conveniently forgotten to mentioned these retractions. The proponents of mtNOS have routinely left the above negative papers uncited in their published works. With NIH funding percentages approaching single digits, it is essential that a balanced viewpoint on mtNOS is understood by the field at large. This has not been the case to-date.

C. Mitochondrial ROS & O2 Tension The production of mitochondrial reactive oxygen species (ROS) is oxygendependent, i.e. it requires an electron donor plus molecular oxygen (O2 ) as the electron acceptor. In the case of the mitochondrial electron transport chain (ETC), the electron donors are present as the FMN and Q binding sites of complex I, and the formation of the ubisemiquinone radical during the Q-cycle at complex III (see Section A above). The resulting product of the interaction of these sites with O2 is O•− 2 , which is dismuted to membrane-permeable hydrogen peroxide (H2 O2 ) by Mn-SOD in the matrix (Lambert and Brand, 2004a; Miwa and Brand, 2005; Muller et al., 2004; Nohl, 1990; Nohl et al., 2005). As expected, when O2 levels increase (hyperoxia), the availability of the electron acceptor increases and the result is an increase in ROS production (Chandel et al., 2000). Decreased levels of ROS generation would be expected to occur under conditions where the electron acceptor is limiting (hypoxia) (Hoffman et al., 2007). In addition to O2 sensitivity, the production of ROS is equally sensitive to oxidation state of the metal centers of the electron transport chain. This can be seen in studies examining the generation of ROS by isolated mitochondria under different respiration states: In state 3 respiration (ADP/ATP turnover) ψm is utilized to synthesize ATP, so proton pumping is increased and electrons flow

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freely through the ETC (Chance and Williams, 1955). Under these conditions, the metal centers in the ETC are more oxidized (i.e. more willing to accept electrons) (Severinghaus, 1999), and as-such donation of electrons to make ROS is decreased (Hoffman et al., 2007; Starkov and Fiskum, 2003). Conversely in the absence of ADP (state 4), ψm is not utilized, and protons are pumped across the membrane at a rate corresponding to the leak of protons back into the matrix (Brookes, 2005; Chance and Williams, 1955). This rate of proton pumping is much slower than in state 3, and leads to decreased electron flux and a more reduced ETC which donates electrons to make ROS more readily. The same set of arguments might naively be applied to conditions of hypoxia; i.e. the lack of O2 would result in a reduced ETC (Hoffman et al., 2007; Wilson et al., 1977) which should favor ROS generation. However, given the lack of O2 to function as an electron acceptor, ROS generation cannot occur. Currently there is much debate regarding the generation of ROS under hypoxic conditions, and the mechanism by which this may occur. It has been shown that mitochondrial ROS, specifically H2 O2 , is essential for the stabilization of the hypoxia-inducible factor 1α (HIF-1α) under hypoxic conditions (Guzy et al., 2005, 2007; Schroedl et al., 2002). Stabilization of this transcription factor leads to the activation of several genes required for hypoxic survival (Semenza, 2000). The mechanism of ROS dependent HIF stabilization is thought to occur through ROS mediated inhibition of O2 dependant prolyl-hydroxylases which regulates ubiquitination and degradation of HIF-1α (Semenza, 2001). However, the underlying mechanism behind the hypoxic ROS burst from mitochondria has been proposed to be one of mitochondrial autonomy (Chandel et al., 2000) and should therefore occur in isolated mitochondria. This proposed mechanism is not only paradoxical, but direct experimental evidence shows that it does not occur in isolated mitochondria (Hoffman et al., 2007). Two possible scenarios may explain this paradox. The first possibility is that ROS generation does not increase under hypoxic conditions (Michelakis et al., 2004), and what is being detected and labeled as “ROS” in whole cell systems is artifactual due to nonspecific ROS detection reagents. Much of the work showing an increase in ROS under hypoxic conditions utilizes the cell permeable fluorescent dye dichlorofluoroscein-diacetate (DCF-DA) (Chandel et al., 2000; Duranteau et al., 1998; Wang et al., 2007). Although cell permeable, DCF is prone to artifacts in the presence of peroxidases (Bonini et al., 2006). Recent evidence using a redox sensitive FRET probe suggests that under hypoxic conditions mitochondrial ROS does increase in cellular systems (Guzy et al., 2005), and thus as detection methods continue to improve, evidence supporting this will no doubt become more concrete. The second possibility, as suggested by recent FRET data, is that the hypoxic ROS burst does occur, but the underlying mechanism involves additional signaling molecules present in cell systems, but absent in isolated mitochondria. An example of such a signaling molecule is the reversible cytochrome c oxidase (complex

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FIGURE 2 Differential response of ROS generation to hypoxia in isolated mitochondria vs. mitochondria residing within cells. As discussed in the text, mitochondria in isolation produce less ROS under hypoxic conditions, but those inside cells produce more ROS in response to the same stimulus. It is hypothesized that the presence of a number of important cell signaling pathways (namely Ca2+ , NO• and protein kinase signaling cascades) confers the positive hypoxia → ROS response upon mitochondria. Such signals are lost when mitochondria are isolated from cells.

IV) inhibitor nitric oxide (NO• ). As a competitive inhibitor of mitochondrial respiration, NO• causes a right-shift in kM of cytochrome c oxidase for O2 (Cooper et al., 2003) leading to reduction of the ETC at much higher O2 levels than expected. Such inhibition would be expected to enhance ROS generation (Brookes and Darley-Usmar, 2002). Other possible contributors to hypoxic signaling that deserve more attention are Ca2+ , and protein kinases. Overall, an emerging consensus is that mitochondria within cells respond very differently to O2 tension than those in isolation (Hoffman et al., 2007). This is conceptualized in Figure 2.

II. EFFECTS OF ROS AND RNS ON MITOCHONDRIAL RESPIRATION The field of ROS, RNS and mitochondrial function has exploded in recent years, and it is far beyond the scope of this article to go into the fine details of all the potential targets of free radicals in mitochondria. For this, the reviewer is directed to several reviews on the subject (Balaban et al., 2005; Brookes et al., 2002a; Gutierrez et al., 2006; Shiva et al., 2005). In this chapter, only a selected sub-set of mitochondrial targets and their sensitivity to selected ROS and RNS will be discussed. Some of these reactions are shown in Figure 3.

A. ROS & Respiration Most of the major enzymes of the TCA cycle and the respiratory chain have at some time been demonstrated as targets of ROS-mediated damage and/or inactivation. The molecular mechanisms for these effects are often not clear. Briefly, the major targets in order of susceptibility are: aconitase > complex

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FIGURE 3 Integrated scheme of complex interactions between ROS, RNS and mitochondria. Although the focus of this review is mitochondrial membrane phenomena, several non-membrane targets of reactive species are also shown, for completeness. This figure is referred to in several sections of this chapter, so for full details and abbreviations, please see appropriate sections of the text.

I > α-KGDH > complex IV, complex II, complex III (Bolanos et al., 1995; Bulteau et al., 2003, 2006; Heales et al., 1997; Lesnefsky et al., 2001; Sadek et al., 2002). The mechanisms of ROS action include oxidation and damage to thiols, disruption of Fe–S centers, of which there are several dozen in mitochondria, and the oxidation of cardiolipin (see Section III). Interestingly, the oxidative modification and inhibition of key metabolic proteins in mitochondria has been proposed to constitute a type of feed-back loop, as depicted in Figure 4 (Armstrong et al., 2004). Briefly, the driving force for the generation of ROS is the entry of electrons into the ETC from reducing equivalents such as NADH and FADH2 . If the enzymes that generate these reducing equivalents are themselves susceptible to inhibition by oxidative stress (such as would be seen with excessive intra-mitochondrial generation of ROS), then such inhibition would serve as a safety-valve, to ensure that reducing equivalents do no continue to be fed into an ETC that is already making too many ROS. This hypothesis has been discussed extensively by Armstrong and colleagues (Armstrong et al., 2004).

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FIGURE 4 TCA-cycle feedback inhibition loop, regulating ROS generation by the respiratory chain. As proposed by Armstrong and colleagues, the oxidative inactivation of several proteins in the TCA cycle (notably α-KDGH and aconitase), will inhibit NADH generation, thereby limiting electron entry into the chain and thus slowing down ROS generation.

B. RNS & Respiration Respiratory enzymes are regulated by and generate a variety of reactive nitrogen species (RNS). Some of these species include nitric oxide (NO• ), Snitrosothiols (RSNOs), peroxynitrite (ONOO− ) and nitroxyl (HNO). The regulation of respiration by RNS gives rise to a number of interesting physiological effects. One of the most intriguing ideas introduced by Lancaster and colleagues suggests that NO• diffusing away from endothelial cells in blood vessels serves to inhibit respiration in cells proximal to the vasculature, and thereby allows O2 to diffuse further away from the vessel, thus in turn controlling hypoxia in cells distal to the blood supply (Thomas et al., 2001). The following section describes how RNS regulate respiration and how respiratory enzymes contribute to RNS generation. Complex IV of the mitochondrial respiratory chain is the best characterized target for RNS modification and regulation. The binding of NO• to the CuB /Heme a3 in complex IV reversibly inhibits mitochondrial respiration, and has been studied using spectrophotmetry, EPR and stop flow kinetic analysis. NO• binds to the heme-copper center via two different mechanisms, resulting in the release of NO• or NO− 2 . The specific nitrosylation reaction is dependent on whether NO• binds to a reduced or oxidized binuclear site (Sarti et al., 2000), which is controlled by a number of parameters including electron flux, O2 tension and ψm (Blackmore et al., 1991; Brookes et al., 2002b, 2003; Brunori et al., 2004; Giuffre et al., 2000). Complex IV thiols are also modified by RNS; nitrosation of thiols within the active site of complex IV has been reported after exposing cells to NO donors, or inducing inflammation as a result of tobacco smoke or polluted air exposure (Zhang et al., 2005). The mechanisms behind complex IV nitrosation are not as well defined as the nitrosylation reaction, and further investigation is required to define the relative physiological impact of this modification. Complex I is a site of electron entry into the respiratory chain, and its inhibition has been observed following exposure to a variety of RNS, including

