Principles of Free Radical Biomedicine [1 ed.] 9781620817780, 9781612097732

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Copyright © 2011. Nova Science Publishers, Incorporated. All rights reserved. Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

Copyright © 2011. Nova Science Publishers, Incorporated. All rights reserved. Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

BIOCHEMISTRY RESEARCH TRENDS

PRINCIPLES OF FREE RADICAL BIOMEDICINE VOLUME I

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BIOCHEMISTRY RESEARCH TRENDS

PRINCIPLES OF FREE RADICAL BIOMEDICINE VOLUME I

KOSTAS PANTOPOULOS Copyright © 2011. Nova Science Publishers, Incorporated. All rights reserved.

AND

HYMAN M. SCHIPPER EDITORS

Nova Biomedical Books New York Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

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All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers‟ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works.

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Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Additional color graphics may be available in the e-book version of this book.

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ISSN: 2159-9343 ISBN: 978-1-62081-778-0 (eBook)

Published by Nova Science Publishers, Inc. † New York

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Contents Preface

vii

Acknowledgments

ix

Chapter 1

The Evolution of Oxidative Stress Nick Lane

1

Chapter 2

Oxygen Radicals and Related Species Ohara Augusto and Sayuri Miyamoto

19

Chapter 3

Nitric Oxide and Derived Oxidants Silvina Bartesaghi, Natalia Romero and Rafael Radi

43

Chapter 4

Sulfur-Centered Radicals Christian Schöneich

75

Chapter 5

Redox-Active Metals: Iron and Copper Willem H. Koppenol and Patricia L. Bounds

91

Chapter 6

Protein Oxidation Annika Höhn, Tobias Jung, Stefanie Grimm and Tilman Grune,

113

Chapter 7

Lipid Peroxidation Angel Catalá

137

Chapter 8

Lipid Nitration Lucía Bonilla Cal and Homero Rubbo

161

Chapter 9

DNA Oxidation Jean-Luc Ravanat

185

Chapter 10

Methods of Investigation of Selected Radical Oxygen/Nitrogen Species in Cell-Free and Cellular Systems Jacek Zielonka and Balaraman Kalyanaraman

Chapter 11

Enzymatic Generation of Hypoxia and Steady-State H2O2 Gabi N. Waite, Gunda Millonig, Lee R. Waite, Helmut K. Seitz and Sebastian Mueller

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vi Chapter 12

Contents Chemiluminescence Detection of H2O2 Sebastian Mueller, Gunda Millonig, Helmut K. Seitz and Gabi N. Waite

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Index

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Preface Recent years have witnessed an avalanche of new knowledge implicating free radicals in virtually every aspect of biology and medicine. It is now axiomatic that the regulated accumulation of reactive oxygen species (ROS) contributes to organismal health and wellbeing and that ROS serve as signaling molecules involved in cell growth, differentiation, gene regulation, replicative senescence and apoptosis. That ROS and other chemical species constitute a primary defense invoked by leukocytes and tissue macrophages against invading pathogens is also well established, although novel insights concerning the nuanced regulation of this adaptive response continue to emerge and captivate. Contrariwise, „unresolved‟ oxidative stress is increasingly recognized as a pivotal pathway for cellular dysfunction and death, and a potentially salient target for therapeutic intervention in disorders as disparate as diabetes mellitus, glomerulonephritis, macular degeneration, drug-induced hepatotoxicity, hypertension, asthma, cancer, Alzheimer disease and infertility. Principles of Free Radical Biomedicine is an interdisciplinary text on the biochemistry and cellular/molecular biology of free radicals, transition metals, oxidants and antioxidants, and the role of oxidative stress in human health and disease. These volumes were conceived as a companion to, and significantly expand on, the contents of a course coordinated and offered by the editors to upper undergraduate and graduate students in the Departments of Medicine and Neurology & Neurosurgery at McGill University (Montreal). The cast of contributors is international – invitations having been extended to senior investigators based on their acknowledged expertise and enthusiasm for their respective fields of fundamental, translational and clinical research into free radical mechanisms and their biological consequences. The authors were requested to provide comprehensive reviews of the current literature in their disciplines encompassing, but not limited to, topic „bullets‟ provided by the editors. They were granted sufficient latitude to provide detail from their own laboratories or clinical practices and encouraged to express their personal viewpoints and perspectives where subject matter remains controversial. We felt that implementation of italics to underscore general principles and key concepts and extensive cross-referencing of the chapters would add value and enhance overall readability. The work begins with a provocative evolutionary perspective on free radicals and oxidative stress. The chapters that follow are grouped in three broad sections, each assigned an independent volume: Vol. I, Free Radical Chemistry. The chapters herein provide detailed accounts of the major classes of reactive chemical species, their biological targets, and

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viii

Preface

popular assays for their detection and measurement. Vol. II, Free Radical Biology. Subsections encompass wide-ranging treatises on non-enzymatic antioxidants, antioxidant enzymes, redox signaling, subcellular redox compartmentalization, and the role of the former in mammalian metabolism and physiology. Vol. III, Oxidative Stress and Disease. The roles of ROS and other reactive species, antioxidant defenses and redox-based therapeutics in diseases of the major organ systems are elaborated here, with emphasis on both clinical and experimental material. Chapters on the participation of oxidative stress in exercise and environmental toxicology complete this section. Free radical biomedicine is galloping ahead and, for investigators at the cutting edge of the field, no textbook can replace perusal of the relevant journals and other primary publications. This being said, there remains a vital need periodically to take stock of and assemble the pertinent literature in a fashion that permits the assimilation of cogent (but frequently strewn) new information, and facilitates the cross-fertilization of disciplines and attainment of a more global understanding. In this vein, we hope that the scope and breadth of this compendium will render it a useful resource to basic scientists interested in various aspects of free radical chemistry and biology as well as to clinically-oriented investigators concerned with the role of oxidative stress in the pathophysiology of diverse medical conditions. In Memoriam

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The editors express great sorrow at the sudden passing of Professor Wulf Dröge, a “Redox Pioneer” (Antioxid Redox Signal 2010 Dec 22), an esteemed colleague and a dear friend. He will be sorely missed.

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Acknowledgments We are deeply indebted to numerous mentors, colleagues and students who have nurtured over the years our interest in „Redox Biomedicine‟, and to the Jewish General Hospital/Lady Davis Institute for Medical Research and McGill University for providing a stimulating work environment. K. Pantopoulos: To my wife Giada for her unconditional and sustained moral support and scientific contribution, and to our baby daughter Elena Maria for her engaging smile which was particularly uplifting while editing this book. H. Schipper: To my wife, Rachel and boys, Joshua and David, go my love and gratitude for their unflagging support and encouragement.

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Kostas Pantopoulos Hyman M. Schipper

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 1

The Evolution of Oxidative Stress Nick Lane* Department of Genetics, Evolution and Environment, University College London, Gower Street, London WC1E 6BT, U.K.

1. Introduction

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“Life is nothing but an electron looking for a place to rest.” Albert Szent-György Oxidative stress has evolved in both senses of the word. It has evolved in semantic meaning. When first defined by Helmut Sies in the 1980s [1], oxidative stress was perceived as largely negative and pathological. Since then, it has become clear that the reactive oxygen and nitrogen species (ROS and RNS) that cause oxidative stress are not merely pathological, but serve as signals in many diverse circumstances (see Chapters 2 and 3). More broadly, oxidative stress is a physiological state that elicits a decisive shift in patterns of gene expression, leading usually to its own resolution. Blocking oxidative stress with antioxidants is rarely beneficial; and most studies of antioxidant supplements have indeed proved ineffective, if not actively damaging. This nuanced conception makes it clear that oxidative stress has evolved in the biological sense too. No longer is it seen as a purely pathological state, over which the body or the cell has little control; it is a central part of cellular homeostasis. Cellular redox state, and specifically the activation of redox-sensitive transcription factors by oxidative stress (Vol. II, Chapters 12-14), is beginning to look as central to cell homeostasis as protein phosphorylation. While the interplay between protein thiol oxidation, S-nitrosylation and phosphorylation is still somewhat murky, it is plain that redox state is a critical regulator of *

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cellular respiration, cell cycle, proliferation, differentiation, sexual development, stressresistance, longevity, senescence and apoptosis. All of this certainly evolved, but the evolutionary tradeoffs underlying ROS signalling are little explored. Often they seem to be in conflict, displaying Janus-like qualities. Is nitric oxide (NO) a „good thing‟ or a „bad thing‟? What about mitochondrial biogenesis? Without an evolutionary narrative, the costs and benefits can be confounding. While ROS are no longer perceived as merely damaging, it is far from certain how damaging they actually are, and under what circumstances. Fenton chemistry, lipid peroxidation, protein carbonyl oxidation, DNA mutation (Chapters 5-9) – the old order of free radical biology – are real enough, but many assays are equivocal, and the standard laboratory conditions are not always relevant. For example, the mutation rate (as opposed to the long term evolution rate) of mitochondrial DNA relative to nuclear DNA is unknown to within several orders of magnitude [2], as is the proportion of oxygen escaping from the respiratory chain as superoxide under physiological oxygen tensions of around 3 µM oxygen (1-2% of atmospheric tension). The frequently cited figure of 1-5% ROS leak during respiration, dating back to Britton Chance‟s work in the 1970s [3], is not correct at lower oxygen tensions; but the actual in vivo figure, or its pathophysiological relevance, is unknown. Likewise, the degree to which protein or DNA damage accumulates in aging tissues is difficult to ascertain, because apoptosis removes the evidence. At the same time, it has become clear that free radicals are not the potent bacteriocidal agents long thought. The oxidative burst of neutrophils, which generates superoxide radicals at high levels, via NADPH oxidase, in fact alkalinizes the vacuoles, activating proteolytic enzymes inside [4]. These, not ROS, degrade bacteria. This inevitably begs the question, why use potentially toxic ROS at all? In the same context, it has also become clear that ROS leak from respiratory chains is not a simple by-product of respiration: the proportion of oxygen consumed that is converted into ROS is not fixed, but varies according to the respiratory rate, training, thyroid hormone levels and calorie intake, and between different tissues and across species [5]. Nor does ROS leak seem to be strictly necessary: cytochrome oxidase achieves the complex four-electron reduction of oxygen to water without any measurable ROS leak at all. So again: why, from an evolutionary point of view, are potentially toxic ROS used as signals? The answer does not relate in a simple manner to oxygen toxicity and antioxidant balance, as long assumed, but in a fundamental way to the nature of energy transduction in cells. Oxidative stress, differences in lifespan and susceptibility to age-related disease are all pleiotropic consequences of the fundamental requirement to regulate energy transduction.

2. The Origin and Significance of Chemiosmotic Coupling From the medical point of view, respiration is typically divided into aerobic and anaerobic. Aerobic respiration obviously requires oxygen; anaerobic respiration – or „life without oxygen‟, in Pasteur‟s words – is typically taken to mean anaerobic glycolysis, or fermentation. The distinction, which may be reasonable enough for humans, makes a mockery of evolution. The true, meaningful, distinction is between substrate-level

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phosphorylations, such as fermentation, in which phosphate groups are transferred directly by chemistry, and oxidative phosphorylation, in which electrons are transferred from an electron donor such as glucose (but which could be many other organic or inorganic donors such as Fe2+) via a series of redox centres to a terminal acceptor. In aerobic respiration this acceptor is oxygen, but in anaerobic respiration a range of other electron acceptors are used, from NO to Fe3+ to protons. In all forms of oxidative phosphorylation, the passage of electrons from the donor to the acceptor is coupled to ATP synthesis by way of an intermediary proton gradient across a membrane – chemiosmotic coupling. Rather than chemistry, which is to say reactions between molecules, the process is basically electrical. This is, at bottom, the cause of oxidative stress. Described by Leslie Orgel as „the most counter-intuitive idea in biology since Darwin‟ [6], the mechanism of chemiosmotic coupling was controversial for three decades, a period known as the „ox-phos wars‟, culminating in the award of the Nobel Prize to Peter Mitchell in 1978 [7]. The passage of electrons along respiratory chains – essentially the flow of electricity down electrical wires – drives conformational changes in respiratory proteins (or more simply, electro-chemical loops across the membrane, as in the Q cycle) to generate a proton gradient across the membrane. Driven by the proton-motive force (the combination of electrical potential and pH gradient), the passage of protons through the rotary motor of the ATPase in turn drives ATP synthesis. But why, and how, did such a counterintuitive system evolve? The answer relates to the ability of a gradient to uncouple exergonic reactions from ATP synthesis. Consider the direct reaction of hydrogen with carbon dioxide, known as the acetyl CoA pathway, and probably the most ancient chemolithotrophic pathway in life. In ancient methanogens (archaea) and acetogens (bacteria), this pathway provides both the carbon and energy metabolism of life – there is no need for solar power, primordial soup, ATP or any other accoutrements. The acetyl CoA pathway is exergonic right through to pyruvate, and releases enough energy (captured as ATP) to power all intermediary metabolism. But there is one drawback: kinetics. While thermodynamically probable, the reaction between H2 and CO2 is slow because a kinetic barrier must be surmounted to convert CO2 into formate (CHO2-). In modern organisms, this conversion requires an initial input of energy, usually in the form of ATP. The problem is that, by substrate-level phosphorylation, one ATP must be split to gain one ATP from the reaction: there is no net gain and therefore there can be no growth. Life would be impossible [8,9]. The reaction of H2 with CO2 is by no means alone in „technically‟ not generating enough energy for growth. Many other reactions that do in fact sustain growth – such as the anaerobic oxidation of methane using nitrite (the anammox reaction) – could not do so by substratelevel phosphorylation alone. Put more broadly, if life operated by chemistry alone, it could never have got started. In the absence of oxygen or photosynthesis, there is insufficient energy available from anaerobic reactions to power both carbon assimilation and ATP synthesis by substrate-level phosphorylations alone. There are in fact only five known pathways of carbon assimilation across all life, including the Calvin cycle (used in oxygenic photosynthesis), the reverse Krebs cycle (found in many vent bacteria), and the acetyl CoA pathway. All but the acetyl CoA pathway require an input of energy, in the form of ATP or some equivalent, which is provided by sunlight in photosynthesis and oxygen in the case of chemosynthesis in vents. Because oxygen is derived from photosynthesis, the profusion of life in black smoker vents is ultimately powered by photosynthesis and could not exist

