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

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Author: Sadis Matalon

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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

AMSTERDAM · BOSTON · HEIDELBERG · LONDON NEW YORK · OXFORD · PARIS · SAN DIEGO SAN FRANCISCO · SINGAPORE · SYDNEY · TOKYO Academic Press is an imprint of Elsevier

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First edition 2008 Copyright © 2008, Elsevier Inc. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://www.elsevier.com/locate/permissions, and selecting: Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made

ISBN: 978-0-12-373879-0 ISSN: 1063-5823

For information on all Academic Press publications visit our website at books.elsevier.com

Printed and bound in USA 08 09 10 11 12

10 9 8 7 6 5 4 3 2 1

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 92 IV. The Mitochondria-AOS-Kv Axis, Metabolism and Apoptosis 97 References 108

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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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 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

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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

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.

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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.

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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

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