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ONOO− and S-nitrosothiols. The downstream NO modifications that have been detected include nitrosated thiols and nitrated tyrosine residues. Complex I is inhibited by S-nitrosation, and the inhibition is reversed by light, and by reductants such as glutathione or dithiothreitol (Borutaite et al., 2000; Clementi et al., 1998; Zhang et al., 2005). Studies from our lab have identified the 75kDa subunit as a site of complex I S-nitrosation (Burwell et al., 2006). This is in agreement with other groups who have predicted this site to be S-nitrosated based on amino acid sequence (Chinta and Andersen, 2006). It is interesting that complex I is only inhibited ∼30–40% by S-nitrosation, and this may be important to reversibility of this modification (Nadtochiy et al., 2007). In addition to S-nitrosation, complex I tyrosine residues are targets of nitration. Nitration is not as reversible as nitrosation, and is seen under pathological conditions where there is a prolonged generation of NO• and O•− 2 (Tompkins et al., 2006). Complex II is a second site of electron entry in the respiratory chain, and is a component of the TCA cycle. There is evidence that complex II is reversibly inhibited by a nitroxyl (HNO) donor (Shiva et al., 2004a). This inhibition was found to be independent of S-nitrosation and was reversible by glutathione. Reversible inhibition of either or both complex I and complex II would serve to control electron entry into the mitochondrial respiratory chain, which may be important under pathological conditions such as ischemia-reperfusion injury. Like complex I, complex II is sensitive to ONOO− inhibition after prolonged NO• and • O•− 2 exposure (Cassina and Radi, 1996). There is also some evidence that NO may inhibit at complex III (Poderoso et al., 1999), although this has not been widely corroborated by other laboratories. When considering RNS-dependent control of respiration, it is also important to look at electron transport chain-independent modifications. For example, aconitase of the TCA cycle and pyruvate dehydrogenase are inhibited by ONOO− (Han et al., 2005; Martin et al., 2005). Inhibition of these enzymes would lead to a decrease in the production of NADH and FADH2 , and in turn lead to a decrease in respiration. RNS could also affect the transport of respiratory regulators, such as Ca2+ , into and out of mitochondria (see Section IV below). Previous studies have confirmed that Ca2+ transport is regulated by S-nitrosation in other areas of the cell (Sun et al., 2006; Yoshida et al., 2006), but the impact S-nitrosation has on mitochondrial Ca2+ transport proteins is difficult to study since a specific mitochondrial Ca2+ transporter awaits identification. Besides being modified by RNS, the respiratory chain also generates RNS. For example, ONOO− formation within mitochondria would be favorable, since NO• preferentially accumulates in mitochondria based on its hydrophobicity (partition coefficient ∼8) and O•− 2 would be produced from sites within the mitochondrion (Packer et al., 1996). It has also been observed complex IV serves a NO− 2 reductase under hypoxic conditions, to produce NO• (Castello et al., 2006). This nitrite reductase capability of complex IV may have evolved to control respiration. Some

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of the discussed effects of RNS and ROS on mitochondrial function are depicted in Figure 3.

III. MITOCHONDRIAL MEMBRANE LIPIDS The more common aspects of lipid oxidation and membrane biology are dealt with adeptly in several other chapters in this volume, and thus the following section will attempt to discuss only those aspects of membrane lipidology relevant to mitochondria.

A. ROS & Membrane Lipids When considering mitochondrial membrane lipids, two key parameters need to be considered: the head group and the fatty-acid side chains. Mitochondrial membrane lipids are approximately 90% phospholipid, and mitochondria possess a unique phospholipid called cardiolipin (CL), which is essentially a double phospholipid, i.e. 2 head groups covalently linked, with 4 acyl chains (Hoch, 1992). The biochemistry of CL is beyond the scope of this review, but it should be mentioned that the function of the respiratory chain complexes and mitochondrial carrier proteins is utterly dependent on CL (Kadenbach et al., 1982; Kuan and Saier Jr., 1993; Schlame et al., 2000). All of the respiratory complexes bind CL, and a role for CL is also proposed in the assembly of “super-complexes”, or the so-called “respirasome” (Schagger, 2002). Loss of CL alone is sufficient to account for respiratory chain dysfunction in a number of pathologies. Most notable of these is cardiac ischemia-reperfusion injury, which leads to a severe depletion of CL that is somewhat responsible for the bioenergetic crisis associated with this condition (Paradies et al., 2001, 2004, 1997, 1998; Petrosillo et al., 2003). Why is CL so susceptible to oxidative damage? Primarily, it is due to the very high complement (around 90%) of unsaturated fatty acids in its acyl chains (Hoch, 1992). Mitochondria are enriched in unsaturated lipids such as oleic, linoleic, arachidonic, and docosahexaenoic acids. This complement of fatty acids correlates highly with metabolic rate (i.e. faster metabolism equals more polyunsaturated membranes), and this parameter is also linked to the proton leak of the inner membrane (Brookes et al., 1997, 1998a; Brookes, 2005; Porter et al., 1996; Porter and Brand, 1993, 1995) (Figure 5). Thus, it is proposed that mitochondrial membrane fatty acid composition is a key determinant of basal metabolic rate. However, such a relationship between these parameters also has a downside, since it has been proposed that higher levels of lipid polyunsaturation in mitochondrial membranes may be a link between metabolic rate and aging; i.e.,

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FIGURE 5 Relationships between standard metabolic rate, mitochondrial H+ leak, and mitochondrial membrane fatty acid polyunsaturation levels. Data were collected by analyzing the H+ leak and fatty acid composition of liver mitochondria from a variety of organisms of varying metabolic rate, as pictured below the graphs. Organisms included: Rattus norvegicus, Mus musculus, Xenopus laevis, Pogona vitticeps, Trachhydosaurus rugosa, Crocodylus johnstoni, Onchoryncus mykiss, Oryctolagus cuniculus, Columba livia, Sus domesticus, Equus caballus, Ovis aries, Bufo marinus, Luchesa dugessi, obese Zucker rats, hyperthyroid and hypothyroid rats. Unsaturation index refers to the # of double bonds per 100 fatty acid molecules. Data are means of at least 3 independent measurements for each organism, and are collated from numerous studies (reviewed in Porter et al., 1996; Porter and Brand, 1995). * Note that axes are log–log, and thus correlation coefficients (r 2 ) between parameters span data across several orders of magnitude.

smaller animals live shorter lives because their membranes are more polyunsaturated and therefore more susceptible to oxidative damage (Pamplona et al., 1998, 1996; Rojas et al., 1993). The complex interplay between ROS generation, H+ leak, aging, and lipid polyunsaturation, at the level of the mitochondrion, has been extensively explored in a recent review article (Brookes, 2005).

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The oxidation of cardiolipin has also recently taken on significance due to discoveries regarding its role in apoptosis. It has been known for some time that the release of cytochrome c from mitochondria during apoptosis proceeds via a two-step pathway, i.e. first the protein must be released from the outer surface of the inner membrane before being released across the outer membrane (Ott et al., 2007). The regulation of the first stage was poorly understood until recent work from Kagan’s laboratory, showing that cytochrome c itself can adopt a peroxidase activity, and acts to oxidize cardiolipin’s side chains during apoptosis. This side-chain oxidation then causes a structural change in the head group of CL, decreasing its affinity for cytochrome c and permitting the latter’s release from the inner membrane surface (Basova et al., 2007; Belikova et al., 2007; Kapralov et al., 2007). The precise molecular mechanism by which cytochrome c adopts a peroxidase activity (i.e. what is the trigger for dys-coordination of the heme), is not yet known. Mitochondria have not (yet) been shown to contain cyclooxygenase and lipoxygenase enzymes, and thus the primary source of oxidized lipids in the mitochondrion is non-enzymatic lipid oxidation. Mitochondria are known to contain multiple lipid oxidation reaction products, and in addition it has been shown that addition of such molecules (e.g. 15d-PGJ2 ) to intact cells leads to their accumulation in mitochondria (Landar et al., 2006b). Thus, the mitochondrion has been proposed as a critical mediator for the signaling effects of oxidized lipid products (Shiva et al., 2004b). Some of the effects of lipid oxidation products (reactive lipid species, RLS) on mitochondria are consistent with their effects elsewhere in the cell, i.e. the posttranslational modification of nucleophilic protein amino-acid residues (Cys, His, Lys) by electrophiles such as 4-HNE, 15dPGJ2 . The so-called “electrophile reactive proteome” of the cell is in the process of being identified, and early reports suggest that a substantial proportion of such is mitochondrial (Bailey et al., 2005; Ceaser et al., 2004; Landar et al., 2006a). Key enzymes of the TCA cycle (αKGDH) and the respiratory chain (complexes I & II) have been identified as being inhibited by RLS (Lashin et al., 2006; Martinez et al., 2005), and the role of such inhibition in a variety of pathologic conditions is currently under exploration. The unique pH of the mitochondrial matrix may also render mitochondrial proteins particularly susceptible to electrophilic attack by virtue of the pKa of reactive protein residues. Interestingly, reactive lipids such as HNE may also mediate protective effects via signaling at mitochondria (see below).

B. RNS & Membrane Lipids While far less is known about the reactions of RNS with mitochondrial membrane lipids, it has been shown that ONOO− leads to extensive lipid oxidation, and with mitochondria being proposed as a likely source of ONOO− (Packer et

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al., 1996) this may be a major pathway for mitochondrial lipid damage in pathologic states (Gadelha et al., 1997). We showed several years ago that ONOO− induces a H+ leak (uncoupling) in brain mitochondria that is mediated via induction of secondary lipid oxidation (Brookes et al., 1998b). Considering the interactions of RNS with mitochondrial membrane lipids, it is essential to consider the hydrophobicity of NO• and the impact this has on the biochemistry of the free radical at the organelle level. It has been shown that the reaction between NO• and O2 , which serves as a sink for NO• in cells, is enhanced approximately 300-fold in the presence of biological membranes, vs. in the aqueous phase (Liu et al., 1998a). The mitochondrion is one of the primary membraneous organelles within the cell, and therefore this reaction is a potent regulator of NO• bio-availability at the organelle level. In-fact, we showed that at least part of the competition between NO• and O2 at the level of cytochrome c oxidase can be accounted for by a direct reaction between the two molecules within the hydrophobic membrane compartment of mitochondria (Shiva et al., 2001). A final recent development in the field of NO and membranes, is the discovery of a novel series of NO-derived lipid products, termed the nitro-alkenes (Kalyanaraman, 2004; O’Donnell et al., 1999). The mechanisms by which these species are generated is currently not clear, but they possess a wide range of anti-inflammatory and other signaling roles (Baker et al., 2007, 2004, 2005; Lim et al., 2002; Schopfer et al., 2005b, Schopfer et al., 2005a). By virtue of their electrophilic character, some of these signaling effects are mediated via post-translational protein modifications (Batthyany et al., 2006). Recent unpublished work from this laboratory has shown that nitro-alkenes are generated endogenously inside mitochondrial membranes, in a variety of patho-physiological conditions. The unique polyunsaturated lipid complement of the mitochondrion, coupled with a unique biochemical environment (e.g. pH, local ROS/RNS generation), suggest that mitochondria may be a very important source of nitro-alkenes within the cell.