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without it. Only the acetyl CoA pathway, the direct reaction of H2 with CO2, can provide the energy required for growth in the absence of light or oxygen; and even this pathway can only do so by way of chemiosmotic coupling. Why does chemiosmotic coupling make such a decisive difference? Because an exergonic reaction, such as the reaction of CO2 with H2, can be uncoupled in both time and space from an endergonic reaction, such as the reaction of ADP with Pi to give ATP. This means that there is no direct equation between the energy released by one reaction and consumed by the other, as there must always be in substrate level phosphorylations, or indeed, in any form of chemistry. In chemiosmotic coupling, in contrast, the energy released by an exergonic reaction is used to transfer one or more protons across a membrane. So long as the energy released is sufficient to transfer a single proton at least part of the way across the membrane, the reaction can be repeated indefinitely to generate, in the end, a proton gradient. And then that gradient can be used independently to power ATP synthesis, enabling growth. In the case of the acetyl CoA pathway, it remains true that 1 ATP must be spent to overcome the kinetic energy „hump‟; but instead of reclaiming just one ATP, so precluding growth, chemiosmotic coupling makes it possible to gain about 1.5 ATPs per CO2. The nonstoichiometric number gives it all away: stoichiometry is a property of chemistry, not gradients [10]. From this point of view it is no accident that all life on earth is powered by gradients, mostly proton gradients; and even exceptions like fermenting bacteria use their ATP to pump protons to maintain their membrane charge, used in turn to power motility, ionic homeostasis and the uptake of organic matter. Fermentation arose later, of that there can be little doubt. The glycolytic pathway evolved independently in archaea and bacteria, and no fermenters are found anywhere near the base of the phylogenetic tree. The conception of fermenting a primordial soup lacks any thermodynamic basis [9]. Without chemiosmotic coupling, early life would have been impossible, and even today, after four billion years of evolution, the fact that chemiosmotic coupling is as universal as the genetic code goes to show that the mechanism is close to being energetically unimprovable. But the inevitable penalty, at least in the presence of oxygen, is oxidative stress. The daunting complexity of modern respiratory chains long concealed the conceptual simplicity of chemiosmotic coupling – the flow of electrons powers the transfer of protons over a membrane. If membranes, proton gradients or electron-transferring respiratory chains required eons of natural selection to evolve, then they could certainly not have powered early life. But in fact all are provided „free of charge‟ in alkaline hydrothermal vents, which are driven by the global geological process of serpentinization [11]. In serpentinization, seawater reacts with ultramafic magnesium-rich rocks derived from the upper mantle, such a peridotite, containing the mineral olivine. Olivine is hydroxylated to metamorphose into serpentine, hence the name of the process. The dark green mottled mineral serpentinite is commonly used as a building stone, often on the facade of banks or public buildings, including the United Nations. The hydroxylation of olivine oxidises Fe2+ to Fe3+, releasing hydrogen gas and hydroxide ions into solution, and producing enough heat to drive these alkaline fluids back up to the seafloor, where the dissolved minerals precipitate out into the colder seawater to form alkaline hydrothermal vents: porous calcium carbonate towers, riddled with microscopic, roughly cell-sized pores, lined with flimsy aragonite membranes. While their existence and periaxial location was predicted in detail two decades ago by geochemist Michael Russell [12,

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13], the first such vent system was not discovered until ten years later – Lost City Vent, just off the mid-Atlantic ridge [14]. Lost City conforms to most of Russell‟s predictions but does not do so in every way. The two main differences are highly significant. On Earth 4 billion years ago there was virtually no oxygen, and far higher levels of CO2 – anything up to a thousand-fold more. The absence of oxygen meant that iron was in the form of Fe2+, which dissolves in water. The oceans were full of it; later, as oxygen levels rose, Fe2+ was oxidised to Fe3+, which precipitated out as rusty minerals to form vast banded-iron formations; but 4 billion years ago, before all this iron had been precipitated from the oceans as Fe3+ in banded-iron formations the oceans would have been nearly saturated in dissolved Fe2+. Unlike today, therefore, ferrous iron was available for incorporation into vent systems. In ancient alkaline hydrothermal vents, this ferrous iron is likely to have precipitated out with sulfide to form bubbly membranous films of iron sulfur minerals, perhaps mixed with aragonite. Fossils of such structures exist, albeit only 350 million years old (the ocean crust is constantly recycled by tectonics, making older structures rare) and they have been reproduced in the lab [15]. Critically, on the molecular scale, such minerals form into iron-sulfur clusters (Vol. II, Chapter 19) with an identical structure to the Fe-S clusters still found at the heart of respiratory complexes today, notably in complexes I and II. Recent functional studies of respiration suggest that electron transfer is independent of the protein groups that embed these Fe-S clusters; what matters is the distance between clusters [16]. If the distance between successive redox centres is less than around 14 Ångstroms, electrons „tunnel‟ via quantum processes down the respiratory chain. Thus, electron flow depends on a property – distance – that is geometric, and geochemical, rather than biological in provenance. Furthermore, Fe-S clusters have the important intrinsic factor of transferring single electrons, by dint of the electron configuration of the transition metal iron. Free-radical chemistry is the chemistry of single electrons, not the electron pairs characteristic of covalent bonds, and it began with Fe-S clusters precipitating in ancient alkaline vents, doing what they do today: transferring single electrons from the alkaline fluids emerging from serpentinized rocks into the acidic ocean waters above. Acidic. That is the difference CO2 made. When dissolved in water, CO2 forms carbonic acid, hence the concerns today that global warming, in acidifying the oceans, is destroying the delicate reef systems that depend on slightly alkaline waters to precipitate carbonates into corals. Back then, the pH of the oceans was likely to have been in the range of 5 to 6 [17]. The pH scale, of course, is defined in terms of proton concentration, each pH unit representing a tenfold difference in proton concentration. As the alkaline fluids percolated into acidic oceans, through a labyrinth of interconnected micropores, lined with hydrophobic iron-sulfur membranes, the vent system would have developed a natural proton gradient of at least 4 pH units across the mineral walls – a difference in proton concentration of about 10,000-fold, equating to a membrane potential of 200-400 mV, with the correct polarity, remarkably similar to that across bioenergetic membranes today. How the first cells began to tap the energy of this natural proton gradient is beyond the scope of this Chapter; suffice to say that polyphosphates, such as ATP and pyrophosphate, form under acidic conditions at low water activity (in hydrophobic membranes) whereas their hydrolysis is favoured under alkaline aqueous conditions [9]. It is therefore plausible that natural proton gradients could have driven the cycling between pyrophosphate and phosphate, or ATP and ADP, in the vent environment.

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Whatever the mechanism, the first cells could not have left the vents without first mastering the art of chemiosmotic coupling – nothing else could have provided the necessary energy. Luckily the vents also equipped the first cells with all the necessary tools – proton gradients, electron-conducting iron-sulfur clusters, and charged membranes. When the first prokaryotic cells did emerge, these were the tools of their trade. Without them, oxidative stress would not exist. With them, the stage was set for the second act: photosynthesis.

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3. The Origin and Consequences of an Oxygen Atmosphere Respiration, by necessity, evolved early. Not aerobic respiration, but anaerobic respiration – the transfer of electrons from an electron donor (probably hydrogen) to an acceptor (initially CO2). At some point early in evolution, electrons would have been transferred to a range of other electron acceptors, as energy metabolism diverged from carbon assimilation. By its nature, chemiosmotic coupling splits exergonic reactions that provide energy from endergonic reactions which utilise that energy in carbon metabolism. Simply substituting alternative electron acceptors such as NO or Fe3+, probably plentiful on the early Earth, required no change in membrane structure, and may have been among the earliest forms of prokaryotic specialisation [18]. As a general rule, it is fair to say that prokaryotes can be classified not by their morphology but by their metabolic capabilities. And the most significant of those was photosynthesis. Photosynthesis reverses respiration, not only in the global chemical sense (the equation for aerobic respiration can be represented as CH2O + O2 → CO2 + H2O and oxygenic photosynthesis CO2 + H2O → CH2O + O2) but also in particulars. In oxygenic photosynthesis, for example, electrons are transferred from water – split by chlorophyll photo-oxidised by the sun – via an electron-transport chain that is exactly analogous to the respiratory chain, ultimately onto CO2 to form sugars. The flow of electrons drives the transfer of protons across the thylakoid membranes to generate a proton gradient, which in turn drives ATP synthesis. Thus, in a curious way, photosynthesis also reverses the specialisation of metabolism into carbon assimilation and energy metabolism, reuniting them in a system driven by the power of light. Of course, cyanobacteria, and later algae and plants, are also capable of normal respiration to provide energy during darkness or to bleed off excess reduction potential. Like respiration, photosynthesis is not particular about the source of electrons. Indeed, drawing on the power of the sun it has even less need to be. On the early Earth, H2S and Fe2+ would have been major electron donors, as they are in many bacteria today. The principle is exactly the same: photo-oxidation of chlorophyll transforms it into an oxidant, which can strip electrons from many sources, passing them ultimately onto CO2. Only when water itself is used as an electron donor – a difficult operation even for cyanobacteria, and never yet achieved by any other form of bacteria, or for that matter, solar scientists – is the waste product oxygen. If H2S is the electron donor, the waste product is sulfur. While the origin of oxygenic photosynthesis is obscure, it probably arose 3 billion years ago, if not earlier, by coupling in series the two reaction centres used by anoxygenic photosynthesizers, PSI and PSII [19, 20]. But however it evolved, photosynthetic water-splitting utterly transformed the

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planet. Drawing on water as a fuel, rather than reduced chemicals derived from volcanic and hydrothermal processes, probably increased global biomass 10-fold [21]. The waste, molecular oxygen, initially reacted with sulfur, iron or methane but ultimately accumulated in the atmosphere, from around 2.4 billion years ago in the Great Oxidation Event, perhaps precipitating the first global ice age – a „snowball earth‟ – as it did so [22]. Oxygen changed the world. The extent to which complex life is possible at all without oxygen is an interesting question, thrown into sharp relief by the recent discovery of animals, Loriciferans, that complete their life cycle in the absence of oxygen [23]. In fact, oxygen produces only about an order of magnitude more power than fermentation; and the difference between aerobic and true anaerobic respiration is somewhat less than that. While this is substantial, it is orders of magnitude less than the difference made by mitochondria, as discussed later in this Chapter; and probably differences in nutrient availability or concentration gradients outweighed any metabolic advantages of oxygen, at least among bacteria. Oxygen hardly wrought a global revolution in prokaryotic physiology. Even in the presence of oxygen, no prokaryote ever came close to evolving the morphological complexity of eukaryotes. In this context, the evolution of aerobic respiration may have made a difference, but the most immediate impact of the rising tide of oxygen was its juxtaposition with the electron-transport chains of bacteria, all of which transfer single electrons. The reactivity of oxygen, of course, is limited by kinetics in much the same way as CO2. If it were not so, the biosphere, even the atmosphere, would spontaneously combust – as it would if it were filled with singlet oxygen. The kinetic limitation on the reactivity of oxygen relates to its unusual electron outer orbital structure, giving molecular oxygen two electrons in parallel spin. Unlike singlet oxygen, triplet oxygen is unable to accept a pair of electrons, and must be reduced by one electron at a time – by single-electron donors, among which iron is prominent. Because the roots of respiratory chains are in geochemistry, notably the Fe-S clusters already discussed, all these respiratory chains become badly insulated wires in the presence of oxygen. Oxygen is hardly toxic if left to itself; but it is readily activated in the presence of the very respiratory chains that are necessary for life. ROS leak has more to do with the speed of electron flow down electron transport chains than it does with the concentration of oxygen itself. In general, ROS leak is lower in state III respiration (when ATP consumption is fast) than it is in state IV respiration, when electron flow is limited by ADP deficiency [5]. If the respiratory complexes become highly reduced, they become more reactive with oxygen; and the higher membrane potential can drive electrons in reverse back into complex I, again increasing the rate of ROS leak. Without compensation, then, ROS leak is largely defined by poor growth: by a low demand for ATP and highly reduced respiratory complexes. There are various ways out of this „high-voltage‟ situation, from mild uncoupling to complete depolarization of the membrane, or the use of alternative oxidases, which pass electrons directly on to oxygen, without coupling to proton translocation. All of them, in effect, short-circuit the membrane potential, enabling faster electron flow, less reduced respiratory complexes and lower ROS leak. Just how important such mechanisms might be in prokaryotes is hinted at by the sheer scale of „energy spilling‟ in bacteria, an apparently wasteful frittering away of up to 50% of cellular energy charge, for unknown reasons [24]. It is plausible that such energy spilling works to short-circuit overcharged membranes in the presence of oxygen, so restricting ROS leak and cellular damage.

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In this context, it is revealing that the only bacteria to have evolved a form of controlled cell death analogous to apoptosis in eukaryotes (Vol. II, Chapters 25 and 26) – right down to the use of metacaspase enzymes, which are closely related to metazoan caspases – always deal with oxygen. The best example is cyanobacteria, which form oceanic blooms on a similar scale to eukaryotic algae. Like algal blooms, cyanobacterial blooms often disappear overnight, in effect killing themselves in response to some stimulus, usually a viral infection. But viruses are not the only cause: light stress and iron deficiency can also literally liquefy cyanobacterial blooms – and the common denominator in all these cases is oxidative stress, brought about by ROS leak [25]. Why single-celled algae and cyanobacteria kill themselves is a complex question, and can only be interpreted in light of their differentiation within colonies – ROS leak can signal cell death or differentiation into spores or biofilms. Selection can therefore be seen as acting at the group level, where the bloom has at least some of the properties of a multicellular organism, assisted by the genetic relatedness of the clonal cells in the bloom; or in terms of selfish genes. But regardless of the genetic interpretation, perhaps the most significant factor determining the fate of the bloom is the potential for serious cellular damage in the presence of oxygen. Only bacterial cells capable of oxygenic photosynthesis or aerobic respiration appear to undergo controlled cell death in the context of blooms [25]. Oxygen introduces a new penalty for failure, controlled cell death, that later played a central role in the evolution of true multicellular organisms. The basic problem, which is central to eukaryotic evolution too, is that the rates of photooxidation and electron transfer, being essentially quantum events, differ from the rates of chemical reduction and carbon assimilation. This means that conditions such as high light intensity (which rapidly photo-oxidizes chlorophyll), low temperatures (electron transfers are barely slowed, but chemical reactions are much slower), and iron deficiency (leading to poor respiratory stoichiometry) all cause high ROS leak. If this high ROS leak is not brought under control quickly, the caspase enzymes are activated – significantly by the loss of the respiratory carrier cytochrome c, in plants as well as animals – and the cell is eliminated. Controlled cell death offers the advantage of recycling scarce nutrients, and so can be beneficial to the larger grouping, whether an organism, a colony, or selfish genes. Much the same problems affect the respiratory chains of non-photosynthetic aerobic bacteria, such as some α-proteobacteria, among them the free-living ancestors of mitochondria, which likewise are capable of controlled cell death using metacaspase enzymes. The later development of apoptosis in metazoans makes use of enzymes that are bacterial in ancestry, notably the caspases, but also the Bcl-2 family and other mitochondrial apoptotic proteins [26]. The point is that the evolution of metazoan cell death – and with it the complex developmental programs that require apoptosis – all build on a system that evolved in relatively complex clonal bacteria capable of an apoptotic-style of cell death in response to oxidative stress. The single greatest danger is the failure to pass electrons on swiftly down respiratory chains, resulting in highly reduced complexes in an aerobic atmosphere. The way in which these factors played out in the respiratory chains of eukaryotes may have been one of the most significant selective forces in eukaryote evolution.

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4. Oxidative Stress and the Chimeric Origin of Eukaryotes All truly complex life on Earth is composed of eukaryotic cells. All eukaryotes are closely related in cellular structure, and this structure is totally unlike bacteria or archaea. All eukaryotes by definition share the nucleus, with its double membrane, pitted with large protein pore complexes. All have straight chromosomes, telomeres, centromeres, chromatin structures, introns and exons, mitosis, meiosis, reciprocal sex, dynamic cytoskeleton, endoplasmic reticulum, lysosomes, mitochondria and so on: undoubtedly these complex traits were all inherited from a common ancestor, which must already have been a complex cell quite unlike any known prokaryote. These detailed similarities reach into the deepest structure of genes, with the same introns occupying the same site in the same gene in fungi, algae, plants, animals and protists. The deep unity of eukaryotic cells is arguably evidence for the evolution of sex in the earliest eukaryotes; but whatever the cause, the sheer number of shared traits testifies to the common and unique origin of eukaryotic cells. Unique, because if complex cells had arisen more than once, then there should be various disparate types of eukaryote today, each with a spectrum of different traits, unless all fell extinct without trace. While it is not possible to rule out this possibility, there is no evidence in its favour, and much to suggest a genuinely unique origin. The most significant piece of evidence testifying to the unique origin of eukaryotes is the realization, over the last two decades, that all known eukaryotes either currently possess, or once had and later lost, mitochondria. There are in fact around a thousand species of apparently primitive single-celled eukaryotes that do not have mitochondria – a paraphyletic grouping described as „archezoa‟ by the cell biologist Tom Cavalier-Smith [27]. These species – mostly parasites such as Giardia and entamoeba, albeit with free-living relatives – were hypothesized to be representatives of the earliest eukaryotes, primitive phagocytes that had never acquired mitochondria, but which survived in marginal niches. But closer scrutiny of their cell structure (all contain double-membraned structures derived from mitochondria, known as hydrogenosomes or mitosomes) and their genomes (which contain genes derived from mitochondria) betrayed the fact that these apparently primitively amitochondriate groups had once possessed mitochondria, but had later lost them in the course of specializing into anaerobic environments [28]. This discovery has two major implications. First is the fact that the niche itself is perfectly viable: it is occupied by bona fide amitochondriate eukaryotes, and from their cell structure and diverse habitats there is no obvious reason why they should have driven all genuinely primitively amitochondriate eukaryotes to extinction. Positing extinction as an explanation for the unique origin of eukaryotic cells is therefore, at the least, counter to Occam‟s razor. Second, the fact that all eukaryotes once possessed mitochondria pushes the origin of mitochondria and the origin of the eukaryotic cell back to the same time frame, and plausibly the same event, as first proposed by evolutionary biologist Bill Martin [29]. There is now large-scale genomic evidence that this is exactly what happened: the eukaryotic cell originated in some kind of chimeric endosymbiotic event between two prokaryotes, the host cell being an archaeon and the endosymbiont the ancestor of the mitochondria [30, 31]. Exactly what benefit mitochondria brought is not obvious, despite 40 years of research since Lynn Margulis marshalled the evidence to demonstrate the bacterial origin of mitochondria [32]. The advantage was not aerobic respiration, as many prokaryotes

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respire aerobically, and many mitochondria are anaerobic, notably among protists and fungi. Nor did mitochondria protect against oxygen toxicity: mitochondria, being full of respiratory chains, are among the most potent free-radical generators known (Vol. II, Chapter 15). They were part of the problem, not the solution. Mitochondria did not enable anaerobic host cells to adapt to rising oxygen levels as anaerobic environments disappeared; on the contrary, rising oxygen levels actually gave rise to sulfidic „Canfield‟ oceans, in which the oceans stratified in the same manner as the Black Sea today, with the subphotic zone remaining anoxic for more than a billion years, from 1.8 to 0.7 billion years ago – the period during which eukaryotes almost certainly evolved [33]. And while it is true that mitochondria compartmentalized respiration within the cell, many prokaryotes internalize their bioenergetic membranes in circumscribed regions of the cell, including cyanobacteria, nitrosococcus and nitrosomonas, so compartmentalization per se could not have been the decisive advantage either. That decisive advantage probably lies in the mitochondrial genes. All eukaryotes that have retained the capacity for oxidative phosphorylation have also retained a small genome – a core set of genes encoding predominantly integral inner membrane proteins essential for respiration. Mitosomes and hydrogenosomes have lost the capacity for oxidative phosphorylation and almost invariably also lost the complete mitochondrial genome, although in one single case the retention of a hydrogenosome genome was instrumental in establishing the mitochondrial ancestry of hydrogenosomes . While the reasons for the retention of a mitochondrial genome are beyond the scope of this Chapter, the likely answer, as argued by biochemist John Allen, is that the mitochondrial genes form part of a necessary regulatory loop that enables highly hydrophobic proteins to be transcribed and translated quickly and specifically, immediately adjacent to the bioenergetic membranes they service, in response to sudden changes in membrane potential, substrate availability, or oxygen tension [34, 35]. A large body of data shows that the rate of cell respiration depends on the copy number of mitochondrial DNA (mtDNA), with active cells having more copies of the genome, and vice versa. Cells depleted in mtDNA have a low respiratory capacity, while mutations that cause mtDNA depletion are typically associated with mitochondrial diseases [36]. Likewise, the rate of respiration is controlled directly by transcription of the ND5 subunit of complex I, encoded by mtDNA [37] (the rate of transcription depending in part on mtDNA copy number). Thus there is compelling evidence that mtDNA is a necessary component of the regulatory loop that controls respiration in all eukaryotic cells capable of oxidative phosphorylation. The requirement for genes to control respiration explains why prokaryotes do not usually expand up to eukaryotic size. While certain bacteria do internalise their respiration, as already noted, they only do so across a relatively restricted membrane area – no larger than the cristae area of mitochondria, or the thylakoid area of chloroplasts. But eukaryotic cells contain thousands of mitochondria, indeed hundreds of thousands in the case of large amoebae, so the surface area of bioenergetic membranes is four or five orders of magnitude greater than most prokaryotes. A simple and powerful hypothesis is that prokaryotes cannot expand up to eukaryotic size (on average five orders of magnitude larger than bacteria) without colocalizing cytoplasmic genes with their bioenergetic membranes [38, 39]. If correct, this hypothesis predicts that giant bacteria, of which there are a number of examples, must control respiration by colocalizing genes with their bioenergetic membranes. On the evidence of a few cases this does seem to be true. Epulopiscium for example, a parasite of the surgeonfish gut, is a true giant that, with a length of about 0.6 mm, dwarfs even ciliates such as

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paramecium. Epulopiscium exhibits extreme polyploidy, with as many as 600,000 copies of its full genome [40]. Other giant bacteria, such as Thiomargarita (even larger, but filled with a massive, metabolically inert vacuole) has between 6,000 and 17,000 copies of its full genome (H. Schultz-Vogt, personal communication). There is as yet no proof that polyploid genomes are necessary to control respiration in giant bacteria; but their tight association with the bioenergetic membranes is suggestive, at least. The available evidence is therefore consistent with a requirement for genes to control respiration. But the really significant point about giant bacteria, with enormous implications for the evolution of oxidative stress, is the nature of their polyploid genomes: they are invariably full genomes. This has an intractable energetic cost [39]. The significant fact about mitochondria is that they have lost the vast majority of their genes, leaving only the handful needed for the control of respiration across a wide area of membranes. Gene loss is the normal outcome of reductive evolution in intracellular bacteria and is characteristic of all endosymbionts, obligately parasitic bacteria and organelles. Because all of them started out as autonomous cells with their own cell division apparatus (and even mitochondria retain this apparatus) they are not dependent on the cell division machinery of the host cell and can replicate semiautonomously. There is inevitably competition within populations of endosymbionts for succession to the next generation, and the long-term outcome is genome reduction, because the fastest replicators tend to have the smallest, most streamlined genomes, and leave more progeny. Contrast this situation with extreme polyploidy in giant bacteria. Here, the polyploid genomes are non-autonomous copies of the host cell genome, and without some form of cytoplasmic inheritance there is no way in which they can be reduced in size over generations. Competition does not and cannot exist. Each generation passes on a fraction of its polyploid genomes, and the daughter cells are then obliged to amplify the number of copies of this genome in proportion to cell size – large cells have greater polyploidy, presumably at least in part to retain control over the wider area of bioenergetic membranes. The energetic cost is colossal. The genomic weight of 200,000 copies of the Epulopiscium genome, for example, is around 760,000 Mb of DNA, whereas the equivalent cost of 200,000 copies of mtDNA is just 6000 Mb [39]. But if the two cells could support, energetically, the same amount of DNA, then eukaryotes could support the difference, more than 750,000 Mb of DNA, in the nucleus, as genes that can be segregated at low copy number. In other words, the fact that the mtDNA genome is so small means that eukaryotic cells can support five or six orders of magnitude more DNA in the nucleus (and equally, that many more genes) than their prokaryotic counterparts. There are strong arguments to suggest that this massive increase in genomic capacity is the critical difference between eukaryotes and prokaryotes, explaining why only the eukaryotes have evolved true morphological complexity [39]. But there is a significant cost to this arrangement, which explains much about the evolution of oxidative stress [41]. If large complex cells are not possible at all without tiny mtDNA genomes, then there is a necessary interaction between mtDNA and the nuclear genes encoding mitochondrial proteins. In other words, mosaic respiratory chains, whose protein subunits are encoded by two separate genomes, are a strictly necessary feature of eukaryotic cells; eukaryotes could not exist with any other arrangement. The trouble is that the proteins encoded by the two genomes must interact with nanoscopic precision, or electron flow down respiratory chains will be blocked. Any blockage of electron flow in an aerobic world leads to a high rate of ROS leak, a collapse in energy charge (which is to say, an irreversible fall in

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ATP levels), the oxidation of membrane lipids such as cardiolipin, and the release of cytochrome c. Put another way, any failure of the two genomes to work together correctly leads directly to apoptosis. The surprising involvement of cytochrome c in apoptosis, greeted with „general stupefaction‟ when first reported in the mid 1990s [42], emerges as an explicit prediction of the hypothesis that eukaryotic cells must undergo functional selection for the compatibility of mtDNA and nuclear genes encoding adjoining respiratory chain subunits [41]. There is abundant evidence that selection for genomic coadaptation does indeed take place, ranging from the high proportion of neutral mutations and the concordance of nucleotide substitution rates in the nuclear and mitochondrial genomes, to the loss of respiratory function, fitness and fertility in introgressed populations [43]. The outstanding questions are where and how such selection takes place. What is certain is that selection for mitonuclear coadaptation necessarily involves oxidative stress.

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5. Why Oxidative Stress Is Central to the Evolution of Eukaryotes The speed of electron transfer down respiratory chains depends on the distance between redox centres, and slows down by about an order of magnitude per Ångstrom additional distance, for reasons that relate to the probability of transfer by quantum tunnelling [16]. A likely consequence of even single nucleotide mutations or polymorphisms in mtDNA would be small misalignments in subunit juxtaposition, slowing electron transfer. Slower electron transfer increases the reduction state of respiratory complexes, making them more reactive with oxygen and therefore increasing ROS leak and susceptibility to apoptosis. Thus, any mismatch between mtDNA and nuclear genes encoding respiratory-chain subunits should increase the risk of apoptosis, with manifest costs in terms of embryonic development and aging [41]. Mitonuclear mismatch is unavoidable. The only question is how much can or „should‟ be tolerated. The basic problem is that the tempo and mode of evolution of the two genomes are quite distinct. Nuclear genes evolve by sex, being recombined every generation, but the underlying mutation rate is low [38]. In contrast, mitochondrial genes evolve asexually, passing down the maternal line, but the underlying mutation rate is substantially higher, in animals and fungi at least. In yeast, the petite mutation, which deletes a large part of the mitochondrial genome, occurs at around 10,000 times the frequency of nuclear mutations [44]. Because yeast can survive by fermentation alone, selection against the petite mutants is low, hence the difference in mutation rate is observable. In animals, which usually depend on their mitochondria for oxidative phosphorylation, such mutations would necessarily be eliminated by selection. Thus, in animals, selection for mitochondrial function means that the long-term evolution rate is much lower than the mutation rate. Only the evolution rate is known with any certainty; and this is around 10-20 times faster than the evolution rate of nuclear genes [45]. Thus, again, there is a major discrepancy between the evolutionary rates of the nuclear and mitochondrial genomes. The mtDNA mutation rate is much slower in plants, for unknown reasons, but the principle stands, as there is still a large discrepancy in tempo and mode of evolution.

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These divergent rates of evolution inevitably generate mitonuclear mismatch, which is evidently eliminated by selection over time. The open questions are how much time, and how much selection? The nuclear background changes every generation, following sexual reproduction, so some kind of selective filter must be imposed each generation. Conceptually, there are only three places where selection could operate – during oocyte maturation, after fertilization, or after birth. Selection for mitochondrial function could operate at all three points, but selection for mitonuclear coadaptation cannot take place during oocyte development, because at this time the new nuclear background is not known. Selection for mitonuclear coadaptation must therefore take place during development, or after birth. Presumably, if coadaptation is poor, electron flow will slow down in the new mosaic respiratory chains: ROS leak rises, ATP levels fall and cytochrome c is released, triggering apoptosis. Compromised bioenergetics, combined with high cell loss due to apoptosis, undermines embryonic development, leading to miscarriage or mitochondrial disease [41]. The reality of such selection is unveiled by introgression between different populations of the same species, or between species. In the case of the marine copepod Tigriopus californicus, for example, introgression between neighbouring but reproductively isolated populations leads to a fall in ATP synthesis of about 40%, along with similar (c.a. 40%) reductions in fertility, developmental time and survival, even in the F2 generation: a serious, global fitness penalty [46]. Backcrossing to the maternal population completely eliminates the fitness reduction, proving that these effects are mitonuclear incompatibilities [46]. Such changes may well be responsible for the first stages of speciation in some populations, especially where the mtDNA mutation rate is high [45, 47]. Because these genes evolve faster than the nuclear average (and because nuclear genes encoding mitochondrial subunits are obliged to co-adapt, and so evolve faster themselves) the bioenergetic/apoptotic axis evolves more quickly than most nuclear genes. This drives the rate of change, and so the rate at which incompatibilities accumulate between populations, leading first to hybrid breakdown, and ultimately to speciation. It is not a long stretch to imagine that the hybrid breakdown in copepods is the first step to speciation. The mechanism of speciation, in this case, can be ascribed entirely to mitonuclear incompatibilities, which is to say the consequences of poor electron flow down respiratory chains, leading to a higher rate of ROS leak. But if poor coadaptation leads to directly to apoptosis and high rates of infertility, what constitutes „poor coadaptation‟? The answer is likely to vary among species. Species that need a high aerobic capacity, such as flighted birds or bats, could not get airborne at all if they did not have an aerobic capacity several-fold higher than even fast runners like the cheetah. On the other hand, rats do not require a high aerobic capacity, and so could presumably tolerate a poorer mitonuclear match. Put another way, there must be an adjustable threshold, above which ROS leak stimulates apoptosis and developmental failure, and below which ROS leak is tolerated, or might even be beneficial as a redox signal [45]. This leads to an eminently testable hypothesis. Birds should be highly sensitive to ROS leak from their mitochondria, whereas mammals like rats should be more tolerant of ROS leak, and also mitochondrial heteroplasmy (a mixture of different mtDNA genotypes in the same cell) [45]. This is because heteroplasmy undermines selection for mitonuclear coadaptation, as there is no means of selecting an optimal match. Given a mix of different mtDNAs, some must function against the nuclear background better than others. The overall aerobic capacity depends on the average function of mtDNA, and this average is necessarily lower than the optimum, which is, again necessarily, homoplasmic. Mitochondrial heteroplasmy is normally

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eliminated by the combination of uniparental inheritance and a mitochondrial bottleneck during oocyte development – which is to say, by the existence of two sexes, one of which (the female) bequeaths a homoplasmic population of mitochondria, and the other (the male) has his mitochondria destroyed [38]. Whether the requirement for mitonuclear coadaptation is partly responsible for the maintenance, perhaps even the origin, of two sexes, is an interesting, unanswered question; but the consequences are equally pervasive. A variable apoptotic threshold has profound implications for fertility, fecundity, adaptability, fitness, aging and age-related disease. The reason is simple. Setting the apoptotic threshold high – meaning a high tolerance of ROS-leak before apoptosis is triggered – enables high fertility and fecundity. Poor mitonuclear match is overlooked, and embryos that would fail to develop in more discerning animals develop full term. Some degree of heteroplasmy is tolerated and indeed can be beneficial, as a range of mtDNA enables greater adaptability to changing environments. However, the offspring are less fit, and more likely to suffer from mitochondrial diseases. They will have lower aerobic capacity. Worst of all, they will leak ROS from their mitochondria at a faster rate, without triggering apoptosis. The outcome is a shorter lifespan, and a greater tendency to oxidative stress and chronic inflammatory conditions linked with aging, such as diabetes, cardiovascular disease and cancer. In short, there is a trade-off between fertility, fecundity and adaptability, on the one hand, and aerobic capacity, life-span and susceptibility to age-related disease on the other [45]. The trade-off is mediated by sensitivity to oxidative stress. In birds, the apoptotic threshold is low: they are sensitive to ROS leak from mosaic respiratory chains and quickly trigger apoptosis. A poor mitonuclear match leads to slow electron flow, high ROS leak and swift apoptosis, translating into infertility and low fecundity. An intolerance of heteroplasmy means a low incidence of mitochondrial disease but also a low adaptability to changing conditions. On the positive side, birds have a high aerobic capacity, a long lifespan and low susceptibility to the chronic inflammatory conditions characteristic of old age in mammals. The difference is not trivial. Pigeons and rats have a similar body size and similar metabolic rate, even a similar foraging lifestyle, to the point that pigeons are often dismissed as flying rats. Far from it. Rats live for 3 – 4 years, pigeons for up to 35, ten times longer. Their mitochondrial ROS leak is nearly 10-fold lower [5]. While this difference makes no sense in terms of the efficiency of respiration (the proportion of ROS leak as a fraction of total oxygen consumption is too small) it makes a big difference in terms of functional selection for mitonuclear match, and it makes a big difference in terms of lifespan and healthspan [45]. “Nothing in biology makes sense except in the light of evolution”, wrote the evolutionary theorist Theodore Dobzhansky. The same applies to medicine. For decades, researchers have wrestled with oxidative stress, and found it surprisingly refractory. It is time to replace the simplistic notion that ROS leak is merely a trivial by-product of respiration, or that aging and age-related disease can be blocked with antioxidants [48]. An evolutionary perspective makes it clear why antioxidants don‟t work: they must not be allowed to interfere with sensitive ROS signals that affect everything from fertility to speciation. But equally, the evolutionary perspective reveals the costs and benefits of oxidative stress, many of them hitherto unsuspected [45]. Aging and age-related diseases are the outcomes of flexible evolutionary trade-offs, with oxidative stress at the heart of it all. But if birds can solve the aging problem, surely we can too.