IV. ROS, RNS & MITOCHONDRIAL ION TRANSPORT A. Overview When considering mitochondrial ion transporters, the phraseology of Winston Churchill in 1939 comes to mind. . . “a riddle, wrapped in a mystery, inside an enigma”. Mitochondria are known to transport a multitude of ions across their membranes, but the molecular identity of the ion channels responsible for these transport functions, is unknown (for an extensive review, see O’Rourke, 2007). Apart, of course, from the H+ pumps of the respiratory chain, the major ion fluxes in mitochondria are often characterized by their pharmacology, with the most

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widely characterized being the ruthenium-red sensitive Ca2+ uniporter (Gunter et al., 2004), the diazoxide-sensitive K+ ATP channel (Inoue et al., 1991), and the mitochondrial ryanodine receptor (Beutner et al., 2001). In addition there are several channels and transporters which are known gene products, but whose precise mechanisms of action is not yet clear (Kuan and Saier Jr., 1993), or which appear to adopt different functional properties depending on the stress status of the mitochondrion. This includes the permeability transition (PT) pore, the adenine nucleotide translocase (ANT), and the mitochondrial uncoupling proteins (UCPs). Each of these channels will now be discussed in detail: B. Ca2+ Uniporter The mitochondrial Ca2+ uniporter (MCU) is a high-capacity, highly specific channel of unknown molecular identity (Brookes et al., 2004; Gunter et al., 2004; Kirichok et al., 2004). This channel has been mostly studied via its sensitivity to the poorly-defined mixture of compounds ruthenium red (RuRed), of which the active component has been identified as a UV absorbing species termed “Ru360”, with an IC50 of ∼5 nM (Zazueta et al., 1999). There are a variety of effects of ROS and RNS on mitochondrial Ca2+ transport. Ca2+ uptake is driven in an electrophoretic manner by the membrane potential (ψ). Since both ROS and RNS affect the machinery of the electron transport chain, which generates the ψ, then the most obvious effect of RNS and ROS on Ca2+ uptake is to limit the driving force (e.g. we showed that NO• inhibition of complex IV inhibits mitochondrial Ca2+ uptake (Brookes et al., 2000)). Since the MCU is not yet identified at the molecular level, the role of RNS and ROS in regulating the function of this (presumed) protein by post-translational modification, is unknown. Interestingly, the MCU (or rather, ruthenium red sensitive mitochondrial Ca2+ uptake), has been identified as a potential target for p38 MAP kinase, which is induced under conditions of oxidative stress (Montero et al., 2002). Such a mechanism may act to link the redox status of the cell with mitochondrial Ca2+ homeostasis. On the flip-side regarding the effects of ROS/RNS on mitochondrial Ca2+ , is the equally interesting effect of Ca2+ on mitochondrial ROS generation (reviewed in Brookes et al., 2004). It has been reported that an increase in the matrix levels of Ca2+ can lead to an increase in ROS generation rates from the respiratory chain, but the mechanism underlying this effect is not clearly defined. Several proposed mechanisms have been proposed with the most favored current schemes including the following: (i) Ca2+ influx activates the TCA cycle dehydrogenase enzymes, thus increasing electron flux through the ETC and thus increasing ROS by simple mass action; (ii) Ca2+ activates mtNOS, although see earlier comments regarding this controversial entity; (iii) Ca2+ stimulates PT pore opening, leading to loss of cytochrome c, thereby inhibiting the ETC and leading to increased ROS;

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(iv) Related to this, PT pore opening leads to release of matrix GSH that scavenges ROS, so ROS release increases; (v) Ca2+ induces structural changes or possibly phosphorylation events in key ETC proteins, leading to ROS generation. All of these above mechanisms must also be reconciled with the simple fact that electrophoretic Ca2+ uptake into mitochondria consumes ψ, and thus will itself decrease ROS generation. Thus, it is often reported that Ca2+ decreases mitochondrial ROS on a short time scale (Brookes et al., 2004). Overall, there remains much to learn regarding the molecular events that link mitochondrial Ca2+ fluxes and ROS. The identification of proteins involved in these processes, most notably the MCU, would greatly advance the field. C. Ryanodine Receptor While the ryanodine receptor (RyR) has traditionally been thought of as a channel of the sarcoplasmic reticulum in excitable tissues such as heart, brain and skeletal muscle, a growing body of evidence suggests that the mitochondrial inner membrane also contains an RyR, termed “mRyR” (Beutner et al., 2001). The Ca2+ dose response of the mRyR is significantly left-shifted relative to the MCU (i.e. it responds to lower amounts of Ca2+ ), and this has led to the suggestion that the MCU and mRyR may act in cohort, to cover the entire range of physiologic Ca2+ concentrations. The rapid response of the mRyR is also proposed to be responsible for the rapid changes in [Ca2+ ]m , in response to changes in cytosolic [Ca2+ ], that have been documented (Beutner et al., 2005). Such changes may be important in processes such as excitation-metabolism coupling. Whether the mRyR actually constitutes the molecular identity of the so-called rapid uptake (RaM) mode of Ca2+ entry into mitochondria, remains to be determined (Gunter et al., 2004). Like other Ca2+ channels, the response of the SR RyR to RNS is well documented, and it is known that the channel is affected by changes in cellular redox status (Favero et al., 1995; Lipton et al., 1998; Sun et al., 2006; Yoshida et al., 2006; Zable et al., 1997). Unfortunately, the amounts of RyR found in mitochondria are vanishingly low, and thus-far all attempts to purify the protein from mitochondria have been unsuccessful. Thus, without the molecule itself to work with, it is difficult to know if any of the ROS- or RNS-mediated post-translational modifications that have been shown on the SR RyR will also be found on mRyR. D. K+ ATP Channel + The mitochondrial ATP-sensitive K+ channel (mK+ ATP ) is a K channel located in the inner mitochondrial membrane (Bednarczyk et al., 2004; Inoue et al., 1991) and plays a critical role in the protection provided by ischemic precon-

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ditioning (IPC) and pharmacological preconditioning (Armstrong et al., 1995; Auchampach et al., 1992; Garlid et al., 1997; Gross and Auchampach, 1992; Hide and Thiemermann, 1996; Liu et al., 1998b; Mizumura et al., 1995; Schultz et al., 1997). The mechanism of protection mediated by the activation of mK+ ATP remains unclear (Costa et al., 2006) and like many mitochondrial channels, mK+ ATP lacks a specific molecular identity. The pharmacology of mK+ is similar to that ATP of the known sarcolemmal K+ ATP channel, suggesting similar structural components; however many of the pharmacological agents also have nonspecific effects on mitochondria. For example, the commonly used mK+ ATP agonist diazoxide also inhibits complex II (Schafer et al., 1969). Despite extensive investigation, the molecular identity and in-situ regulators of this channel (apart obviously, from ATP) remain elusive. Active research focuses on the signaling pathways involving the channel. One such pathway involves ROS, which are necessary for IPC-mediated cardioprotection (Baines et al., 1997; Tritto et al., 1997; Vanden Hoek et al., 1998). While the mK+ ATP is involved in IPC (Armstrong et al., 1995; Gross and Auchampach, 1992; Mizumura et al., 1995), the influence of ROS on mK+ ATP is disputed. For instance, the activation of mK+ has been demonstrated to both inhibit ROS release in ATP isolated mitochondria (Facundo et al., 2007; Ferranti et al., 2003) and to increase ROS in cells (Forbes et al., 2001; Krenz et al., 2002). This discrepancy may be attributable to different model systems but a decrease in ROS coincides with the uncoupling effect of mK+ ATP activation, as well as the ability of mitochondrial uncouplers to decrease ROS (Starkov and Fiskum, 2003). Upstream of the channel, ROS is known to increase mK+ ATP activity through an unknown mechanism (Facundo et al., 2007; Zhang et al., 2001). Current evidence describes the channel as a redox-sensor which controls mitochondrial ROS release physiologically and pathologically (Facundo et al., 2007). The mK+ ATP is central to IPC signaling cascades and is therefore modulated by various protective signaling molecules such as NO• (Ljubkovic et al., 2007; Sasaki et al., 2000; Schafer et al., 1969). Most evidence is based upon the administration of pharmacological agents and the activation of the channel is determined by protection from an ischemic insult. The role of NO• in activating the channel was demonstrated with the addition of NO• donors which provided protection from an ischemic insult in a manner that was sensitive to mK+ ATP antagonists (Sasaki et al., 2000). Furthermore, the addition of a NOS inhibitor (Ockaili et al., 1999) or NO• scavenger (Sasaki et al., 2000) abolished • the mK+ ATP protective effects. Despite the direct interaction of NO with the + channel, the mKATP is thought to be a downstream target for PKG, in classic cGMP/NO• signaling (Costa et al., 2005; Cuong et al., 2006; Sato et al., 1998; Xu et al., 2004). In isolated mitochondria, exogenous PKG and cGMP resulted in the opening of the mK+ ATP , and the activation of the channel is blocked by inhibitors of the kinases and by mK+ ATP antagonists. The mechanism of PKG

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signaling remains unclear since the signal needs to transmit through mitochondrial membranes to reach the channel in the inner membrane. This mechanistic dilemma has been suggested to be overcome by PKCε which can activate mK+ ATP in cardiomyocytes (Sato et al., 1998). Despite advances in the mK+ ATP field, the lack of a molecular identity hampers the design of specific pharmacological tools. The limited knowledge of the mK+ ATP components impedes the establishment of the channel’s pathophysiological roles.

E. Permeability Transition (PT) Pore According to the classical definition, the PT pore is a non-selective high conductance channel of the inner and outer mitochondrial membranes, which assembles from existing proteins including but not limited to: ANT, VDAC, cyclophilin D and creatine kinase (Crompton, 1999) (Figure 3). While much of the phenomenology of the pore has been obtained via the use of inhibitors such as cyclosporin A (CsA) (Griffiths and Halestrap, 1993), the recent availability of knockout animals has forced a re-think of the composition of the pore. From a contemporary viewpoint, it is now widely accepted that cyclophilin D regulates PT pore formation (Baines et al., 2005; Nakagawa et al., 2005), however the involvement of ANT and VDAC is still a matter of debate since opening of PT pore has been demonstrated in ANT−/− (Kokoszka et al., 2004) or VDAC−/− (Baines et al., 2007; Krauskopf et al., 2006) mitochondria. Presumably, these proteins could be replaced by other mitochondrial membrane transporters, of the mitochondrial carrier family, suggesting that a degree of redundancy exists in the formation of an active PT pore complex. Opening of the PT pore establishes a non-selective mitochondrial permeability to solutes of less than 1.5 kDa. The consequences of mitochondrial permeabilization include mitochondrial swelling, rapid loss of ψm due to massive proton leak, followed by depression of ATP production (for review see Halestrap, 2006). In addition, the PT pore facilitates release of 12kDa cytochrome c and several other proteins from mitochondria, including AIF, endonuclease-G, Smac/Diablo and Omi/Htr2A. Release of pro-apoptotic proteins initiates caspase activation leading to proteolytic activation of crucial cellular targets in the apoptosis/necrosis cascade, finally resulting in cell death (for review see Gustafsson and Gottlieb, 2007). The PT pore is exquisitely redox sensitive, with mitochondrial ROS serving to activate the pore, and reductants such as glutathione and NADH serving to inhibit it (for review see Brookes et al., 2004). In addition, the PT pore offers a bi-phasic response so NO• , such that low doses of NO are protective (consistent with the anti-apoptotic role of such), whereas high doses of NO associated with ONOO− generation are detrimental, and promote pore formation (Brookes et al., 2000; Brookes and Darley-Usmar, 2004).