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References [1] [2] [3] [4] [5]

[6] [7] [8] [9] [10] [11] [12]

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[13]

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[16]

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[19] [20]

Sies H, Oxidative stress: introductory remarks. In: Sies H, ed. Oxidative stress. London: Academic Press, 1985. Beckman KB, Ames BN. Endogenous oxidative damage of mtDNA. Mutat Res 1999;424:51-8. Boveris A, Chance B. The mitochondrial generation of hydrogen peroxide. Biochem J 1973;134:707-716. Segal AW. How neutrophils kill microbes. Ann Rev Immunol 2005;23:197-223. Barja G. Mitochondrial oxygen consumption and reactive oxygen species production are independently modulated: implications for aging studies. Rejuv Res 2007;10:21524. Orgel LE. Are you Serious, Dr. Mitchell? Nature 1999;402:17. Mitchell P. David Keilin's respiratory chain concept and its chemiosmotic consequences. Science 1979;206:1148-1159. Martin W, Russell MJ. On the origin of biochemistry at an alkaline hydrothermal vent. Phil Trans Roy Soc Lond B 2007;367:1887-1925. Lane N, Allen JF, Martin W. How did LUCA make a living? Chemiosmosis in the origin of life. BioEssays 2010;32:271-280. Lane N. Life Ascending: The ten great inventions of evolution. New York: Norton, 2009. Martin W, Baross J, Kelley D, Russell MJ. Hydrothermal vents and the origin of life. Nat Rev Microbiol 2008;6:805-14. Russell MJ, Daniel RM, Hall A. On the emergence of life via catalytic iron-sulphide membranes. Terra Nova 1993;5:343-7. Russell MJ, Daniel RM, Hall AJ, Sherringham J. A hydrothermally precipitated catalytic iron sulphide membrane as a first step toward life. J Mol Evol 1994;39:231243. Kelley DS, Karson JA, Blackman DK, et al. An off-axis hydrothermal vent field near the Mid-Atlantic Ridge at 30 degrees N. Nature 2001;412:145-149. Mielke RE, Russell MJ, Wilson PR, McGlynn S, Coleman M, Kidd R, Kanik I. Design, fabrication and test of a hydrothermal reactor for origin‐ of‐ life experiments. Astrobiology 2010; in press. Moser CM, Page CC, Dutton PL. Darwin at the molecular scale: selection and variance in electron tunnelling proteins including cytochrome c oxidase. Phil Trans R Soc B 2006;361:1295-1305. Russell MJ, Arndt NT. Geodynamic and metabolic cycles in the Hadean. Biogeosciences 2005;2:97-111. Nitschke W, Russell MJ. Hydrothermal focusing of chemical and chemiosmotic energy, supported by delivery of catalytic Fe, Ni, Mo, Co, S and Se forced life to emerge. J Mol Evol 2009;69:481-96. Allen JF. A redox switch hypothesis for the origin of two light reactions in photosynthesis. FEBS Lett 2005;579:963-968. Allen JF, Martin W. Evolutionary biology: out of thin air. Nature 2007;445:610-612.

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[21] Canfield DE, Rosing MT, Bjerrum C. Early anaerobic metabolisms. Phil Trans R Soc B 2006;361:819-1836. [22] Lane N. Oxygen: The molecule that made the world. Oxford, England: OUP, 2002. [23] Danovaro R, Dell‟ Anno A, Pusceddu A, Gambi C, Heiner I, Kristensen RM. The first metazoa living in permanently anoxic coditions. BMC Biology 2010;8:30. [24] Russell JB. The energy spilling reactions of bacteria and other organisms. J Mol Microbiol Biotechnol 2007;13:1-11. [25] Bidle KD, Falkowski PG. Cell death in planktonic, photosynthetic microorganisms. Nat Rev Microbiol 2004;2:643-655. [26] Koonin EV, Aravind L. Origin and evolution of eukaryotic apoptosis: the bacterial connection. Cell Death Differen 2002;9:394-404. [27] Cavalier-Smith T. Eukaryotes with no mitochondria. Nature 1987;326:332-333. [28] van der Giezen, M. Hydrogenosomes and mitosomes: Conservation and evolution of functions. J Eukaryot Microbiol 2009;56:221-231. [29] Martin W, Müller M. The hydrogen hypothesis for the first eukaryote. Nature 1998;392:37-41. [30] Cox CJ, Foster PG, Hirt RP, Harris SR, Embley TM. The archaebacterial origin of eukaryotes. Proc Natl Acad Sci USA 2008;105:20356-20361. [31] Pisani D, Cotton JA, McInerney JO. Supertrees disentangle the chimeric origin of eukaryotic genomes. Mol Biol Evol 2007;24:1752-1760. [32] Sagan L. On the origin of mitosing cells. J Theoret Biol 1967;14:225-274. [33] Lyons TW, Anbar AD, Severmann S, Scott C, Gill BC. Tracking euxinia in the ancient ocean: A multiproxy perspective and Proterozoic case study. Annu Rev Earth Planet Sci 2009;37:507-534. [34] Allen JF. Control of gene expression by redox potential and the requirement for chloroplast and mitochondrial genomes. J Theor Biol 1993;165:609-631. [35] Allen JF. The function of genomes in bioenergetic organelles. Phil Trans Roy Soc Lond B 2003;358:19-37. [36] Rocher C. et al. Influence of mitochondrial DNA level on cellular energy metabolism: implications for mitochondrial diseases. J Bioenerg Biomembr 2008;40:1573-1581. [37] Bai Y, Shakeley RM, Attardi G. Tight control of respiration by NADH dehydrogenase ND5 subunit gene expression in mouse mitochondria. Mol Cell Biol 2000;20:805-815. [38] Lane N. Power, Sex, Suicide: Mitochondria and the meaning of life. Oxford, England: OUP, 2005. [39] Lane N, Martin W. The energetics of genome complexity. Nature 2010;467:929-934. [40] Mendell JE, Clements KD, Choat JH, Angert ER. Extreme polyploidy in a large bacterium. Proc Natl Acad Sci USA 2008;105:6730-6734. [41] Lane N. Genomic matching and the evolution of eukaryotes. BioEssays 2011; in press. [42] Blackstone NW, Green DR. The evolution of a mechanism of cell suicide. Bioessays 1999;21:84-88. [43] Blier P, Dufresne F, Burton RS. Natural selection and the evolution of mtDNAencoded peptides: evidence for intergenomic co-adaptation. Trends Genet 2001;17:400-6.

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[44] Linnane AW, Ozawa T, Marzuki S, Tanaka M. Lancet 1989;333:642-645. [45] Lane N. On the origin of barcodes. Nature 2009;462:272-274. [46] Ellison CK, Burton RS. Interpopulation hybrid breakdown maps to the mitochondrial genome. Evolution 2008;62:631-8. [47] Gershoni M, Templeton AR, Mishmar D. Mitochondrial biogenesis as a major motive force of speciation. BioEssays 2009;31:642-650. [48] Lane N. A unifying view of ageing and disease: the double agent theory. J Theoret Biol 2003;225:531-40.

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 2

Oxygen Radicals and Related Species Ohara Augusto* and Sayuri Miyamoto Departamento de Bioquímica, Instituto de Química, Universidade de São Paulo, Caixa Postal 26077, 05513-970, São Paulo, SP, Brazil

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1. Introduction1: Oxygen and its Metabolites Imprinted the Evolution of Life As discussed in Chapter 1, around three billion years ago, life on Earth consisted of anaerobic microbes subsisting on the energy provided by cycles of electron transfer between prevalent populations of electron donors and acceptors. Potential donors were molecular hydrogen (H2), hydrogen sulfide (H2S) and methane (CH4), whose electrons were transferred to acceptors such as carbon dioxide (CO2) and, to a lesser extent, sulfate (SO42-). The biggest electron-donor pool, water (H2O), remained biologically inaccessible until the first photosynthetic organisms evolved between ~3.2 and 2.4 billion years ago. Employing the energy of the sun and H2O as the reductant to fuel metabolism, these ancient organisms produced molecular oxygen (O2) as a waste product. Over a relatively short period of geologic time, around 100 million years, O2 atmospheric levels increased, indicating that the energy-transducing process of photosynthesis was replacing the old metabolic networks. As a consequence, the life forms that evolved in anaerobic conditions had to adapt to O2, hide or become extinct. With O2 overwhelming the Earth‟s reducing atmosphere, these primitive microbes became restricted to hypoxic or anaerobic environments. However, some of them were able to escape and adapt to the oxidizing atmosphere. They either modified parts of their metabolism, or formed symbiotic associations that permitted the coupling of nutrient oxidation to reduction of O2 back to H2O. This respiratory pathway was far more efficient in extracting energy from nutrients, but it came with costs. The triplet ground state of O2 is * 1

Email: [email protected] Due to space constraints, we cited only a small fraction of the original investigations that contributed to the current understanding of the multiple roles of radicals/oxidants in Biology. Whenever possible, we cited reviews that referred to the original papers.

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prone to one-electron transfers, which yield species that are toxic to life (see section 6.3). This pushed the development of mechanisms to protect the genetic investment in the metabolic pathway that drive the “H2O-H2O” cycle upon which most life on Earth would come to depend. The protective pathways evolved further, driving the evolution of single-cell eukaryotes, from which all current plants and animals on Earth descend [1, 2]. Adaptation to O2, however, went beyond the creation of redox energy couples that allowed complex life forms to evolve. As a recent meta-metabolomic analysis demonstrated, it also pushed an evolutionary explosion of random, alternative and novel metabolic networks yielding a wide variety of gene products that increased fitness of the organisms. Although we have a long way to go to understand how life evolved, it is clear that O2 and its derived metabolites imprinted the evolution of complex life forms [1, 2]. Thus, the participation of oxygen-derived metabolites in cell regulatory and signaling pathways could be anticipated [2]. Nevertheless, until recently, oxygen-derived metabolites were mostly considered for their involvement in cell damaging mechanisms [3, 4]. In this Chapter, the chemical properties of O2 that cause its propensity to produce free radicals and oxidants as metabolites will be presented. These species, known by the general term reactive oxygen species (ROS) will be historically contextualized. The discoveries pointing to their roles as mediators of physiological and pathophysiological circuits will be summarized. Finally, the reactivity of specific ROS towards biomolecules will be presented in the context of health and disease - topics that will be extended in Volume III of this book.

Figure 1. Electronic distribution of the electrons of two oxygen atoms (8O) in atomic orbitals, which combine in molecular orbitals to form molecular oxygen (O2) (left side). O2 contains two unpaired electrons. This is because in distributing its electrons in molecular orbitals, the last two electrons have two molecular orbitals of the same energy to occupy (2 *) and each electron occupies one (Hund´s rule). Because the presence of two electrons in antibonding orbitals (2 *) energetically cancels out one of the bonding orbitals (2 ), the two oxygen atoms in O2 are bound by two covalent bonds (3 bond-occupied orbitals minus 2 antibound semioccupied orbitals). B. Electronic distribution of the electrons of one nitrogen atom (7N) and one oxygen atom (8O) in atomic orbitals which combine in molecular orbitals to form nitric oxide (NO●) (right side). Distribution of the 17 electrons of NO● in molecular orbitals leaves an unpaired electron in the 2 * orbital making it a free radical in the ground state. Nitrogen and oxygen in NO● are bound by 2.5 covalent bonds (3 bond-occupied orbitals minus 0.5 antibond orbitals).

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2. Molecular Oxygen, a Sluggish Oxidant 3. Producer of Reactive Species Long before the formation of molecules and ions was conceptualized by quantum mechanics, the discoverers of molecular oxygen in the late 18th century - Priestley, Scheele and Lavoisier - reported its beneficial and toxic effects on living organisms. These opposing effects result from the properties of molecular oxygen, which is formed by the combination of two oxygen atoms in covalent bonds (O2). The characteristics of a covalent bond, such as strength, length and direction, depend on the occupied molecular orbitals. O2 is bound by two covalent bonds (Figure 1), and requires 402 kJ/mol to break into two oxygen atoms (O). This is roughly the amount of energy required to bring 1 liter of H2O to boiling, indicating that the bond strength between the two O atoms in O2 is strong. In other words, O2 is a considerably stable molecule, as attested by our daily experience. Most covalent bonded molecules are equally stable, possessing energy bonds from 150 to 950 kJ/mol. However, O2 has its peculiarities. Although stable, a spark triggers its reaction with fuels (combustion) liberating energy to move our cars. Likewise, the oxidation of the foods we eat (respiration) produces energy to sustain our lives. Indeed, most reactions of O2 are slow but, once initiated or catalyzed, liberate a considerable amount of energy. In other words, reactions of molecular oxygen are favored by thermodynamics but not by kinetics. This is because O2 has two unpaired electrons in the ground state, which makes it a triplet molecule (Figure 1A). In contrast, most organic molecules possess all electrons paired, being singlet in the ground state. To react with them, O2 has to receive a pair of electrons, but this requires spin inversion, an event that is prohibited by the spin conservation rule (Figure 2). According to its structure, O2 tends to receive electrons by one-electron steps, reacting rapidly only with species capable of one-electron transfer. Examples of such species are biomolecules that produce stable free radicals, such as flavin enzymes and coenzymes, and species that contain unpaired electrons, such as other free radicals and transition metal ions (Figure 2). Thus, it is not surprising that iron artifacts are rapidly turned into scrap and that most enzymes that catalyze biological oxidations contain transition metal ions and/or flavin coenzymes in their active sites. O2, free radicals and transition metal ions are closely related. In fact, O2 and transition metal ions are also free radicals because they contain unpaired electrons. Nevertheless, „free radical‟ is more frequently defined as a species (molecule, cation or anion) that contains one unpaired electron. Only in these cases is the unpaired electron denoted by a superscript dot to the right preceding any charge in the radical formula [5]. The term free radical is historical because organic radicals, which are used to refer to the chemical groups in a molecule, are bound as opposed to free. Currently, both terms free radical and radical are used in the literature and understood by their contexts. A classical free radical in the ground state is nitric oxide (NO ) (Figure 1B), whose role in signal transduction pathways was established in the last decade of the 20th century [3, 4, 6].

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Figure 2. Schematic representation of the main reactions of O2. Due to its unpaired electrons, O2 does not react with most organic molecules because they have paired electrons. Such reactions are extremely slow because they would require spin inversion which is prohibited by the spin conservation rule. O2 reacts rapidly with molecules that produce stable radicals by one-electron transfer (BM●+) or have unpaired electrons, such as free radicals (R●) and transition metal ions (Mn+). If O2 receives energy (from light, excited molecules, etc.), it is excited to singlet oxygens (1 gO2 or 1 g+O2) that are more reactive towards biomolecules because there is no spin restriction.

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3. Free Radicals in Biology: An Historical Account In the first half of the 20th century, organic radicals were recognized as intermediates of chemical reactions, and many of their properties were studied and explored in polymerization processes. Most investigators considered radicals irrelevant to biology, except for some pioneers. For instance, the known propensity of O2 to react by one-electron steps (Figure 2) led some investigators, such as Otto Warburg and Leonor Michaelis, to propose that during respiration O2 would produce one-electron intermediates, such as superoxide (O2 -) and the hydroxyl radical (HO ) (Reaction 1).

O2 + 4H+ + 4e e O2

e /2H+ O2∙

2 H2O

e /H+ H2O e /H+ H2O2

HO∙

H2O

(Reaction 1) Evidence that O2 produced radicals in animals was first provided by Rebeca Gerschman and co-workers in 1954 while studying the toxic effects of high O2 pressures and X-ray irradiation in mice [7]. In the resulting paper, they concluded: “it would appear that irradiation and O2 poisoning produce some of their lethal effects through at least one common mechanism, possibly that of the formation of free radicals”. In addition, they showed that

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protectors against irradiation, such as glutathione and ethanol, protected against oxygen toxicity, or as currently defined, they acted as antioxidants. Another pioneer was Denham Harman, who proposed in 1954 that aging was a consequence of free radical attack on biomolecules (see Vol. II, Chapter 23). These views remained largely ignored up to 1970, while the properties of free radicals were extensively studied by radiation chemists. The employment of biochemical approaches to explore fundamental biological questions, such as the molecular basis of life, provided a turning point. In this background, McCord and Fridovich discovered the enzyme superoxide dismutase (SOD) in 1969 [8] (see also Vol. II, Chapter 5). They showed for the first time that mammals possess an enzyme whose function is to catalyze the dismutation of O2●- (k= 1.6 x 109 M-1 s-1), which also occurs spontaneously at a slower rate (k= 5 x 105 M-1 s-1) (Reaction 2). Dismutation is a typical reaction of most free radicals; two molecules of the same radical (such as O2●-) react in an electron transfer process to produce oxidized (O2) and reduced (H2O2) products.

O2∙ + O2∙ + 2 H+

H2O2 + O2

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(Reaction 2) The fact that evolution preserved an enzyme whose function is to dismutate a free radical indicated that radicals were constantly produced during normal metabolic processes. This fundamental discovery inaugurated a new area of research, “free radicals in biology”, which continues to expand. The discovery of SOD provided evidence that cells generate O2●-. Soon thereafter, it became clear that during biological processes cells also produce additional related intermediates, generically termed reactive oxygen species (ROS), such as H2O2, HO● and hypochlorous acid (HOCl). ROS were shown to be produced during mitochondrial respiration, phagocyte-mediated killing of pathogens, and xenobiotic metabolism. These investigations brought new concepts into pathophysiology. One was the importance of reactions catalyzed by transition metal ions (Fenton chemistry; Chapter 5) to produce HO●, an extremely potent oxidant, from the less reactive O2●- (Reactions 3 and 4).

Fe 3+ + O2∙ H2O2 + Fe2+

Fe2+ + O2 Fe3+ + HO∙ + HO

(Reactions 3 and 4) Another was the notion that if not eliminated by the antioxidant defenses, ROS would attack DNA, lipids and proteins (Chapters 6-9) causing cell and tissue injury. The most influential one was the concept of oxidative stress defined as an imbalance between free radicals/oxidants and antioxidants in favor of the former [9]. As a consequence, radicals and oxidants became associated with most human diseases and many intervention studies were designed to examine the effects of antioxidant vitamins on diseases, particularly cardiovascular and neurological.

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By the last decade of the 20th century, however, most data collected from intervention studies with antioxidants were inconclusive, suggesting that the classical view of oxidative stress required revision [4, 6]. This was reinforced by the discovery of NO● (Figure 1), a gaseous free radical, as a major autocrine and paracrine mediator of vascular relaxation, immune regulation and many other physiological effects. In mammals, NO● is mainly produced from arginine oxidation, catalyzed by a family of enzymes, the nitric oxide synthases (NOS) (Reaction 5) [10, 11].

Arginine + NADPH + O2

NOS

NO∙ + Citruline + NADP+ + H2O

(Reaction 5)

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Soon thereafter, it was demonstrated that O2●- reacts rapidly with NO● (Reaction 6) and not only regulates NO● bioavailability but also leads to potent oxidants such as peroxynitrite (ONOO-/ONOOH)†, HO●, nitrogen dioxide (NO2●) and carbonate radical (CO3●-) (Reactions 7 and 8) [12-14] (see also Chapter 3).

Figure 3. Schematic representation of the roles of radicals and oxidants as mediators of physiological and pathophysiological circuits. Present evidence indicates that free radicals and oxidants are constantly produced in vivo from exogenous and endogenous sources and are, directly or indirectly, derived from O 2. The main endogenous sources of free radicals/oxidants are: the mitochondrial electron transport chain and enzymatic reactions catalyzed by nitric oxide synthases (NOS), NADPH oxidases (NOX) (from phagocytes and other cell types), xanthine oxidase (XO) and hemeperoxidase enzymes, such as myeloperoxidase (MPO). Organisms evolved enzymatic antioxidant defenses and the capability to use antioxidants from the diet to control the physiological levels of free radicals/oxidants. High levels of some oxidants may cause dysfunction because they oxidize biomolecules that lead to cell and tissue injury if not repaired by the evolved repair systems. Low levels of certain free radicals/oxidants also compromise physiological functions that evolved to depend on them. On the other hand, transient and small increases of some oxidants trigger redox-sensitive signaling pathways. †

The term peroxynitrite refers to the sum of peroxynitrite anion (ONOO-, oxoperoxonitrate (-1)) and peroxynitrous acid (ONOOH, hydrogen oxoperoxonitrate) unless otherwise specified. Other abbreviations are defined in the text.

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k= 1.9 x 109 M-1 s-1

O2∙ + NO∙

k= 0,17 s-1 0.7 NO + 0.7 H+ 3

pKa= 6.4

ONOO + H+

ONOO

ONOOH

k= 2.6 x 104 M-1 s-1

ONOO + CO2

[ONOOCO2 ]

0.3 HO∙ + 0.3 NO2∙

0.65 NO3 + 0.65 CO2 0.35 NO2∙ + 0.35 CO3∙

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(Reactions 6-8) Furthermore, in the same period, several lines of evidence indicated that H2O2 can act as a second messenger for receptor agonists, such as growth factor and hormones, prompting the development of the concept of redox signaling. This involves cellular signal transduction networks in which the integrative element is a series of interconnected electron transfer reactions. By exerting second messenger effects, oxidants can regulate major cellular pathways [15-17]. Such function is likely rooted in the oxygen-dependent evolution of complex life forms. Our current knowledge about redox signaling mechanisms is still in its infancy. For decades, free radical research focused on understanding the formation of reactive species in vivo, and elucidating how proteins, lipids and DNA are damaged by them, resulting in cellular injury and disease. The complete picture is more complex, though. Exceeding levels of reactive species may cause dysfunction, but low levels of certain oxidants also compromise physiological functions that evolved to depend on them, such as microbicidal activity, proliferative responses and vasodilation. Moreover, transient and small increases in some oxidants trigger redox-sensitive signaling pathways. Thus, radicals and oxidants are presently considered to control signaling circuits involved in physiological and pathological responses (Figure 3) [4, 6]. Unraveling these interrelated processes requires a better understanding of cellular oxidative mechanisms, including the identification of the involved oxidants, the pathways regulating their generation, and their molecular targets. Further advances in the field will depend on an interdisciplinary effort that combines rigorous chemical thinking and tools with relevant biological data and insights [16, 17].

4. Oxygen Radicals and Related Species: General Aspects of the Reactivity of Oneand Two-electron Oxidants Although discrimination between different biologically relevant oxidants became critical, the difficulties in detecting these short-lived species in biological media stimulate the use of general terms, such as ROS, in the literature. As previously noted by Christine Winterbourn [16, 17], terms such as ROS and antioxidants are appropriate to refer to general classes of compounds but are counterproductive for understanding mechanisms. This is because not all antioxidants act by the same mechanism, and the species encompassed by ROS have widely different reactivities. The same is true for another frequently used general term, RNS (reactive

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nitrogen species). RNS comprise reactive metabolites derived from NO●, such as peroxynitrite and NO2● (Chapter 3). However, RNS and ROS are closely interrelated (see, for instance, Reactions 6-8), and only one general term may be more appropriate. Indeed, radicals typically react rapidly with other species containing unpaired electrons (Figure 2), making it often difficult to establish which oxidant is involved in a particular physiological process. To make progress in this direction, it is necessary to recognize the widely different characteristics of the species encompassed by the term ROS (and RNS). Some are radicals and undergo one-electron reactions, whereas others are non-radicals and promote two-electron oxidations. Some are strong and others weak oxidants, and the second-order rate constant of their reactions with biomolecules vary considerably [16, 17]. For instance, the second order rate constant of some physiologically relevant oxidants with glutathione (GSH) varies from undetectable to diffusion-controlled (k ~ 109-1010 M-1 s-1 in aqueous media) (Table 1). The tripeptide GSH ( -L-glutamyl-L-cysteinylglycine), which is present at millimolar concentrations in most cell types, is a major intracellular reductant because of its cysteine thiol group (-SH) (Vol. II, Chapter 1). In this book, oxygen radicals and related species (this Chapter), nitrogen radicals and related species (Chapter 3) and sulfur radicals and related species (Chapter 4) are discussed separately. Nevertheless, it is useful to stress some general aspects concerning the reactivity of biologically relevant oxidants. The oxidizing strength of a radical is given by its oneelectron reduction potential (Eo), with higher values corresponding to more potent oxidants. This is usually reflected in the rates at which radicals react because of the low activation energy of radical reactions (Table 1). Thus, it is not surprising that HO● reacts with most biomolecules with second-order rate constants close to the diffusion limit. That is, the reaction occurs as fast as the reagents encounter each other. As one-electron oxidants, radicals favor radical chain reactions as occurs during lipid peroxidation (Chapter 7). A remarkable exception is NO●, which rapidly interacts with other radicals interrupting radical chain reactions (Reaction 9; see also Chapter 8). k~ 109 M-1 s-1

NO∙ + ROO∙ NOOOR (Reaction 9) Although low NO● levels inhibit lipid peroxidation, the radical can produce potent oxidants by reacting with O2●- (Reaction 5) and O2 (Reaction 10). k= 2

2

NO∙

+ O2

106 M-2 s-1

2 NO2∙ (Reaction 10)

In the case of two-electron oxidants, their reduction potential also determines their oxidizing strength; however, kinetics determines their reactivity because of the high activation energy involved in these oxidations. For instance, H2O2 has a higher reduction potential than peroxynitrous acid (ONOOH) but reactions of H2O2 have higher activation energies and slower rates (Table 1).

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Oxygen Radicals and Related Species Table 1. Relative reactivity of selected radical and non-radical oxidants Oxidant

Reduction potential (E ‟, V)

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Radicals (one electron)a -0.80 NO●/3NO 0.92 RS●/RS (Cys) ● + 0.94 O2 , 2H /H2O2 1.06 HO2● , H+/H2O2 ROO●, H+/ROOH 1.00 ● 1.04 NO2 /NO2 RO●, H+/ROH 1.60 1.78 CO3● , H+/HCO3 ● + 1.80 O3 , 2H /H2O, O2 HO●, H+/H2O 2.31 Non-radicals (two electron)b 1.40 ONOOH, H+/NO2 , H2O 1.28 HOCl, H+/Cl , H2O H2O2, 2H+/2 H2O 1.77 a Data for one-electron reduction potential collected from [74]. b Data for two-electron reduction potential collected from [17, 36]. c Second-order rate constants for GSH collected from [17, 22]. n.d., not determined.

kGSH (M-1 s-1)c

non detectable 8.0 x 108 10 to 103 n.d. n.d. 3.0 x 107 n.d. 4.6 x 107 7.0 x 107 1.0 x 1010 6.6 x 102 3.0 x 107 0.9

The kinetics of the reactions of radicals and oxidants with biotargets contribute to the understanding of oxidant actions under physiological conditions. The relevance of the target depends on its local concentration ([BM]) and on the second-order rate constant of its reaction with the oxidant (k). Indeed, the product (k x [BM]= k´(s-1)) allows the ranking of the targets of an oxidant in homogenous media. For instance, the main target of intracellular peroxynitrite is likely to be CO2 because it reacts rapidly with peroxynitrite (Reaction 8) and has a high intracellular concentration (~1.3 mM) due to the bicarbonate buffer. The value of k´ for CO2 (k´= 2.6 x 104 x 1.3 x10-3 ~ 34 s-1) is unmatched by that of intracellular GSH (~5 mM) (k´= 6.6 x102 x 5 x10-3 ~ 3.3 s-1) (Table 1) and other possible targets, except for some heme and thiol proteins. Among the latter, peroxiredoxins (Prx), which are abundant and react rapidly with peroxynitrite (k= 105 -107 M-1 s-1) (Reaction 11), deserve special attention [1620] (see also Vol. II, Chapter 10). k~ 105-107 M-1 s-1

ONOOH + Prx-S PrxSOH + NO2 (Reaction 11) At 5 µM concentration, a Prx whose k= 1 x107 M-1 s-1 (k´= 50 s-1) will compete almost equally with CO2 for peroxynitrite. It is noteworthy that Prx also react rapidly with H2O2 (k= 105 -107 M-1 s-1). The high second-order rate constant of the reactions of these enzymes with peroxides is one of the reasons why they are being examined as mediators in redox signaling

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mechanisms [16-20]. It is also important to point out that the k´value permits an estimation of the diffusion distance of selected oxidants under specific circumstances [21]. As an example, Figure 4 shows the putative diffusion distance of peroxynitrite in the presence of physiological concentrations of GSH, CO2 and a Prx. It becomes evident that if the only peroxynitrite target in a cell were GSH (5 mM), the oxidant would diffuse away from one cell to others and thereby oxidize distant targets by two-electron mechanisms. In the presence of both CO2 and GSH, the radicals resulting from peroxynitrite would diffuse for only a few hundred nanometers, and GSH would be oxidized by one-electron mechanisms to the glutathionyl radical (GS●) (Figure 4). These examples show the importance of considering kinetics in planning and interpreting experiments related to oxidant action under physiological conditions. There are databases (NIST Standard Reference DataBase [22]) and several reviews articles that collect second-order rate constants for reactions of radicals and oxidants (see, for instance, [13, 14, 16-18]). It is always useful to consult them, although other factors that affect reactions in physiological media, such as compartmentalization and media heterogeneity, remain more difficult to assess.