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F. ANT ANT is the best characterized and most abundant mitochondrial carrier protein (∼30 kDa) (Kuan and Saier Jr., 1993; Kunji, 2004; Palmieri, 2004), which exchanges non-mitochondrial ADP3− for intramatrix ATP4− in a ψ-dependent manner. Four different isoforms of ANT have been found. Based on RT-PCR, the amount of ANT1-4 mRNA in human tissues indicates that ANT1 is mostly expressed in heart, skeletal muscle and brain; ANT2 is expressed at higher levels in lung, testis and small intestine and less in kidney, liver and pancreas; ANT3 is expressed in very abundant amounts in lung and testis; ANT4 is expressed in liver at levels comparable to those of the other ANT genes (Dolce et al., 2005). Two highly specific inhibitors of ANT carboxyatractyloside (CAT) and bongkrekic acid (BKA) fix the carrier in cytosolic “c” and matrix “m” facing conformations respectively. ANT contains 3 critical cysteine residues (Cys57 , Cys160 and Cys257 based on the rat sequence) located on the matrix side, and redox modifications to these residues cause conformational and functional changes of ANT (Majima et al., 1993). Under physiological conditions Cys160 plays a significant role because it is a binding site for adenine nucleotides (Majima et al., 1993). Cys57 is a cyclophilin D binding site, thus it is a critical residue for CsA-sensitive PT pore formation. Under events of oxidative stress and Ca2+ overload, ANT may lose its original properties of nucleotide transporter and become a structural component of PT pore (see above). It has been demonstrated that oxidative stress induced by t-butylhydroperoxide (t-BuOOH) stimulates cyclophilin D binding to ANT, sensitizing mitochondria to Ca2+ -induced PT pore opening (Connern and Halestrap, 1994); moreover, ANT isolated from mitochondria treated with Cu2+ /t-BuOOH exhibited a progressive loss of lysine, cysteine, arginine, and valine residues (Giron-Calle and Schmid, 1996). Oxidative stress and thiol reagents may also stimulate PT opening by affecting nucleotide binding to ANT (Halestrap et al., 1997) or cross-linking Cys257 with Cys160 (McStay et al., 2002). Overall, the “c” conformation facilitates PT pore opening, while in contrast events that fix ANT in “m” conformation (BKA, ADP) inhibit the PT pore. The differential exposure of ANT thiols in these two different conformations may partially explain such effects.

G. UCPs Uncoupling proteins (UCPs) belong to a large family of anion carriers that are present in the inner mitochondrial membrane (Palmieri, 2004). Originally uncoupling protein (UCP1) was discovered as an inner mitochondrial membrane protein which regulated thermogenesis in brown adipose tissue by wasting the ψm in the form of H+ leak (Nicholls, 2001). In 1997 two genes for UCP2/3

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and later, in 1998 UCP4/5 were discovered. UCP2 is highly expressed in immune cells, kidney, heart, lung and pancreatic β-cells; UCP3 is expressed at higher levels in skeletal muscles; UCP4/5 were originally found in the brain. Surprisingly both plants (Douette and Sluse, 2006; Echtay, 2007; Nicholls, 2006; Palmieri, 2004) and ectotherms (zebrafish) (Stuart et al., 1999) contain UCPs, which suggests these proteins may play roles other than simple thermogenesis. Despite the fact that UCP2 and UCP3 share only 59% and 57% identity with original UCP1, and UCP4/5 has even less then 35% identity with UCP1, all these proteins were combined into one family because of their property to uncouple mitochondrial OXPHOS by increasing inner mitochondrial membrane H+ leak. UCPs can be activated by free fatty acids and proton conductance is inhibited by purine nucleotides. The physiological function of UCP2-5 is unlikely to be thermogenesis regulation because they are expressed in very low amounts, less then 1–2% compared to UCP1 in brown adipose (Harper et al., 2002; Pecqueur et al., 2001). Alternatively, several different functions for UCPs have been proposed including regulation of insulin secretion, fatty acid metabolism and oxidative stress (for reviews see (Echtay, 2007; Green et al., 2004; Negre-Salvayre et al., 1997)). The last function is the direct subject of this chapter. As discussed above (see Section A), a small degree of uncoupling can significantly decrease mitochondrial ROS generation (Votyakova and Reynolds, 2001) with no effect on ATP production. However, far more interesting is the discov+ ery that O•− 2 induces H leak via activation of UCPs (Echtay et al., 2002, 2003; Murphy et al., 2003) Despite the fact that direct activation of H+ leak by O•− 2 has been questioned (Cannon et al., 2006), it has been demonstrated that the electrophilic lipid oxidation product 4-hydroxy-2-nonenal (HNE) activates H+ leak via both UCPs and ANT (Echtay et al., 2003), and on the basis of these data suggested that electrophiles may trap UCP and ANT into a heterodimer. In addition, peroxynitrite (ONOO− ) can induce mitochondrial H+ leak via a mechanism involving lipid oxidation (Brookes et al., 1998b). It is unclear if this latter effect proceeds via UCPs or ANT, but interestingly, unpublished work from this laboratory has found that nitrated lipid products (nitro-alkenes, e.g. nitro-linoleate, see Section B) can activate mitochondrial H+ leak, and this occurs via direct covalent modification of thiols on UCPs and ANT.

V. COMPLEX INTERACTIONS & CONCLUDING REMARKS An example of the way in which the complex pathways detailed in the previous sections come together to form an integrated signaling pathway, is shown in Figure 6. In this scheme, ROS generated by the respiratory chain can initiate lipid oxidation, leading to the generation of reactive lipid species (e.g. 4-HNE).

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FIGURE 6 Auto-regulation of mitochondria ROS generation via cross-talk with UCPs. ROS generated by the mitochondrial electron transport chain (ETC, depicted schematically as complexes I–IV), leads to the generation of reactive lipid species (RLS) in the mitochondrial inner membrane. These RLS then covalently modify and activate H+ conductance channels in the membrane, namely UCPs and ANT. Activation of H+ leak then serves to decrease ROS generation. For full details see text.

These electrophiles can then covalently modify cysteine residues on key mitochondrial proteins (ANT & UCPs), activating a H+ leak channel activity. This H+ leak leads to mild uncoupling of the mitochondria, thereby decreasing the membrane potential (10–15 mV), which in turn decreases the rate of ROS generation. Thus, the stimulus for this process (ROS) acts to switch itself off again, in an auto-regulatory feedback loop (Brookes, 1998, 2005). Of course, the missing link in this loop is how it regenerates, since the post-translational modification of proteins by reactive lipids is currently only thought to be an irreversible process. Thus, the only way to switch off an electrophile-induced H+ leak via UCP is to destroy the UCP and make a new protein. This may in-fact be the case, as it has recently been demonstrated that UCPs have incredibly high turnover rates (Rousset et al., 2007). It is not difficult to imagine that similar relationships exist between ROS/RNS, and the other major ions transported by mitochondria, suggesting a highly orchestrated network of ion movements and free radicals. Overall, such interactions between mitochondrial membranes, including both their protein and lipid components, and ROS/RNS, are incredibly complex. Just the simple act of answering a question such as “why does Ca2+ uptake increase ROS” is impossible to answer at this point in time, and there is much to learn regarding the molecules involved, both proteinaceous and otherwise. The integration of such multi-component systems into useful working models will require massive computational power, in order to predict the effects of something so simple as NO on overall mitochondrial function. The fundamental involvement of mitochondria in both the life of the cell, and the processes of apoptosis, necrosis, and disease pathology, assures that the investigation of these processes at the molecular level will yield many new targets for pharmacologic targeting, to combat disease.

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CHAPTER 11 Oxidant Stress and Airway Epithelial Function1 Jenora T. Waterman and Kenneth B. Adler Department of Molecular Biomedical Sciences, North Carolina State University, College of Veterinary Medicine, 4700 Hillsborough Street, Raleigh, NC 27606, USA

I. Introduction II. Sources of Reactive Oxygen Species III. Antioxidant Defenses in Airway Epithelium A. Non-enzymatic Antioxidants B. Enzymatic Antioxidants IV. Oxidant-induced Airway Epithelial Responses A. NF-κB Activation, Cytokine Release and Oxidant Stress in Airway Epithelium B. Oxidative Stress and Airway Epithelial Mucus Hypersecretion V. Conclusions References

I. INTRODUCTION Radicals of oxygen such as superoxide anion (O•− 2 ) and hydroxyl radical (OH− ), and metabolites including hypochlorous acid, (HOCl) and hydrogen peroxide (H2 O2 ) are referred to as reactive oxygen species (ROS) because they readily react with biological molecules such as DNA, lipids and proteins. ROS are generated primarily by normal metabolic reactions in all cell types. In mammalian systems, ROS production by several inflammatory cells may be increased in chronic respiratory diseases such as asthma. Although substantial evidence exists characterizing ROS as chemicals capable of causing damage to cellular structures, they also play important roles as signaling molecules within cells. The doubleedged phagocytic cells of the immune system—neutrophils and macrophages— must generate ROS to kill certain types of bacteria and other microbes. However, 1 Supported by grant # R37 HL36982 from NIH.

Current Topics in Membranes, Volume 61 Copyright © 2008, Elsevier Inc. All rights reserved

1063-5823/08 $35.00 DOI: 10.1016/S1063-5823(08)00211-1

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neutrophil retention in airways along with overproduction of ROS and/or prolonged exposure of tissues to ROS are features common to several respiratory diseases, such as asthma, chronic bronchitis, and cystic fibrosis. This review will discuss the various sources of oxygen radicals and antioxidant defenses in the airways, and the role of the oxidant-sensitive transcription factor, NF-κB, and its downstream targets on airway epithelial function.