Figure 4. Estimated diffusion distance of peroxynitrite in the presence of physiological concentrations of GSH (5 mM), CO2 (1.3 mM) and a peroxiredoxin (Prx) (5µM) in a generic tissue composed of cells with a 20 µm diameter. The scheme shows peroxynitrite migration from the center of the circle over different distances in the presence of the specified targets. Diffusion distances (l) for a tenfold decrease in the oxidant concentration are represented as the radius (µm) of a circle and were calculated from the expression l=2.3(D/k[BM])1/2, where D is the oxidant diffusion coefficient, k is the second-order rate constant value and [BM] is the concentration of the target [17, 21]. D values for peroxynitrite are unknown and a value of 1000 µm2/s was used for the calculation. For comparison, the diffusion distances of the radicals produced from peroxynitrite reaction with CO2, nitrogen dioxide (NO2 ) (188 nm) and carbonate (CO3 ) radical (152 nm) in the presence of GSH, are also shown.

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5. Chemistry of Biologically Relevant Oxygen-Derived Radicals The biologically relevant oxygen-derived radicals include O2●- and its conjugated acid (HO2●), HO●, CO3●-, peroxyl (ROO●) and alkoxyl (RO●) radicals. The sources, properties and main reactions of these species in physiological environments are discussed below.

5.1. Superoxide (O2●-) and Hydroperoxyl (HO2●-) Radical Superoxide (O2●-) is formed by the one-electron reduction of O2 (Reaction 1). In biological systems it is produced by enzymatic reactions catalyzed by oxidases, such as NADPH oxidases (NOX) and xanthine oxidase (XO), and non-enzymatically by redox active compounds, such as the semi-ubiquinones of the mitochondrial electron transport chain. O2● can act as oxidant or reductant, and the dismutation reaction is an example of this double action (Reaction 2). The second-order rate constant of the dismutation reaction is higher at acidic pH due to the increase in the concentration of the hydroperoxyl/perhydroxyl radical (HO2●) (pKa = 4.8) in equilibrium with O2●- (Reaction 12). The determined second order rate constant value for the spontaneous dismutation varies from 102 M-1 s-1 at pH 11 to 5 x 105 M-1 s-1 at pH 7. As noted above, the dismutation reaction catalyzed by SOD is much faster (k= 1.6 x 109 M-1 s-1).

O2∙ + H+

HO2∙

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(Reaction 12) O2●- reactivity towards a variety of organic and inorganic targets has been studied. Despite its moderately high reduction potential (0.94 V) (Table 1), its reactivity with nonradical targets is limited [23]. However, O2●-may induce harmful effects by reacting with radicals, such as NO● (Reaction 6), and species containing transition metal ions. For instance, O2●- is highly reactive towards iron-sulfur ([Fe-S]) clusters [24] (see also Vol. II, Chapter 19), with the reaction occurring at near diffusion-limited rates (Reaction 13). O2●- causes oneelectron oxidation of the [4Fe-4S] clusters to form H2O2 and an unstable intermediate that decomposes losing iron (II). Relevantly, this free iron ion can react with H2O2 and catalyze the production of HO● by Fenton chemistry (Reactions 3 and 4). k > 109 M-1 s-1

[4Fe-4S]2+ + O2∙ + 2 H+

H2O2 + [4Fe-4S]3+ (Reaction 13)

[3Fe-4S]+ + Fe2

Radical-radical reactions of O2●- typically occur at near diffusion-controlled rates [25]. Among them, the reaction of O2●- with NO∙ has received special attention due to the generation of peroxynitrite (Reaction 6), a strong biological oxidant (Chapter 3). Reactions of O2●- with radicals formed on aromatic amino acids, such as the tyrosyl radical (Tyr●), have

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also been studied. The second-order rate constant of the reaction with Tyr● is about 3-fold higher than that of Tyr● dimerization to dityrosine [25]. Upon reaction with O2●- the Tyr● radical can be repaired back to Tyr or be transformed into Tyr hydroperoxides depending on the position of the Tyr residue in the polypeptide chain. Reactions of O2●- with biologically important thiol compounds have been investigated [26]. Literature data on the second-order rate constant for these reactions vary from 10 to 105 M-1 s-1. This variation has been attributed to the use of inappropriate assays and the complexity of the chain reactions involving sulfinyl and thiyl radicals [27]. According to Winterbourn and colleagues, the best estimates are in the range of 30 to 103 M-1 s-1. These values indicate that biothiols do not react rapidly enough with O2●- to compete with SOD. Therefore, proteins containing [Fe-S] clusters are likely to be O2●- sensors in redox signaling (Reaction 13). In agreement, bacteria rely on the SoxR transcription factor, which contains [Fe-S] clusters, to upregulate resistance genes in response to sublethal O2●- levels.

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5.2. Hydroxyl Radical (HO●) HO● is the strongest oxidant produced in biological systems (E ‟ =2.31 V) (Table 1). It reacts very rapidly and indiscriminately with most biological targets present at its site of formation (see, for comparison, Figure 4). Indeed, the rate constants for its reactions with most of biomolecules, including, lipids, proteins, carbohydrates and DNA are very close to the diffusion-controlled limit. HO● can be generated in vivo by four major processes: ionizing-radiation (e.g. UV, X-rays, -rays); transition metal ion-catalyzed reactions (Reaction 4); proton-catalyzed decomposition of peroxynitrite (Reaction 6); and decomposition of ozone (O3) (see section 6.5). Irradiation with high-energy radiations generates HO● by homolytic fission of water molecules (Reaction 14). HO● produced by this process is thought to be responsible for DNA damage and tumor development (e.g. skin cancer), while irradiation can also be employed for targeted killing of tumor cells (e.g. radiotherapy).

H2O HO∙ + H∙ (Reaction 14) Reactions of the HO● can be classified into three main types: hydrogen abstraction, addition and electron transfer reactions. HO● is extremely fast in abstracting hydrogen atoms from organic compounds, especially from those that are weakly bound. The reaction produces H2O and a carbon-centered radical (RC●), which, in the presence of O2 generates peroxyl radicals (ROO●), such as in the case of carbon-centered lipid radicals. In the absence of O2, a covalent bond can be formed between two RC● producing a crosslink. An example is the reaction between two Tyr● to produce dityrosine. Reactions of HO● with aromatic compounds usually involve the addition of the radical to produce hydroxylated radical adducts. An important example is the addition of HO● to the guanine moiety of DNA/RNA to produce 8hydroxyguanine and 2,6-diamino-4-hydroxy-5-formamidopyrimidine (Chapter 9). Reactions of HO● with anions produce radicals by electron transfer mechanisms. For instance, the

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abundant Cl- and the NO●-metabolite, NO2-, are oxidized to chlorine (Cl●) and NO2●, respectively (Reactions 15 and 16). k= 4.3

Cl

+

HO∙

NO2

+

HO∙

109 M-1 s-1

Cl∙ +

HO

1010 M-1 s-1

k= 1

NO2∙

+

HO

(Reaction 15 and 16)

5.3. Carbonate Radical (CO3●-) Although less oxidizing than HO●, CO3●- (Eo´ =1.78 V) (Table 1) is a very strong oneelectron oxidant. Relatively recent data demonstrated that CO3●- is an electrophilic oxygencentered radical and a strong acid (pKa 1.0 V) (Table 1). They can be generated from organic hydroperoxide (ROOH) decomposition induced by heat or radiation Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

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and by ROOH reaction with transition metal ions and other oxidants capable of abstracting hydrogen (Reactions 18 and 19). ROO● are also important intermediates in processes involving carbon-centered radicals, which react rapidly with O2 (k>109 M-1 s-1). In addition, biomolecule-derived ROO● and RO● can be generated from the oxidation of lipids, proteins and nucleic acids.

ROOH + Fe2+

RO∙ + HO + Fe3+

ROOH + Fe3+

ROO∙ + H+ + Fe2+

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(Reaction 18 and 19) The reactivity of ROO● and RO● is influenced by the substituents at the -carbon. An electron-withdrawing group increases the reactivity (for instance, chloroperoxyl radical, CCl3OO●), whereas electron-donating groups decreases it (for instance, phenoxyl radicals). Aromatic ROO● and RO●∙ tend to be less reactive because of unpaired electron delocalization. The reactions of ROO● and RO● with biomolecules often involve hydrogen-abstraction, which is facilitated in compounds containing weakly bound hydrogens. This is the case for lipids, thiols, and several chain-breaking antioxidants. Lipids are particularly susceptible to hydrogen abstraction and this reaction is the rate-limiting step in the propagation of lipid peroxidation chain reactions (Chapter 7). The second-order rate constant value of ROO●mediated hydrogen abstraction from lipids is low (k < 102 M-1 s-1) [29, 30] and increases with the number of allylic or double-allylic hydrogens. For unsaturated fatty acids the secondorder rate constant value decreases in the sequence 22:6>20:5>20:4>18:2>18:1. Notably, the second-order rate constant values of RO● reactions (k 106-107 M-1 s-1) [29] are about 4-5 orders of magnitude higher than those of ROO●. Both ROO● and RO● can undergo rapid monomolecular rearrangements or fragmentations that compete with hydrogen abstraction reactions. ROO● formed on aromatic rings and those with -carbon linked to hydroxy or amino groups can decompose to liberate O2●- or HO2●. This type of reaction has been reported for amino acids, such as lysine. ROO●∙ can also react with another ROO●∙ by the Russell mechanism, generating a ketone, an alcohol and singlet molecular oxygen (1O2) (see section 6.3) [31].

6. Chemistry of Biologically Relevant Non-radical Oxygen Species Biologically relevant two-electron oxidants derived from oxygen include H2O2, HOCl and related species, 1O2, biomolecule-derived hydroperoxides and ozone (O3). These species are discussed below with regard to main properties and reactions in physiological environments.

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6.1. Hydrogen Peroxide (H2O2) Biological production of H2O2 can occur by chemical and photochemical processes and by enzymatic reactions catalyzed by several oxidases, such as NOX, XO and monoamine oxidase (MAO). In addition, H2O2 is produced continuously from O2●- dismutation, either spontaneous or catalyzed by SOD (Reaction 2). In physiological environments, H2O2 can be rapidly inactivated to H2O by seleno-, heme- and thiol- peroxidases. For instance, the secondorder rate constant value of the reaction of H2O2 with glutathione peroxidases (GPx), catalases and Prx are ~108 M-1 s-1, ~106 M-1s-1 and 105 -107 M-1s-1, respectively. Thus, steadystate concentrations of H2O2 in cells and tissues are low, being estimated to be around 10-7 to 10-8 M. H2O2 is a powerful two electron-oxidant (E = 1.77 V, pKa 11.6) (Table 1). However, its reactivity toward most of biological molecules is low because of the high activation energy of these oxidations. Most damaging effects of H2O2 in vivo are considered to be mediated either by transition metal ions or enzymes, such as heme-peroxidases. These processes generate secondary species, which are more reactive and include radicals, such as HO● and NO2●, and non-radical species, such as HOCl and related species. Reaction of H2O2 with reduced transition metal ions, such as copper (I) and iron (II), leads to the generation of HO● (Reaction 4). This reaction has been studied for more than a century (Chapter 5). The secondorder rate constant values of Fenton reactions are dependent on the metal-ligand and on the medium pH, ranging from 102 to 104 M-1 s-1 for iron ions. In addition to being substrate for several heme-peroxidases, H2O2 can be consumed by other hemoproteins, such as hemoglobin, myoglobin and cytochrome c, in processes that oxidize the proteins. In phagocytic cells, myeloperoxidase (MPO) uses H2O2 mainly for the production of HOCl (Reactions 20 and 21). MPO has also been reported to use H2O2 to oxidize NO2- to NO2● [32] and Tyr to Tyr● [33]. Thus, MPO can be an efficient mediator of protein nitration, particularly at acidic pH [34]. MPO is released during phagocytosis and is thought to play an important role in microbial killing [35]. However, excessive MPOmediated production of HOCl, NO2● and other oxidants can cause host tissue injury, contributing to the development of several diseases [36, 37].

MPO + H2O2 MPO + Cl

k1

k -1 k2

MPO-I + H2O MPO + HOCl

(Reaction 20 and 21) Among the physiological reactions of H2O2, the oxidation of biothiols is receiving increasing attention in the literature [15, 16]. The second-order rate constant values of the oxidation of low molecular weight thiols (RSH) by H2O2 are relatively low (1-5 x 102 M-1 s-1). The value increases for thiols with low pKa [26], indicating that the effective substrate is the thiolate form (RS-). In fact, it is proposed that a nucleophilic attack of RS- on peroxide oxygen occurs to produce sulfenic acid (RSOH) (Reaction 22), which subsequently reacts with a second thiol to produce disulfide (RSSR) (Reaction 23).

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Ohara Augusto and Sayuri Miyamoto

H2O2 + RS

RSOH + H2O

RSOH + RSH

RSSR + H2O

(Reaction 22 and 23) It is important to emphasize that some classes of proteins possess Cys residues whose pKa values are in the range of ~4.0 to ~6.5, which are much lower than that of free Cys (pKa=8.4) or GSH (pKa=9.2). Examples of these proteins are the Prx, which react extremely rapid with H2O2 (k~105-107 M-1 s-1). Currently, it is accepted that Cys residues with low pKa are only one of the factors contributing to the high reactivity of Prx towards peroxides [18]. It will be important to unravel Prx catalysis at the molecular level because these enzymes are likely to participate in H2O2-mediated signaling cascades [16-20] (see also Vol. II, Chapter 10).

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6.2. Hypochlorous acid (HOCl) and Related Species HOCl and related species (HOX, X= Cl, Br, I and SCN) are moderately strong twoelectron oxidants (E ‟HOCl/Cl =1.28 V, HOBr/Br =1.13 V, HOSCN/SCN =0.56V) generated during inflammatory processes (Vol. II, Chapter 21). They are primarily produced from the reaction of H2O2 with halide and pseudo-halide ions (Cl , Br , I and SCN ) catalyzed by MPO and eosinophil peroxidase [36, 37]. In terms of plasma concentration, the most abundant halide ion is Cl (100-140 mM) followed by Br (20-100 µM), SCN (20-120 µM) and I (0.1-0.6 µM) [38]. Thus, it is believed that HOCl is the major hypohalous acid produced during phagocytosis [37, 39]. However, the second-order rate constant value for the halogenation reaction is about 10-20 fold higher for SCN and Br compared to Cl and significant amounts of HOSCN and HOBr can also be produced. At physiological pH, HOCl is in equilibrium with its conjugate base, hypochlorite (OCl-, pKa 7.59 at 25 C) [36] (Reaction 24), and both forms appear to be responsible for oxidation and/or halogenation reactions. In acidic conditions, HOCl can be in equilibrium with molecular chlorine (Cl2, pKa 3.3) [40] (Reaction 25). In vitro studies suggest that Cl2 might be the agent that mediates formation of chlorinated products during phagocytosis [40].

HOCl

pKa = 7.4

HOCl + Cl + H+

H+ + OCl

pKa = 3.3

Cl2 + H2O

(Reaction 24 and 25) HOCl is reactive towards several biomolecules. Amino (RNH2) and thiol (RSH) groups of amino acids and peptides are among the most important targets (Table 1). Oxidation of these groups by HOCl yields unstable chloramines (RNHCl) and sulfenyl chlorides (RSCl), respectively [41-43]. Both intermediates induce further reactions that lead to an increased oxidative damage to biomolecules [36]. HOCl is also reactive towards aromatic rings in

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35

amino acids and nucleobases. Reaction of HOCl with Tyr yields 3-chlorotyrosine and 3,5dichlorotyrosine [44, 45]. With lipids, HOCl adds across carbon-carbon double bonds in fatty acids and cholesterol yielding chlorohydrins [46]. HOCl also reacts with H2O2/ROOH. The reaction with H2O2 produces stoichiometric amounts of 1O2 (Reaction 26) [47, 48]. Similarly, HOCl can react with lipid hydroperoxides to yield 1O2 through a mechanism involving the generation of ROO● [49].