II. SOURCES OF REACTIVE OXYGEN SPECIES ROS are formed when electrons are added to molecular oxygen, which under normal conditions has two unpaired electrons distributed over different shells in its outer orbital. This configuration and its affinity for electrons make O2 ideal for aerobic energy production and consequently radical formation. When O2 accepts one electron O•− 2 is produced, adding a second electron will yield H2 O2 and a third produces OH− . These three molecules are termed ROS because they readily react with other molecules such as DNA, protein and lipids. When oxygen gains a fourth electron it is reduced to water and is therefore no longer reactive. ROS in airways may originate from endogenous and/or exogenous sources. Endogenous sources include respiratory bursts from activated inflammatory and immune cells, normal metabolic reactions with the electron transport chain of the mitochondria being the major source and resident airway epithelial cells themselves (Martin et al., 1997). As electrons are passed along the chain from protein to protein, electrons leak from the electron transport chain onto oxygen molecules to produce superoxide anion. Neutrophils and macrophages produce ROS to destroy engulfed bacterial or fungal pathogens. In neutrophils, engulfed bacteria are compartmentalized into phagosomes which fuse with ROS- and hydrolytic enzyme rich-lysosomes. The consumption of oxygen during the generation of ROS is termed the “respiratory burst.” The respiratory burst involves activation of the enzyme nicotinamide adenine dinucleotide phosphate (NADPH) oxidase, which produces large quantities of O•− 2 . Superoxide dismutates subsequently catalyze the conversion of O•− anions to H2 O2 . High levels of neutrophils 2 in bronchoalveolar lavage fluid of patients with chronic airway disease correlates with poor pulmonary function (Welsh et al., 1995; Stanescu et al., 1996; Senior and Shapiro, 1998). Analysis of sputum from children with cystic fibrosis showed there was a correlation between increased concentrations of neutrophils, elastase activity and interleukin-8 (IL-8) with matrix metalloprotease-9 (MMP9) and its inhibitor tissue inhibitor of metalloprotease-1 (TIMP-1) (Sagel et al., 2005). Exogenous sources of ROS include exposure to environmental air pollutants (Becker et al., 2005a, 2005b), ozone (Kierstein et al., 2006) and cigarette smoke (Macnee, 2005, 2007). Cigarette smoke, an extremely rich source of oxidants,

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elicits proinflammatory response in the airways via NF-kB activation and resultant cytokine production and release (Yang et al., 2006).

III. ANTIOXIDANT DEFENSES IN AIRWAY EPITHELIUM The airways are well-equipped for protection against exogenous ROS and oxidants. Studies on animal models have shown that oxidants can increase the level of mucins, a large family of heavily glycosylated proteins secreted into airway epithelial lining fluid (Adler and Li, 2001), which can trap and facilitate removal of microbes, dusts and other airway irritants. In addition to mucins, the epithelial lining fluid (ELF) contains high levels of glutathione (GSH) (Cantin et al., 1987), a tripeptide considered to be the primary antioxidant defense in the lung as it is constitutively active and inducible by oxidative stress. Normal airways are replete with antioxidants to defend against intracellular and extracellular oxidants. There are two classes of antioxidants in the lungs; non-enzymatic and enzymatic antioxidants. Non-enzymatic forms work to scavenge free radicals and prevent ROS formation, while enzymatic antioxidants catalyze reactions that ultimately reduce ROS to non-reactive oxygen-containing compounds.

A. Non-enzymatic Antioxidants Non-enzymatic antioxidants consist of low molecular weight compounds such as vitamin E (tocopherol), vitamin C (ascorbate), β-carotene and glutathione; and high molecular weigh antioxidants include albumin, lactoferrin and transferrin, which act by binding heavy metals, making them unavailable for radical generation (Burton and Ingold, 1989). In a murine ovalbumin model of allergic airway inflammation, oxidant stress was more prevalent in a vitamin E-restricted group compared to a vitamin E-supplemented group (Talati et al., 2006). Cells particularly affected were airway epithelial (Clara) cells and macrophages. Diets low in vitamins C and E, fruits, manganese and n − 3 fatty acids have been associated with decreased pulmonary function and increased symptoms of bronchitis and asthma (Patel et al., 2006; Burns et al., 2007). Glutathione (L-γ -glutamyl-L-cysteinylglycine) is probably the most important non-protein antioxidant for protection against ROS damage, and is present in virtually every living cell (Meister, 1988). This tripeptide, with its reactive sulfhydryl-containing cysteine center, serves as a target for radical attack (Kidd, 1997; Pastore et al., 2003). In cells, glutathione is present mainly in its reduced form which becomes oxidized to glutathione disulfide (GSSH) during oxidative stress. The reduced form, GSH, is generated in a redox cycle with glutathione reductase and NADPH. Under normal physiological conditions reduced and oxidized forms of glutathione (GSH:GSSH) occur within cells at concentrations

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between 1 and 10 mM, with GSH being the dominant form (Pastore et al., 2003). Glutathione levels are a critical determinant of tissue redox status and an increase in GSSH levels over GSH is indicative of oxidative stress (Pastore et al., 2003; Day, 2005). The lung has a remarkable ability to concentrate GSH in epithelial lining fluid (ELF). In fact, GSH levels are approximately 100-fold greater in ELF than in serum (Cantin et al., 1987). Glutathione levels are diminished in a number of respiratory disorders, such as cystic fibrosis (Roum et al., 1993), adult respiratory distress syndrome (ARDS) (Pacht et al., 1991) and human immunodeficiency virus (HIV) infection (Buhl et al., 1989), following lung transplantation (Baz et al., 1996), or due to environmental and occupational exposures including asbestos (Brown et al., 2000). GSH levels are increased in bronchoalveolar lavage fluid of asthmatics (Smith et al., 1993) and ELF of cigarette smokers (Cantin et al., 1987).

B. Enzymatic Antioxidants Major enzymatic antioxidants are superoxide dismutase (SOD), catalase (CAT) and glutathione peroxidase (GPO) (Burton and Ingold, 1989; Rahman et al., 2006). Superoxide dismutases are enzymes which catalyze dismutation of two •− O•− 2 molecules to H2 O2 , which is substantially less toxic than O2 , and O2 . Al•− though the conversion of O2 molecules to H2 O2 can occur under non-catalytic conditions, SOD accelerates this detoxification reaction as much as 10,000fold over the non-catalyzed reaction (Burton and Ingold, 1989; Rahman et al., 2006). There are three major mammalian SODs; copper-zinc SOD (CuZnSOD) which is located in the cytoplasm, manganese SOD (MnSOD) which is most abundant in the mitochondria, and extracellular SOD (ECSOD) (Kinnula and Crapo, 2003). Superoxide dismutases are present in essentially every cell of the body and have been shown to play a critical role in protection against oxidative stress. In airway epithelium of asthmatics, SOD activity, particularly CuZn SOD, is decreased (Smith et al., 1997) which is proportional to an increase in O•− 2 anion production (Jarjour et al., 1992). Mn SOD, considered one of the most essential antioxidant components of a cell, is induced by altered redox states, inflammatory cytokines, cigarette smoke and hyperoxemia (Rahman et al., 2006) and is critical for survival since Mn SOD-deficient mice die within 10– 21 days after birth from neurodegeneration and myocardial injury (Li et al., 1995; Lebovitz et al., 1996). Extracellular SOD is abundant in blood vessels, pulmonary fluids and airways and together with glutathione peroxidase serves as a first line of defense against inhaled oxidants (Rahman et al., 2006). Catalase is one of three major enzymes responsible for H2 O2 elimination. It completes the detoxification commenced by SOD by producing H2 O and O2 from H2 O2 (Burton and Ingold, 1989; Rhee et al., 2005). Tracheal epithelial cells isolated from catalase transgenic mice had lowered dichlorodihydrofluorescein

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(DCF) levels resulting from diminished baseline H2 O2 levels (Reynaert et al., 2007). Glutathione peroxidases are a family of antioxidant enzymes that degrade H2 O2 and lipid peroxides (Hou et al., 1996; Siest et al., 1997). When the airway is exposed to pollutants or irritants such as cigarette smoke or dust, an inflammatory response may ensue. The trachea may take the brunt of the offense, and it has been reported that depletion of antioxidant reserves in the trachea may reflect persistent oxidant and inflammatory processes within bronchoalveolar spaces (Deaton et al., 2006). In smokers with COPD, exogenous antioxidant defenses such as vitamins (C and E) have been insufficient in combating smoke-related lung injury (Kinnula, 2005). Exogenous antioxidants such as vitamins have had little effect in combating smoke-induced airway trauma, however, N-acetylcysteine, a glutathione related synthetic therapeutic, has shown promise (Comhair and Erzurum, 2002; Blesa et al., 2003; Kinnula, 2005).

IV. OXIDANT-INDUCED AIRWAY EPITHELIAL RESPONSES Mammalian airways are lined with morphologically distinct epithelial cells types which may be classified into three major categories: basal, ciliated and secretory (Knight and Holgate, 2003). These cells function as an assimilation unit providing a physical barrier between the internal lung milieu and the external environment. It has become clear that the airway epithelium performs many functions critical to maintaining homeostasis including metabolism and clearance of inhaled agents, attraction and activation of inflammatory cells and phagocytes in response to injury, regulation of lung fluid balance and airway smooth muscle function via secretion of numerous mediators (Holgate, 2000; Holgate et al., 2000; Knight, 2001; Knight and Holgate, 2003).

A. NF-κB Activation, Cytokine Release and Oxidant Stress in Airway Epithelium In the airways, systemic inflammatory responses such as sepsis, severe trauma, and burns, as well as many noxious and/or inflammatory stimuli including bacterial products and ozone, have been known to activate the transcription factor, NF-κB. In quiescent cells, NF-κB is sequestered to the cytosplasm in an inhibitory complex with inhibitor of kappa B (I κB). Following cellular stimulation, I κB molecules become phosphorylated at several serine residues in the amino terminus by IκB kinase 2 (IKK2) which targets them for ubiquitination and proteosome-degradation (Rothwarf and Karin, 1999; Karin and Ben-Nerian, 2000; Häcker and Karin, 2006). This process allows nuclear translocation of NFκB for transcriptional activation of a variety of pro-inflammatory genes including

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cytokines/chemokines, immunoreceptors, cell adhesion molecules, growth factors and their modulators (Pahl, 1999). Increasing evidence is mounting for the involvement of nuclear factor kappa B (NF-κB) in chronic respiratory diseases such as asthma and COPD. In a study by Pierrou and MacNee (2007) inflammatory signaling pathways involving NF-κB and AP-1 were overexpressed in smokers with COPD. These findings are consistent with mouse models for NFκB-mediated lung inflammation and injury (Cheng et al., 2007). Transgenic mice with inducible activation of NF-κB in airway epithelial cells elicited cytokine production, inflammatory cell recruitment (primarily neutrophils) and lung injury by day 3; and high mortality in as few as 7 days of constitutive NF-κB signaling (Cheng et al., 2007). Expression of a dominant negative inhibitor of NF-κB (IKK 2) prevented lung inflammation and injury associated with NF-κB in airway epithelium (Cheng et al., 2007). NF-kB, considered a pivotal transcription factor in chronic inflammatory diseases, is sensitive to oxidants and other stimuli (Schreck et al., 1992). For example, low levels of H2 O2 have been shown to induce NF-kB activation, which can be inhibited by antioxidants (Schreck et al., 1992). Several reports indicate a relationship between cytokines (Chung and Barnes, 1999), oxidant generation/stress (Bowler and Crapo, 2002a, 2002b; Crapo, 2003) and consequential mucin gene expression and secretion (Adler et al., 1990, 1994; Wright et al., 1996; Fischer et al., 1999; Krunkosky et al., 2003). Tumor necrosis factor-alpha (TNF-α) is a primary inflammatory mediator produced by macrophages and a variety of other cells types including mast cells (Pendl et al., 1997), cardiac monocytes (Röntgen et al., 2004), endothelial cells (Edelman et al., 2007), and bronchial epithelial cells (Carter et al., 1997). The role of TNF-α in airway inflammation has been well documented. Studies in our laboratory revealed that in primary cultures of guinea pig tracheal epithelial cells, TNF-α provoked mucus secretion in a time- and concentrationdependent manner (Fischer et al., 1999). TNF-α also stimulates expression of adhesion molecules such as intercellular adhesion molecule (ICAM)-1 and Vascular cell adhesion molecule (VCAM)-1 (Marui et al., 1993; Gosset et al., 1994; Krunkosky et al., 2003) in airway epithelium and pulmonary endothelium. It is worth noting that the promoter region of ICAM-1 contains NF-kB binding sites and its expression is oxidant sensitive (Marui et al., 1993). Once stimulated, TNFα can provoke secretion of secondary mediators of inflammation including IL-6 and GM-CSF (Cohn et al., 1997). Importantly, TNF-α has NF-kB binding sites in its regulatory region, suggesting a key role for NF-kB in mediating TNFα-mediated airway inflammation. Moreover, NF-kB activation may result from TNF-α secretion and can lead to ROS generation (Rahman et al., 1996). TNFα plays an important role in oxidant stress in that it can trigger expression of genes including MUC5AC, iNOS and ICAM-1 whose function can contribute to the pathogenesis of inflammatory diseases such as asthma (Wuyts et al., 2001; Fischer and Voynow, 2002; Krunkosky et al., 2003).

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IL-8, a potent chemoattractant, is a secondary inflammatory mediator produced in response to primary mediators of inflammation such as TNF-α (Martin et al., 1997) and other stimuli including oxidant stress (DeForge et al., 1993; Becker et al., 2005b) via a mechanism involving Toll-like receptor-2 (TLR-2) (Becker et al., 2005a). In vitro studies have shown that normal human bronchial epithelial (NHBE) cells and alveolar macrophages respond to particulate matter as found in ambient air (e.g., diesel particles) by generating proinflammatory mediators including IL-8 and COX-2 (Becker et al., 2005b). Similar reports have shown that hydrogen peroxide elicits IL-8 release in NHBE cells (Pelaia et al., 2004), immortalized human bronchial epithelial cells (BEAS-2B) (Oslund et al., 2004) and primary human tracheobronchial epithelial cells (Oslund et al., 2004).

B. Oxidative Stress and Airway Epithelial Mucus Hypersecretion Features common to asthmatic airways include hyper-responsiveness, antioxidant depletion (Caramori and Papi, 2004) persistent inflammation, mucus hypersecretion and epithelial damage. Oxidant stress can lead to mucus hypersecretion (Macnee, 2005, 2007) and decreased clearance of potential pathogens (Wright et al., 1994). Airway retention of neutrophils, possibly in response to cytokine stimuli from resident airway epithelial cells or in response to bacterial presence, is accompanied by increased levels of oxidants that prolong the inflammatory process and ultimately lead to cell and tissue damage. Mucus overproduction is common to airway disease states and is a major cause of airway obstruction in COPD (Macnee, 2005, 2007), asthma, cystic fibrosis and bronchiectasis and is associated with increased oxidant levels (Fischer and Voynow, 2002). MUC5AC, believed to be the major mucin contributing to chronic airway diseases, is closely related to mucus hypersecretion and goblet cell metaplasia (Voynow, 2002; Casalino-Matsuda et al., 2006) and its regulation is critical to the pathogenesis of cystic fibrosis and other chronic respiratory illnesses. Work by Basbaum and colleagues has dealt with understanding the relationship of smoke-induced ROS generation and mucin gene expression in the airway. One of the earliest effects of airway epithelial cell exposure to tobacco smoke is oxygen radical generation (Lemjabbar et al., 2003). DCF fluorescence studies indicated that smoke exposure raised levels of ROS in lung cells, which was inhibited by antioxidants including diphenyliodonium chloride, an inhibitor of NADPH oxidase, indicating an essential role for NADPH oxidase in the early response of airway cells to tobacco smoke (Lemjabbar et al., 2003). Further studies by Basbaum and colleagues showed that tobacco smoke can stimulate MUC5AC through cooperation of JNK and ERK via pathways involving ROS (Gensch et al., 2004). Results of in situ hybridization studies done on human autopsy bronchial tissues from smokers and non-smokers indicated that smoke stimulated MUC5AC expression in vivo in airways as well as in cell cultures (Gensch et al., 2004). In

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transcription activation studies, airway epithelial cells exposed to smoke extract had a 7-fold increase in luciferase activity, indicative of MUC5AC transcription induction. Deletion mutagenesis studies revealed a smoke-responsive element in the MUC5AC gene regulation region which contained four binding sites for three major transcription factors; 2 AP-1 sites, one RXR site, and one NF-kB site. Deletions in each of these transcription binding sites resulted in decreased MUC5AC gene expression, with the greatest effect arising from mutation of the AP-1 sites. It has been documented that neutrophil elastase (NE) mediates MUC5AC gene expression and stability via ROS-dependent pathways (Voynow et al., 1999; Fischer and Voynow, 2000, 2002). Results of nuclear run-off and mRNA stability assays revealed that NE does not initiate transcription of new MUC5AC mRNA, but instead NE causes an increase in MUC5AC transcript stability, enhancing message half-life in airway epithelial cells from 4.5 hr in resting cells to 14.75 hr (Voynow et al., 1999). There was an increase in DCF fluorescence in NE-treated A549 and NHBE cells which correlated with suppression of NEinduced MUC5AC mRNA expression when cells were preincubated with the antioxidant dimethylthiolurea (DMTU) (Fischer and Voynow, 2000). Other studies indicated that co-incubation of NE with antioxidants inhibited NE-mediated MUC5AC gene expression (Fischer and Voynow, 2002). Although there is clear evidence linking NE-induced oxidant injury and MUC5AC regulation, the mechanism and source of ROS remains unclear. Recent studies from Voynow and colleagues have shown that NE-induced MUC5AC mucin gene expression and oxidant stress in airway epithelial cells can be prevented in A549 and NHBE cells in a concentration-dependent manner by dicumarol, an inhibitor of NADPH:quinone oxidoreductase 1 (NQO1) (Zheng et al., 2007). This study presented evidence that NQO1 activity is essential for NE-induced MUC5AC mRNA expression and that NQO1 may be responsible for NE-induced ROS generation. Suppression of NQO1 gene and protein expression via small inhibitory RNAs (siRNA) significantly inhibits NE-mediated induction of MUC5AC gene expression. Zheng et al. (2007) investigated NE regulation of NQO1 expression and discovered that NE increased NQO1 protein levels by 45 ± 10% and 400 ± 79% in A549 and NHBE cells, respectively, which consequently increased NQO1 activity (and resultant oxidant stress) in these cells. Therefore NQO1 regulation by NE appears to be a necessary part of the oxidant-dependent mechanism governing MUC5AC regulation. A novel family of calcium-activated chloride channels (CaCC), present in human secretory organs, may play a mechanistic role in facilitating mucus secretion in airway epithelium. A member of the CaCC family, Gob-5, was found in goblet cells in a murine allergic asthma model (Nakanishi et al., 2001). Importantly, human ClCa1 (hClCa1) shares a high degree of sequence and structure homology with mouse Gob-5 (Gruber et al., 1998, 2000). Recent studies in our laboratory have shown a correlation between mucus secretion and hClCa1 upregulation in airway epithelial and intestinal cells (Raiford, unpublished results).

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Other researchers have reported an increase in hClCa1 mRNA in non-cancerous bronchial tissues from Chinese asthmatics that correlated with MUC5AC expression and mucus secretion (Wang et al., 2007). An intriguing finding by Reynaert and colleagues (2007) implicated a role for H2 O2 in regulating mucin secretion in conjunction with MUC5AC and ClCa3 expression in murine models. In that study CAT overexpression diminished baseline levels of H2 O2 and enhanced mucin secretion. This novel finding represents a direct role for the repression of mucin gene expression or mucus hypersecretion by H2 O2 . Clearly there is a necessary role for H2 O2 and other ROS as signaling molecules (Krunkosky et al., 1996) in normal respiratory function and mucus production.

V. CONCLUSIONS The respiratory epithelium, once thought to be a passive barrier between internal and external environments, plays an active role in protecting against ROS damage. Airway epithelial cells utilize a complex combination of non-enzymatic and enzymatic antioxidant defenses to combat against oxidative assaults. When the balance between oxidants and antioxidant systems is disrupted in favor of oxidants, oxidative stress is said to occur. The airway epithelium essentially is a target and effector tissue that responds to and generates various stimuli that contributed to airway inflammation. As a target, it responds to stimuli such as environmental pollutants and bacterial products through complex mechanisms including ROS production, mucus production and secretion, infiltration and activation of inflammatory cells, and alteration of ciliary dynamics. As an effector, the epithelium generates inflammatory mediators such as autocrine- and paracrine-functioning cytokines. The caveat is that while ROS have an essential role in signal transduction, they also can, when generated in excess, enact deleterious and injurious effects on airway epithelium. References Adler, K.B., Fischer, B.M., Wright, D.T., Cohn, L.A., Becker, S. (1994). Interactions between respiratory epithelial cells and cytokines: Relationships to lung inflammation. Ann. NY Acad. Sci. 725, 128–145. Adler, K.B., Holden-Stauffer, W.B., Repine, J.E. (1990). Oxygen radicals stimulate release of high molecular weight glycoconjugates by cell and organ cultures of rodent respiratory epithelium via an arachidonic acid-dependent mechanism. J. Clin. Invest. 85, 75–85. Adler, K.B., Li, Y. (2001). Airway epithelium and mucus: Intracellular signaling pathways for gene expression and secretion. Am. J. Respir. Cell Mol. Biol. 25, 397–400. Baz, M.A., Tapson, V.F., Roggli, V.L., Van Trigt, P., Piantadosi, C.A. (1996). Glutathione depletion in epithelial lining fluid of lung allograft patients. Am. J. Respir. Crit. Care Med. 153, 742–746. Becker, S., Dailey, L., Soukup, J.M., Silbajoris, R., Devlin, R.B. (2005a). TLR-2 is involved in airway epithelial cell response to air pollution particles. Toxicol. Appl. Pharmacol. 203, 45–52. Becker, S., Mundandhara, S., Devlin, R.B., Madden, M. (2005b). Regulation of cytokine production in humal alveolar marcophages and airway epithelial cells in response to ambient air pollution particles: Further mechanistic studies. Toxicol. Appl. Pharmacol. 207 (2 Suppl.), 269–275.