H2O2 + HOCl

1O 2

+ Cl + H2O + H+

(Reaction 26)

6.3. Singlet Molecular Oxygen (1O2)

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1

O2 refers to the excited states of O2, the 1 g and 1 g+ state. They have energies of 94.3 kJ/mol and 156.9 kJ/mol above the triplet ground state, respectively. The 1 g+ state has an extremely short lifetime in H2O (~10-11 s), decaying rapidly to the 1 g state, which is considered the biologically relevant form of 1O2. The lifetime of 1O2 is greatly influenced by the solvent type, being in the range of 1-5 x 10-6 s in H2O and about ten times higher in deuterium oxide (D2O). Thus, D2O is commonly employed in experiments devised to prove 1 O2 formation. Photosensitization is the most conventional source of 1O2 in biological systems. Typically, this occurs by photoexcitation (type II reactions) of endogenous photosensitizers (porphyrins, flavins, quinones, etc.) exposed to UVA [50]. These processes are especially important in the skin. Excessive production of 1O2 has been associated with disease states, such as porphyrias that are characterized by porphyrin accumulation in the skin. 1O2 promotes tumor cell death and this property is exploited in photodynamic therapy. 1 O2 can also be generated by non-photochemical reactions, usually involving inorganic peroxides (such as H2O2 and peroxynitrite) and organic hydroperoxides (ROOH). For instance, the generation of 1O2 has been evidenced during phagocytosis [35, 51], lipid peroxidation [52] and peroxidase turnover [53, 54]. The reaction between HOCl and H2O2 (Reaction 26) is the likely source of 1O2 during phagocytosis. The generation of 1O2 during lipid peroxidation has been attributed to the combination of ROO● radicals by the Russell mechanism [31, 55]. It is important to note that tertiary peroxyl radicals are unable to generate singlet oxygen because the hydrogen- on one of the ROO● is required for the elimination of O2 in the singlet-excited state [56]. The yield of 1O2 by this mechanism has been estimated to be approximately 3.9 to 14% [57]. 1 O2 is a strong two-electron oxidant that displays considerable reactivity towards electron-rich organic molecules, including nucleic acids, proteins and lipids [58]. Typically, 1 O2 adds to -bonds by three common mechanisms: addition to alkenes containing allylic hydrogen by a ene-type reaction yielding hydroperoxides; 1,4-cycloaddition to 1,3-dienes (Diels-Alder reaction) forming endoperoxides; and 1,2- cycloaddition to electron-rich alkenes producing dioxetanes [59]. Chemical reactions of 1O2 occur with second-order rate constant values that are usually lower than 107 M-1 s-1. For instance, the values obtained for the chemical reaction of 1O2 with unsaturated fatty acids are in the order of 104 M-1 s-1. In contrast, physical quenching of 1O2 occurs at much faster rates and values near the diffusion-

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Ohara Augusto and Sayuri Miyamoto

controlled limit were observed for carotenoids, among which lycopene exhibited the highest rate [60]. Reactions of 1O2 with thiols and ascorbate have been also reported. In the case of thiols, 1O2 reacts preferentially with the thiolate anion yielding several oxidation products [61]. Apparent second-order rate constant values calculated for low molecular weight thiols, such as Cys, N-acetylCys and GSH, were in the order of 106 M-1 s-1 [61]. Ascorbate reacts with 1O2 (k = 3 × 108 M-1 s-1) producing H2O2 and dehydroascorbic acid [62].

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6.4. Organic Hydroperoxides from Biomolecules Several classes of organic hydroperoxides (ROOH) are produced upon oxidation of biomolecules, including lipids, proteins and DNA [63]. Lipid hydroperoxides can be formed enzymatically during lipoxygenase, cyclooxygenase, cytochrome P450 and heme-peroxidase turnover [64, 65]. Moreover, a great number of ROOH are generated non-enzymatically by the oxidation of biomolecules mediated by radicals and 1O2 [64]. ROOH are relatively stable; however, they can participate in reactions that decrease or increase their toxicity. Normally, cells contain enzymes that reduce ROOH to their corresponding alcohols, decreasing their reactivity and toxicity. The enzymatic reduction of lipid hydroperoxides has been extensively studied. Three classes of enzymes are known to mediate their two-electron reduction: GPx, glutathione S-transferases (GST) [64, 66] and Prx [20]. Toxicity is increased when ROOH are converted to RO or ROO [3]. This can occur especially in the reaction of ROOH with transition metal ions, hemoproteins and other one-electron oxidants [3]. ROOH can react with reduced and oxidized states of free transition metal ions to form oxyl radicals. In the first case, RO● is produced whereas in the second case, ROO● is produced [3, 64]. Oxyl radicals are responsible for propagating the oxidation process as well as for the generation of other highly reactive products capable of causing modifications in proteins and nucleic acids. These include electrophilic aldehydes, epoxides, ketones, and excited species, such as 1O2 and electronically excited carbonyl species [63, 67].

6.5. Ozone (O3) O3 is present in polluted atmospheres, and inhalation of this toxic triatomic gas can induce lung injury and inflammation. In biological systems, O3 is thought to be generated during antibody catalyzed oxidation of H2O to H2O2 [68]. In the proposed mechanism, antibodies use H2O as an electron source, facilitating its addition to 1O2 to form hydrogen trioxide (H2O3) as the first intermediate in a cascade of reactions that eventually leads to O3. The generation of O3 in human tissues has been postulated based on the detection of chemical signature products, including isatin sulfonic acid and cholesterol secoaldehyde [69]. However, these markers are not specific for O3, casting doubts on whether O3 can be generated endogenously by cells [70-72]. Reactions of O3 with fatty acids, cholesterol, amino acids and DNA have been characterized. O3 adds directly to the double bonds in fatty acids, giving rise to Criegee ozonide (10%) and a hydroxyhydroperoxide intermediate (90%) that decomposes to form aldehydes and H2O2 (Reaction 27) [70]. With saturated organic targets and inorganic

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Oxygen Radicals and Related Species

compounds, O3 reacts with nucleophiles, especially those containing nitrogen or sulfur atoms by oxygen-transfer mechanism to generate a trioxide intermediate. Decomposition of this intermediate is reported to generate 1O2 and the corresponding oxidized product [73]. In aqueous solutions, one electron-transfer reactions can also occur, leading to the formation of ozonide (O3 ), HO● and O2●- [73].

O3 H2O

+

+ H2O2

(Reaction 27)

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Conclusions In its initial four decades, the field of “free radical research in biology” has focused on understanding the mechanisms by which radicals and oxidants are produced in vivo and how proteins, lipids and DNA are oxidized by them resulting in cell damage and, eventually, disease. The current picture is more complex, though. An increasing body of evidence suggests that apart from being potentially toxic, radicals and oxidants also exert important (patho)physiological signaling functions (Figure 3). The diverse biological activities of oxygen radicals and related species are likely rooted in the oxygen-dependent evolution of complex life forms. Although the chemical properties and reactivities of relevant biological oxidants are becoming well understood, it remains less clear how these properties translate into cellular and tissue responses. Further advances in the field will require interdisciplinary approaches combining chemical reasoning with biological insights.

Acknowledgments Our laboratories are supported by grants from Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP), Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES). The authors are members of the INCT de Processos Redox em Biomedicina-Redoxoma (CNPq/FAPESP/CAPES).

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Ohara Augusto and Sayuri Miyamoto

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[58] Cadet J, Di Mascio P. Peroxides in biological system. In: Rappoport Z, ed. The chemistry of peroxides. West Sussex: John Wiley & Sons Ltd; 2006:915-99. [59] Frimer A. Singlet O2. Boca Raton, FL: CRC; 1985. [60] Di Mascio P, Kaiser S, Sies H. Lycopene as the most efficient biological carotenoid singlet oxygen quencher. Arch Biochem Biophys 1989;274:532-8. [61] Devasagayam TPA, Sundquist AR, Di Mascio P, Kaiser S, Sies H. Activity of thiols as singlet molecular oxygen quenchers. J Photochem PhotobiolB 1991;9:105-16. [62] Kramarenko GG, Hummel SG, Martin SM, Buettner GR. Ascorbate reacts with singlet oxygen to produce hydrogen peroxide. Photochem Photobiol 2006;82:16347. [63] Miyamoto S, Ronsein GE, Prado FM, et al. Biological hydroperoxides and singlet molecular oxygen generation. IUBMB Life 2007;59:322. [64] Girotti AW. Lipid hydroperoxide generation, turnover, and effector action in biological systems. J Lipid Res 1998;39:1529-42. [65] Caro AA, Cederbaum AI. Role of cytochrome P450 in phospholipase A2- and arachidonic acid-mediated cytotoxicity. Free Radic Biol Med 2006;40:364-75. [66] Brigelius-Flohé R. Tissue-specific functions of individual glutathione peroxidases. Free Radic Biol Med 1999;27:951-65. [67] Esterbauer H, Schaur RJ, Zollner H. Chemistry of 4-hydroxynonenal, malonaldehyde and related aldehydes. Free Radic Biol Med 1991;11:81-128. [68] Wentworth P, Jr., McDunn JE, Wentworth AD, et al. Evidence for antibodycatalyzed ozone formation in bacterial killing and inflammation. Science 2002;298:2195-9. [69] Wentworth P, Jr., Nieva J, Takeuchi C, et al. Evidence for ozone formation in human atherosclerotic arteries. Science 2003;302:1053-6. [70] Pryor WA, Houk KN, Foote CS, et al. Free radical biology and medicine: it's a gas, man! Am J Physiol Regul Integr Comp Physiol 2006;291:R491-511. [71] Brinkhorst J, Nara SJ, Pratt DA. Hock ceavage of cholesterol 5α-hydroperoxide: an ozone-free pathway to the cholesterol ozonolysis products identified in arterial plaque and brain tissue. J Am Chem Soc 2008;130:12224-5. [72] Uemi M, Ronsein GE, Miyamoto S, Medeiros MHG, Di Mascio P. Generation of cholesterol carboxyaldehyde by the raction of singlet molecular oxygen [O2 (1Δg)] as well as ozone with cholesterol. Chem Res Toxicol 2009;22:875-84. [73] Munoz F, Mvula E, Braslavsky SE, von Sonntag C. Singlet dioxygen formation in ozone reactions in aqueous solution. J Chem Soc-Perkin Trans 2 2001:1109-16. [74] Buettner GR. The pecking order of free radicals and antioxidants: lipid peroxidation, -tocopherol and ascorbate. Arch Biochem Biophys 1993;300:535-43.

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 3

Nitric Oxide and Derived Oxidants Silvina Bartesaghi1,2,3, Natalia Romero2,3 and Rafael Radi2,3,* 1

Departamento de Histología, 2Departamento de Bioquímica and 3Center for Free Radical and Biomedical Research, Facultad de Medicina, Universidad de la República, Montevideo, Uruguay

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Introduction In mammalian cells the free radical nitric oxide (●NO, also known as nitrogen monoxide) has essential roles in a variety of biological processes, including regulation of blood pressure, platelet aggregation and neuron plasticity, among others [1, 2]. Nitric oxide and its secondary radical and non-radical-derived species can also be cytotoxic intermediates produced by activated macrophages and neutrophils which may participate in immune responses, inflammation and degenerative processes (e.g. arthritis, atherosclerosis, neurodegeneration). The chemistry of ●NO in biological systems can be divided into two main groups of reactions, direct and indirect. Direct effects are defined by those in which ●NO directly reacts fast enough with specific biomolecules such as transition metal complexes and free radicals (e.g. protein- and lipid-derived radicals) among others. Indirect effects result from the reaction with molecular oxygen (O2), superoxide radical (O2●-) and redox-active compounds, which lead to the formation of derived oxidants, collectively known as reactive nitrogen species (RNS) [3-6]. RNS include radical species represented by ●NO and nitrogen dioxide (●NO2) and nonradical species such as nitroxyl (HNO), nitrite (NO2-), dinitrogen trioxide (N2O3), dinitrogen tetraoxide (N2O4), peroxynitrite 2, nitrate (NO3-), and nitryl chloride (NO2Cl) (Figure 1).

* 2

Email: [email protected] The term peroxynitrite refers to both peroxynitrite anion (ONOO-) and peroxynitrous acid (ONOOH). IUPAC recommended name is oxoperoxonitrate (1-).

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Figure 1. Main reactive nitric oxide-derived species and connection with ROS. The steps from ●NO to other derived species are shown. Radicals (i.e. ●NO and ●NO2) are shown in red. The evolution from ●NO to different reactive species depend on redox reactions (e.g. ●NO to HNO, one-electron reduction; ●NO to NO2-, one-electron oxidation) or radical-radical combinations (e.g. ●NO plus ●NO2 to yield ●N2O3). Highly transient species nitrosonium (NO+) and nitronium (NO2+) cations (shown in brackets) readily react with water to yield the stable products nitrite (NO2-) and nitrate (NO3-), respectively. The formation of secondary non-radical species such as nitryl chloride (NO2Cl) and peroxynitrite is indicated; peroxynitrite can secondarily yield ● NO2. MPO and EPO denote myeloperoxidase and eosinophil peroxidase, respectively, and can promote the one-electron oxidation of NO2- to ●NO2 via a H2O2-dependent reaction.

Reactive nitrogen species have unique and distinct reactivities towards biomolecules such as proteins [7], lipids [8, 9] and DNA [10, 11] and promote oxidation, nitrosation and nitration reactions. Importantly, biologically-relevant factors such as the redox microenvironment (e.g. presence of reductants such as ascorbate or glutathione, transition metal centers and oxygen levels), abundance of biotargets, pH, relative hydrophobicity of the milieu and cell/tissue compartmentalization will critically determine the outcome of RNSdependent reactions at the chemical level.

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Nitric Oxide and Derived Oxidants Table 1. Nitrogen oxides (NOx) Compound

Common Name

N2 NOHNO N2O ● NO NO2HNO2 NO+ N2O3 ● NO2 N2O4 NO2+ NO3HNO3 ONOOONOOH

nitrogen nitroxyl anion nitroxyl nitrous oxide nitric oxide nitrite nitrous acid nitrosonium ion dinitrogen dioxide nitrogen dioxide dinitrogen tetroxide nitronium ion nitrate nitric acid peroxynitrite peroxynitrous acid

IUPAC name nitrogen oxonitrate (-1) oxonitric acid dinitrogen monoxide nitrogen monoxide dioxonitrate (-1) dioxonitric acid nitrosyl cation dinitrogen dioxide nitrogen dioxide dinitrogen tetroxide nitryl cation nitrate nitric acid oxoperoxonitrate (-1) oxoperoxinitric acid

Oxidation state 0 +1 +1 +1 +2 +3 +3 +3 +3 +4 +4 +5 +5 +5 +5 +5

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2. Chemistry of Nitric Oxide and Related Nitrogen Species The reaction of ●NO with oxygen leads to the formation of a variety of nitrogen oxides and related species which in turn have different chemical features. Nitrogen has an atomic number (seven) which is situated between carbon (six) and oxygen (eight), and therefore exhibits chemical properties (such as electronegativity) intermediate between oxygen and carbon. The strong electronegativity of oxygen and nitrogen makes most nitrogen oxides strongly oxidizing species.

Figure 2. Enzymatic nitric oxide synthesis by NOS.

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Table 1 summarizes the chemical nature and the oxidation states of different nitrogen compounds, including nitrogen oxides, which in turn lead to different reactivities and properties. The nitrogen compounds have oxidation states that range from -3 (i.e. ammonium and amines, not shown) to +5 (i.e. nitrate and peroxynitrite). In addition, common and IUPAC recommended names of the different compounds are listed.