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Index α-catenin 153, 161, 162, 164, 166, 171 β-catenin 149, 153, 161, 162, 166, 169–171 A actin 8, 55, 153, 154, 171 cytoskeleton 2, 8, 138, 147, 153–155, 161, 162, 172, 173, 175 polymerization 162, 171 activated oxygen species (AOS) 87–93, 95– 98, 101, 103, 108 production 90, 91, 96, 97, 101 activated protein C (APC) 150, 165, 171, 172 activation of AP-1 200–202 lysosomal 18 of NADPH oxidase 150, 151, 163, 173, 175 activator protein-1 (AP-1) 191, 192, 194, 199–203, 248 activity 200 sites 200, 201, 250 transcription factor 200–202 active Na+ transport 50, 131, 133, 134, 136, 139, 140 acute lung injury (ALI) 50, 53, 61, 136, 140, 147, 148, 150, 151, 154, 155, 161, 170, 173, 174 acute respiratory distress syndrome (ARDS) 51, 54, 131, 133, 141, 148, 155, 173, 246 adenine nucleotide translocase (ANT) 211, 225, 228–231 adherens junctions (AJs) 147–150, 153–155, 158, 161, 162, 164, 166, 167, 169–172, 175 components 169, 170 advanced glycation end products (AGEs) 149, 162, 175 aging 15, 18, 116, 122–124, 192, 193, 221, 222 airway 47, 58, 82, 244–247, 249 epithelial cells 72, 76, 248, 250, 251

function 243, 244 epithelium 243, 245–248, 250, 251 ALI, see acute lung injury alveolar epithelial cells 43–45, 47, 49, 54, 58, 133, 137, 138, 140, 141, 203 epithelium 49–51, 53, 61, 133, 139, 141 fluid clearance (AFC) 49, 50, 53, 54, 61, 136, 137 reabsorption 45, 131–134, 136, 138, 140, 141 hypoxia 136 Alzheimer’s disease 4, 15, 17, 18 amiloride 46, 47, 49, 50, 53–56, 60, 61 angiogenesis 100, 147, 156, 192 animal models 92, 93, 105, 106, 245 ANT, see adenine nucleotide translocase antimycin 95, 96, 138, 212, 213 antioxidants 17, 36, 37, 132, 141, 157, 168, 245, 248–250 enzymes 15, 25, 193, 247 non-enzymatic 243, 245 AOS, see activated oxygen species apical compartment 80 membranes 43, 45, 55, 58, 61 apoptosis induction of 107 suppression of 100, 101, 103 ARDS, see acute respiratory distress syndrome asbestos 191–193, 197, 198, 202, 246 fibers 193, 194, 202 asthma 52, 157, 243–245, 248, 249 atherosclerosis 15, 148, 150, 163, 165, 175 ATP 3, 72, 118, 124, 212, 226–228 ATPase 44, 113, 114, 119 B barrier 4, 26, 27, 29, 35, 153, 167 dysfunction 154, 155, 168, 172 basolateral membranes 43, 47, 55, 137, 140 bestrophin 75 257

258 biological functions 1, 3, 132, 148 targets 51, 52 biomembranes 1, 24, 25, 27, 36 biophysical properties 46, 48, 49 birth 4, 44, 47, 50, 92, 93, 246 blood 5, 148 breast cancer cells 199 C cadherin 164, 169, 171 calcium 14, 72, 74, 115, 117 channels 14, 114–117, 165, 173, 175, 226 calmodulin 117, 118, 125, 171 cancer 15, 16, 87, 88, 90, 92, 97, 99–103, 105, 106, 108, 119, 140, 192, 193, 203 cells 87, 99–101, 103, 108 carbonate radical 23, 25, 32, 38 carcinogenesis 99–101, 192 cardiolipin 17, 218, 221, 223 carotid body 93, 95 carrier proteins 4, 5 cell death 17, 87, 100, 101, 115, 148, 193, 196– 198, 200, 201, 228 membrane 2, 6, 12, 15, 30, 57, 73, 89 proliferation 148, 196–198, 200, 203 signaling 24, 25, 115, 119, 191 surface 58, 76, 150, 169 survival 8, 10, 11, 13, 14, 196 types 3, 7, 8, 75, 87, 91, 162, 193, 196, 197, 199, 201, 243 cell-cell adhesions 147, 149, 150, 153, 166, 168 contacts 153, 156, 157, 164, 166, 169, 171, 172 junctions 7, 147–149, 153, 155, 166, 167, 171, 175 cellular processes 8, 13, 198, 200 channels calcium 14, 114–117, 165, 173, 175, 226 chloride 71, 72, 74, 76, 82 potassium 45, 87, 91, 93, 95, 107, 117, 226 protein 25, 26, 29 sodium 43, 45–51, 57, 61, 131, 133, 141 epithelial 46, 51 voltage-gated 113–115, 117, 125 chemical effects 23, 25, 35 chloride channel proteins 71, 72, 75 channels 71, 72, 74, 76, 82

Index ions 43–45 cholesterol 3, 4, 27, 28 cigarette smoke (CS) 203, 244, 246, 247 cyclophilin 228, 229 cysteine 62, 76, 91, 198, 201, 229 residues 57, 136, 196, 198, 199, 229, 231 cystic fibrosis 15, 50, 71, 72, 76, 244, 246, 249 cytochrome 17, 100, 101, 137, 150, 151, 213, 217, 223–225, 228 cytokine production 245 cytoplasm 14, 16, 88, 93–95, 97, 99–101, 115, 198, 246 cytosolic components 135, 148, 151 conductance 46, 56, 61, 72, 74, 77 confluent monolayers 55, 59, 171 conformations 124, 125, 229 conjugate acids 24, 25, 31, 32 D dephosphorylation 170, 198, 199 diabetes 4, 148, 150, 163, 165, 175 mellitus 17, 162, 163, 174, 175 dichloroacetate 102, 104 diesel exhaust particles (DEP) 194, 203 diffusion 24, 29, 31, 34, 35, 51, 76 coefficient 26, 34, 35 disease 15, 17, 18, 72, 83, 87–90, 92, 97, 105, 107, 108, 116, 140, 148, 175, 192, 203 Alzheimer’s 4, 15, 17, 18 processes 175, 212 states 15, 16, 90, 91, 93 distal lung epithelial cells 44, 59 dopamine 48, 131, 140, 141 E ECs, see endothelial cells electrochemical gradient 5, 14, 44, 45 electron acceptor 51, 215, 216 ELF, see epithelial lining fluid endothelial barrier dysfunction 149, 154, 155, 169, 170, 175 function 150, 151, 153, 157, 166, 173 integrity 147, 150, 153, 154 cells (ECs) 52, 140, 147, 148, 150–153, 156, 158–160, 162, 164, 166, 167, 170, 172–175, 199, 219, 248 contraction 147, 148, 155, 156 permeability 147–149, 154–158, 160, 162, 165–168, 170, 175

259

Index increased 150, 154, 155, 158, 161 enzymatic antioxidants 15, 25, 193, 243, 245–247 enzymes 51, 52, 91, 99, 100, 124, 125, 212– 215, 217, 218, 220, 246 epidermal growth factor receptor (EGFR) 8, 136, 195–197 epithelial cells 5, 7, 45, 61, 75, 192–194 distal lung 44, 59 lining fluid (ELF) 245, 246 sodium channel 46, 51 extracellular matrix 6, 8, 158, 161, 196 ezrin 160, 161 F fetal fluid 44, 50 FFAs, see free fatty acids fibers 191, 193, 195, 197, 199 focal adhesion kinases (FAK) 149, 156, 160, 164, 166, 167 free fatty acids (FFAs) 88, 230 function biological 1, 3, 132, 148 membrane 1, 4 G gas exchange 44, 133 gene transcription 100, 200, 201 glycolysis 99 glucose 4, 5, 88, 97, 108 glutathione 12, 136, 202, 220, 228, 245, 247 G-proteins 76, 148, 149, 152, 175, 195 gradient 5, 132, 212, 213 electrochemical 5, 14, 44, 45 growth factor 8, 136, 148, 149, 152, 168, 169, 175, 193, 198, 200, 248 receptors 11, 195 H health 87–89 hydrogen peroxide 12, 15–17, 23, 25, 30, 39, 57, 122, 124, 135, 147, 148, 243 hydroxyl radical 16, 23–25, 31, 39, 57, 151, 193, 243 hyperoxia 48, 50, 131, 139–141, 215 hyperpolarized mitochondria 92, 97, 101, 105 hypoxic conditions 99, 199, 216, 217, 220 pulmonary vasoconstriction (HPV) 87, 88, 92–98

effectors 93, 94 ROS burst 216 I inactivation 115, 121, 122, 124, 153, 164, 170, 196, 198, 199, 217 oxidative 149, 154, 165, 167, 170, 219 increased endothelial permeability 150, 154, 155, 158, 161 increased superoxide production 90 induction of apoptosis 107 inflammation 62, 147, 148, 151, 159, 161– 165, 167, 171, 175, 192, 194, 201, 248, 249 vascular 162, 163 inflammatory diseases 148, 150, 151, 248 inhalation 193, 194, 197, 202 inner membrane surface 3, 5, 223 inner mitochondrial membrane 17, 226, 229 integrins 7, 159, 161 ion transport 1, 4, 5, 29, 45 isoforms 117, 119, 120, 132, 136, 192, 195, 215, 229 K kidney 4, 7, 94, 96, 132, 229, 230 mitochondria 96 kinases 8–10, 13, 73, 136, 148, 160, 165, 168, 169, 195, 196, 227 signal-regulated 195 Kv channels 88, 89, 91–94, 98, 101, 103, 106 L leukocyte 159, 160, 163, 167 adhesion 159 lipid 2, 4, 15, 17, 18, 25, 33, 35–37, 52, 193, 243, 244 bilayers 1, 4, 6, 7, 31, 33–35, 39 hydroperoxides (LOOH) 35–38 membranes 25–27, 30, 33, 37 peroxidation 16, 23, 35–39, 202 rafts 2, 3 lysosomal activation 18 lysosomes 5, 15, 18, 139 low density lipoprotein (LDL) 31, 38 lung cancer 193 epithelial cells 47, 61, 197, 199, 202 mitochondria 96 vascular permeability 156, 161

260 M macrophages 148, 155, 159, 174, 193, 194, 243–245, 248 MAP, see mitogen-activated protein MAPK, see mitogen-activated protein kinase cascades 195, 196, 198, 200 pathways 10, 12, 13, 196, 201 signaling 191, 196, 197 pathways 8, 9, 12, 199 mechanical forces 6, 7 mechanotransduction 4, 6–8 membrane apical 43, 45, 55, 58, 61 basolateral 43, 47, 55, 137, 140 components 25, 135 functions 1, 4 inner 3, 14, 88, 221, 223, 226, 228, 231 lipids 25–27, 30, 33, 35, 37, 39, 212 permeability of 26, 27, 29 proteins 25, 39, 113 resistance 29, 30 systems 1, 2, 4 tension 6, 7 mesothelial cells 197, 202 metabolic phenotype 100, 101 rate 221, 222 metabolism 74, 87, 91, 97, 99, 100, 108, 114, 147, 151, 194, 198, 221, 247 metal centers 37, 215, 216 microparticles 163 microtubules 3, 171, 172 mitochondria hyperpolarized 92, 97, 101, 105 isolated 96, 215–217, 227 mitochondrial Ca2+ uniporter (MCU) 114, 115, 225, 226 damage 18 depolarization 101, 103, 107 DNA 17, 90, 138 electron transport chain 134, 151, 192, 215, 231 function 14, 17, 18, 88, 89, 91, 94, 101, 217, 221, 231 membrane lipids 211, 221, 223, 224 membranes 1, 3, 17, 88, 94, 100, 103, 211, 221, 224, 228, 231 inner 17, 226, 229 respiration 96, 211, 217 ROS 137–140, 216, 226, 228 target sequence 214, 215

Index transition pore (MTP) 101, 103 uncouplers 95, 214, 227 mitogen-activated protein (MAP) 9, 58 mitogen-activated protein kinase (MAPK) 8, 10, 11, 62, 136, 157, 160, 161, 167, 191, 192, 195, 197–201 phosphatase 191, 192, 198, 199 molecular identity 93, 224, 226, 227 monolayers, confluent 55, 59, 171 MTP, see mitochondrial transition pore mucus hypersecretion 249, 251 secretion 250, 251 muscle, skeletal 99, 119, 120, 122, 124, 226, 229, 230 N NADPH 135, 141, 152, 157, 165, 173, 244, 245, 250 oxidase 25, 93, 131, 134, 135, 139, 140, 147, 148, 150–152, 154, 157–159, 163, 165, 167, 173–175, 192, 249 Na,K-ATPase 43–45, 47, 54, 58, 131–134, 137–141 neutrophil 51, 52, 135, 148, 149, 174, 243, 244, 248, 249 elastase (NE) 250 nitration 32, 52, 120, 191, 220 nitric oxide 6, 15, 23, 24, 33, 36, 43, 51, 72, 115, 151, 199, 219 synthases (NOS) 24, 71, 72, 76, 115, 150, 214, 215 activation 72–75, 82 inhibition 74, 75 nitro-alkenes 224, 230 nitrogen dioxide 23, 31, 37, 52, 193 oxides (NOx) 51, 52, 71, 72, 74–76, 82 nitrosation 219, 220 nitrosylating agents 78, 83 nitrosothiols 82 non-enzymatic antioxidants 243, 245 normal lung function 7, 8 NOS, see nitric oxide synthases NOx, see nitrogen oxides O oocytes 46, 47, 55, 56 overexpression 134, 138, 166, 172, 198, 201 oxidant 35, 37, 52, 114, 117, 121, 124, 125, 155, 168, 191, 194, 196–199, 201–203, 244, 245, 248, 249, 251

Index stress 192, 200, 203, 243, 245, 247–250 oxidase 17, 100, 216, 217, 224, 244 oxidation 15, 24, 38, 91, 116, 117, 119, 120, 122–125, 161, 191, 196–198, 218 oxidative damage 16–18, 35–37, 193, 198, 199, 221, 222 inactivation 149, 154, 165, 167, 170, 219 modifications 43, 57, 113, 116, 117, 119– 121, 124, 125, 218 stress 2, 3, 8–18, 113, 114, 116, 122, 124, 136, 151, 157, 163, 171, 193, 194, 197–199, 201, 202, 229, 230, 245, 246 oxidizing species 23–25 oxygen 15, 23, 24, 27, 35, 89, 92, 95, 99, 135, 139, 151, 152, 192, 243, 244, 249 P PA, see pulmonary arteries PAH, see pulmonary arterial hypertension particulate matter 131, 140, 141, 192, 249 partition coefficient 26, 33, 34, 220 PASMC, see pulmonary artery smooth muscle cells perhydroxyl radical 25, 28, 30, 32 permeability 1, 4, 25–27, 29, 30, 32, 35, 39, 46, 118, 156, 158, 162, 167, 168, 170, 175 coefficients 29–31 of membranes 26, 27, 29 permeation 26, 27, 29–32 peroxisomes 1, 4, 15, 16, 28, 192 peroxynitrite 12, 23, 24, 31, 32, 37, 38, 51, 52, 56, 114, 120–124, 151, 193, 219, 230 phospholipid membranes 33 phosphorylates 9–11, 91, 167 phosphorylation 6, 54, 58, 72, 73, 136, 138, 147, 148, 150, 152, 153, 156, 160, 161, 167, 169, 172, 197, 200 pyruvate 3, 88, 99, 102, 103 PKC see protein kinase C activation 12, 14, 57, 168, 175 plasma membrane 1–3, 5, 14, 27, 57, 58, 88, 98, 114, 115, 117, 131, 132, 135, 136, 140, 141, 203 platelet-derived growth factor (PDGF) 136, 164, 193 polycystic kidney disease (PKD) 7, 8

261 potassium channels 45, 87, 91, 93, 95, 107, 117, 226 proliferation 11, 13, 88, 91, 92, 101, 192, 193, 198, 201 protein carrier 4, 5 channels 25, 26, 29 complexes 1, 147, 148 function 113, 114, 124, 196 kinase 10, 118, 136, 168, 172, 196, 199, 217 C (PKC) 10, 43, 57, 58, 62, 136, 149, 156, 168, 175 regulatory 71, 117, 118 structural 55 transmembrane 91, 132 uncoupling 214, 229 protein-tyrosine phosphatase (PTPs) 149, 154, 167–169, 198 PTPs, see protein-tyrosine phosphatase pulmonary arterial hypertension (PAH) 87, 88, 90, 92, 93, 97, 102, 105–108, 194 pathology 105, 106 arteries (PA) 89, 93, 95–97, 105, 108 artery smooth muscle cells (PASMC) 91, 93, 95, 97, 105–107 circulation 92, 94, 95, 105, 133, 136 edema 44, 50, 131, 133, 147, 148, 154, 156, 163 pumps 5, 14, 45, 119, 132, 212, 224 R reactive lipid species (RLS) 223, 230, 231 reactive nitrogen species (RNS) 15, 24, 192, 194, 217–221, 223–225 reactive oxygen-nitrogen species (RONS) 23–27, 31, 35, 37, 39, 43, 51–54, 57 reactive oxygen species (ROS) 4, 12, 13, 15, 16, 18, 114, 116, 131, 134, 137, 147, 148, 192, 215, 243, 244 generation 16, 139, 155, 163, 165, 202, 212–219, 222, 226, 231, 244, 248 production 15–18, 136, 138, 140, 151–157, 160, 161, 163, 174, 215, 243, 251 reactive species 24, 26, 35, 36, 39, 53, 57, 116, 124, 193, 211, 212, 218 receptor tyrosine kinases (RTKs) 136, 195, 196, 203 red blood cell membranes 27–30 release channels 114, 117

262 renal arteries (RA) 94, 96 respiratory burst 197, 244 chain 90, 212, 213, 217, 219, 220, 223– 225, 230 distress syndrome, acute 131, 133, 173 ryanodine receptor 113, 117–119, 125, 211, 226 RLS, see reactive lipid species RNS, see reactive nitrogen species RONS, see reactive oxygen-nitrogen species ROS, see reactive oxygen species rotenone 90, 95, 96, 138 S sarco(endo)plasmic reticulum calcium ATPase (SERCA) 15, 113, 114, 119–125 activity 119–121 inactivation 122, 124 sarcoplasmic reticulum (SR) 119–121, 226 sensors 87–89, 93, 95, 98, 171 shear 162 stress 148, 149, 152, 160, 162, 170, 175 signal-regulated kinases 195 signal transduction 1, 4–6, 48, 119, 149, 192, 200, 202, 251 signaling molecules 10, 13, 131, 132, 135, 136, 148, 216, 243, 251 pathways 8, 12, 15, 136, 149, 172, 196, 198, 201, 227 proteins 191 signals 4–7, 14, 161, 196, 217, 228 smoke cigarette 203, 244, 246, 247 tobacco 200, 219, 249 smokers 193, 247–249 smooth muscle cells (SMC) 72, 74, 91, 94, 97, 105 S-nitrosation 113, 115, 116, 220 S-nitrosylation 73, 82 S-nitrosoglutathione 59, 73 sodium channels 43, 45–51, 57, 61, 131, 133, 141 SR, see sarcoplasmic reticulum steroids 48, 49 structural proteins 55 superoxide 23–25, 29, 32, 36, 38, 51, 57, 73, 78, 89, 95, 135, 147, 148, 192 dismutase (SOD) 25, 90, 135, 151, 246

Index production, increased 90 surfactant proteins 7, 8 T thioredoxin 12, 17, 136, 151, 191 thrombin 54, 58, 140, 141, 150, 154, 160, 167, 169, 172 tyrosine nitration 32, 52, 73 phosphorylation 136, 147, 154, 156, 158, 160–162, 164, 166, 169, 170, 196 residues 11, 32, 52, 55, 220 tobacco smoke 200, 219, 249 transcription factors 10, 11, 13, 89, 103, 174, 191, 194, 196, 200, 201, 203, 216, 247, 250 transient receptor potential 170 transmembrane domains 91, 122, 152 proteins 91, 132 tumor 11, 99, 100 growth 99, 103, 107 U ubiquitination 57, 197, 216, 247 ubisemiquinone 139, 212, 213, 215 uncoupling 212–214, 224, 230, 231 proteins 214, 229 V vascular endothelial growth factor (VEGF) 150, 155–157, 161, 163, 166–169 inflammation 162, 163 permeability 150, 156, 158, 161, 163, 165, 167 stimulation 156, 171 wall 105, 108, 163 VE-cadherin 153, 155–158, 160–162, 165, 166, 169–171 complexes 154, 162, 169 function 164, 166 tyrosine phosphorylation 153, 155, 157, 165, 166 VEGF, see vascular endothelial growth factor vitamins 37, 168, 202, 245, 247 voltage-gated channels 113–115, 117, 125 X Xenopus oocytes 46, 55–58, 61