3. Nitric Oxide

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3.1. Formation and Signaling Actions Nitric oxide is a ubiquitous intracellular messenger which mediates several physiological processes such as neurotransmission, vasodilatation, immune response and platelet aggregation [12]. It is produced from L-arginine, NADPH and O2 in a reaction catalyzed by several isoforms of nitric oxide synthase (NOS) that use FAD, FMN, tetrahydrobiopterin, heme and calmodulin as cofactors. In this complex reaction, electrons are transferred from NADPH through the flavins to heme-bound O2 that adds to the guanidinium group of Larginine forming N-hydroxy-L-arginine as an enzyme-bound intermediate; the latter is oxidized to citrulline and ●NO is formed [13, 14] (Figure 2). Three NOS isoforms have been isolated and cloned and are referred to as neuronal NOS (NOS I or nNOS), inducible NOS (NOS II or iNOS) and endothelial NOS (NOS III or eNOS). Despite this nomenclature, these isoforms are expressed in a broad range of mammalian tissues. The three isoforms differ in their primary sequence, mode of expression and regulation mechanisms, yet they are structurally and catalytically very similar. The NOS I and III isoforms are expressed constitutively and produce transient (lasting seconds) low fluxes (in the nM range) of ●NO in response to intracellular Ca2+ increases. NOS II is expressed in response to cytokines and produces significantly higher ●NO fluxes (> 100-fold more than NOS I) which may last for hours and are independent of cytosolic Ca2+ concentrations. The existence of a fourth constitutive intramitochondrial NOS (mtNOS), encoded by the nNOS gene and containing posttranslational modifications, has also been proposed. This enzyme may play a role in bioenergy metabolism; however this issue remains controversial. There is also sufficient evidence indicating that ●NO can also be formed by an NOSindependent pathway (described in section 6). Aside from its physiological signal transducing actions, it soon became evident that ●NO may also participate as a cytotoxic effector molecule and/or pathogenic mediator when produced at high rates by either nitric oxide synthase (iNOS) induced-inflammation or overstimulation of the constitutive forms (eNOS and nNOS) [1, 2].

Figure 3. Nitric oxide chemical structure.

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Nitric oxide is a relatively stable free radical and is neither a strong oxidant (E0 ' ●NO / NO = 0.39 V) [15] nor a reducing compound (E0 ' NO+ / ●NO- = 1.2 V) [16]. Much of ●NOmediated toxicity in the context of cell/tissue oxidative stress is due to the formation of ●NOderived secondary intermediates which are typically more reactive that ●NO itself. The reactivity of ●NO depends on its physical properties, such as size, diffusion rates and lipophilicity, and also on its particular chemical reactivity towards a variety of cellular targets. -

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3.2. Chemical and Physico-chemical Properties of Nitric Oxide Nitric oxide is a very simple molecule, consisting of one atom of oxygen bonded to one atom of nitrogen by a triple bond. Oxygen has a total of six valence electrons (electrons present in the outer shell with the greatest influence in bonding), while nitrogen contains five valence electrons. Therefore, ●NO has a total of eleven electrons that are organized as shown in the following Lewis dot diagram (Figure 3). Electrons in molecular orbitals are organized in pairs of two electrons, each one holding an opposite spin. Due to the odd number of electrons present in the ●NO molecule, one orbital contains a single electron accounting for the molecule‟s free radical status [3]. According to molecular orbital theory, ●NO has three fully occupied bonding orbitals with an unpaired electron in a fourth antibonding molecular orbital, which is responsible for its rapid reactions with other free radicals (i.e. oxygen and superoxide) and formation of strong ligands with transition metals. The unpaired electron situated in the antibonding orbital weakens the overall bonding between nitrogen and oxygen by one half of a bond, and therefore nitrogen and oxygen are held together by 2.5 bonds (3.0 bonding - 0.5 antibonding orbitals). The bond present between nitrogen and oxygen becomes stronger if one electron is removed from the molecule, yielding nitrosonium ion (NO+), which has the same number and distribution of electrons as molecular nitrogen (N2), with a triple bond and no unpaired electrons. Nitric oxide is not very soluble in water (~2 mM at 25°C) due to its small dipole moment, and does not react with water; thus it is not an acid anhydride. Because of its neutral and hydrophobic nature (octanol/water partition coefficient ca. 8), ●NO can easily permeate through and be concentrated in membranes, which in addition to its small size are important factors in its role as a cellular messenger. 3.3. Reaction Chemistry Nitric oxide is a relatively stable molecule that reacts preferentially with other paramagnetic species, such as other radicals or metal centers [17] with usually fast reaction rates ranging from 107 - 109 M-1 s-1. The major biologically-relevant reactions are indicated in Figure 4 and can be summarized as: i) radical – radical combination reactions between ●NO and radicals such as O2 and O2●- yielding ●NO2 and ONOO- respectively, which are more oxidizing species than ● NO itself; ●NO can also recombine with ●NO2 yielding N2O3, a strong nitrosating species; ii) formation of nitrosyl complexes with transition metal centers or nitroso-adducts with

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biomolecules (i.e. lipid radicals, L●, LO● and LOO●; protein radicals) and iii) redox reactions, such as the ●NO oxidation and reduction to yield NO+ and HNO respectively. The reaction between ●NO and molecular oxygen (O2) is a global third order reaction (k = 3 x 106 M-1 s-1) [18], resulting in the formation of reactive species such as ●NO2, as well as nitrite (NO2-) and nitrate (NO3-), in a process that is more efficient in hydrophobic environments [19] (Eqs 3-4). In addition, ●NO may form other RNS such as N2O3, which participates in nitrosation reactions (Figure 1). As shown, ●NO chemistry may result in oxidant or antioxidant activities, since it can react either with other radicals to give more reactive and toxic species (such as peroxynitrite or ●NO2) or terminate propagation reactions as observed in lipid peroxidation processes, yielding nitroso- and nitrolipids (see Chapter 8). Moreover, ●NO can lead to the formation of nitroxyl (HNO) (Eq. 1), the one-electron reduced and protonated congener of ●NO, which is a chemically unique species with potentially important biological activity [20]. It was also postulated to be formed as a product of the reaction between a thiol and an S-nitrosothiol or as an enzymatic reduction product of ● NO by xanthine oxidase. ●

NO + e- + H+ → HNO

(1)

Nitroxyl pKa has been corrected recently from 4.7 to 11.4 [21], which indicates that it will be fully present as HNO at physiological pH values with significant consequences for its biological activity. It is an unstable molecule, which decomposes to yield nitrous acid, and its reactions are difficult to study since it easily dimerizes to yield ●NO [22]. The one-electron oxidation of ●NO yields nitrosonium cation (NO+) which readily evolves in aqueous systems to nitrite (Eq. 2). H2O NO - e → NO → NO2-

+

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●

Figure 4. Selected biologically-relevant nitric oxide reactions.

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In biological systems, NO+ does not exist by itself unless it is bound to a polarized carrier as the one formed by reaction of ●NO with some metalloproteins [17] (see section 10.1). Thus, considering all the possible reactions shown in Figure 1, the evolution of ●NO to other reactive species will promote ●NO-dependent physiological or pathophysiological effects that are due to either direct reactions or to indirect processes through the intermediacy of RNS. The direct reactions of ●NO relate to the formation of reversible complexes or covalent adducts. The indirect processes involve species such as peroxynitrite and ●NO2 and result in chemical modifications in biomolecules such as lipids, proteins and nucleic acids.

4. Nitrogen Dioxide 4.1. Formation

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Nitrogen dioxide (●NO2) is an oxidizing free radical capable of generating damage in a variety of living systems, and a common pollutant present in air. Nitrogen dioxide (●NO2) can be produced by the autoxidation of ●NO (Eq. 3), the oxidation of NO2- by heme-peroxidases (Compound I or II) such as myeloperoxidase (MPO) [23, 24] and eosinophil peroxidase (EPO) [25] in the presence of H2O2; the homolysis of peroxynitrous acid (ONOOH); and by the reaction of ●NO with organic peroxyl radicals which can yield an intermediate compound that decomposes to ●NO2 and alkoxyl radicals. The exposure of ●NO to air (O2) leads to the formation of ●NO2 (brownish gas) which is in equilibrium with its dimer (N2O4) (Eq. 3) in the gas phase. However, in the presence of water N2O4 reacts quickly to yield nitrite (NO2-) and nitrate (NO3-) (Eq. 3). In addition, ●NO may react with ●NO2 yielding N2O3 which is mainly a nitrosating agent (Eq. 4) and in the presence of water yields nitrite. 2 ●NO + O2 → 2 ●NO2 ●

NO + ●NO2

H2O [N2O4]→ NO2- + NO3- + 2H+

H2O [N2O3] → 2 NO2- + 2H+

(3)

(4)

In the strongly acidic environment of the stomach, NO2- can also be transformed to N2O3 which in turn is in equilibrium with ●NO and nitrogen dioxide [26, 27] (see section 6).

4.2. Chemical Properties As ●NO, ●NO2 is a free radical due to its unpaired electron and can either undergo oneelectron oxidation to form nitronium ion (NO2+) or be reduced by one electron to yield nitrite (NO2-) (Figure 5, Table 1) [3]. In ●NO2 and NO2-, the presence of electrons on the lone nitrogen orbital repels the two oxygen atoms generating an asymmetric bent structure. This effect is more important for NO2than for ●NO2, due to the presence of two electrons in nitrite [3].

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Figure 5. Nitronium ion, nitrogen dioxide and nitrite chemical structures.

4.3. Reaction Chemistry Nitrogen dioxide is a relatively potent oxidant formed in biological systems (E0 ' ●NO2 / NO2- = 0.99 V) [28]. It can concentrate in membranes and react with several biomolecules, and participate in electron transfer reactions, recombination with other radicals, addition to double bonds and abstraction of hydrogen atoms in unsaturated compounds, phenols and thiols [23]. It is especially important in the reactions of protein tyrosine nitration (Chapter 6) and is also involved in processes of lipid oxidation and nitration (Chapters 7-8).

5. Peroxynitrite

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5.1. Formation Peroxynitrite is a powerful oxidant (E0´ ONOOH/ NO2- =1.4 V) [29], formed in vivo by the diffusion-controlled reaction between O2●- and ●NO (k= 4-16 x 109 M-1s-1) [30-32] (Eq. 5). ●

NO + O2● - → ONOO-

(5)

Peroxynitrite is a reactive peroxide continuously formed under basal metabolic condition, though its biological actions become particularly important under enhanced production of O2●- and ●NO, and has been implicated in the pathophysiology of numerous diseases [33-37]. Peroxynitrite anion (ONOO-) is in equilibrium with peroxynitrous acid (ONOOH), which is a rather strong acid (pKa= 6.8) [36] (Eq. 6); thus, under biological conditions both species (ONOO- and ONOOH) will be present at ratios depending on local pH. ONOO- + H+

ONOOH → ●NO2 + ●OH

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5.2. Chemical Properties Peroxynitrite anion is a relative stable species, while peroxynitrous acid is a short-lived species which undergoes unimolecular decomposition either by homolysis to hydroxyl radical (●OH) and ●NO2 or isomerization to nitrate in 30 and 70 % yields, respectively (k = 0.9 s-1 at 37°C and pH 7.4). The oxygen-oxygen bond in peroynitrous acid has a low bond dissociation energy (BDE), therefore protonation of peroxynitrite anion favors the homolytic cleavage of this peroxide. Peroxynitrite exists under two possible conformations, as proposed by quantum mechanical calculations and spectral data. The bond between the nitrogen and the peroxidic oxygen has a partial double bond character. Two possible conformers can be defined around this bond, cis and trans, however, in aqueous solutions both rotamers exchange in a rapid equilibrium [29] and despite initial belief, do not play major roles in peroxynitrite reaction biochemistry (Figure 6).

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5.3. Reaction Chemistry In biological systems, peroxynitrite undergoes different types of reactions which can be classified in three main groups: i) direct oxidation reactions, ii) nucleophilic addition to carbon dioxide and iii) homolytic cleavage of peroxynitrous acid (Figure 7). In direct reactions, peroxynitrite promotes one or two-electron oxidations, such as those observed for transition metal centers (Figure 7, I) and thiols (Figure 7, II), with rate constants that range between 103 -107 M-1s-1 [29, 38]. The nucleophilic addition of ONOO- to CO2 (1-2 mM in cells / tissues) is an important reaction regarding the fate of peroxynitrite, yielding a transient nitrosoperoxo-carboxylate adduct. This process renders ●NO2 and carbonate radical (CO3●) with yields of about 35% [39-41] which in turn mediate several nitroxidative processes (Figure 7, III). Finally, in the absence of other targets, peroxynitrous acid homolytically decomposes to release ●NO2 and ●OH with 30% yields, while the remaining ONOOH directly isomerizes to nitrate (Figure 7, IV). However, under physiological conditions this reaction is quantitatively not very important (probably with some exceptions like in acidic or hydrophobic environments) due to the high concentrations of peroxynitrite biotargets that outcompete homolysis.

Figure 6. Peroxynitrite anion chemical structure and conformations.

Through these processes, peroxynitrite mediates several direct oxidation reactions (e.g thiols) or indirect reactions such as tyrosine nitration and lipid peroxidation involving peroxynitrite-derived radicals (i.e. ●NO2, CO3●- and ●OH). Hydroxyl radical is a more powerful oxidant compared to ●NO2 and CO3●-; however, due to its high reactivity, it usually

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reacts rapidly (~109 M-1s-1) with most biomolecules in a non-selective manner. Peroxynitrite, ● NO2 and CO3●- react at slower rates and are much more selective oxidants than ●OH.

Figure 7. Peroxynitrite reaction pathways. Extracted from [29].

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6. Nitrite and Nitrate Nitrite is the one-electron reduction product of nitrogen dioxide, and is relatively stable in biological systems in the absence of hemoglobin. Nitrite in acid media yields nitrous acid (HNO2), pKa of 3.5, which in turn evolves to nitrosonium ion (NO+), capable of performing nitrosation reactions mainly (Eq. 7-9): NO2 - + H+ 2HNO2 N 2O 3

HNO2

(7)

N2O3 + H2O

(8)

●

NO + ●NO2

(9)

While typically considered as “end points” of ●NO metabolism, in recent years NO2- (and NO3-) anions have been documented as potential sources of ●NO. Nitrite/nitrate metabolism in blood and tissues can yield ●NO and may be considered an “alternative” route of ●NO synthesis especially under low O2 concentrations where the oxygen-dependent NOS activity is greatly diminished. Several enzymatic and non-enzymatic pathways of “hypoxic” reduction of NO2- have been postulated, such as the one mediated by the electron transport chain in mitochondria, xanthine oxidoreductase and deoxyhemoglobin (see section 10.1); however, the physiological relevance of this route remains unclear. Additionally, the high level of NO3-

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present in some vegetables in our diet may constitute an important source of ●NO in the digestive tract, acting mainly as a tissue protective mediator in the gastric lumen [42, 43]. Nitrate is initially reduced to NO2- by bacteria present in the oral cavity, which can then generate a variety of nitrogen oxides, including ●NO, in the acidic milieu of the gastric lumen. Nitric oxide can also be produced non-enzymatically from NO2- in other acidic environments, for example during ischemia which may contribute to post-ischemic injury [44].

7. Dinitrogen Trioxide and Dinitrogen Tetraoxide Dinitrogen trioxide (N2O3) is formed during ●NO autoxidation when ●NO2 reacts with another ●NO molecule (Eq. 4, Figure 1), or by NO2- in acidic environments (Eqs. 7-9) and can participate in nitrosation reactions. Additionally, N2O3 can participate in deamination reactions in DNA bases [45] (see section 10.7). In turn, N2O4 is formed by dimerization of ●NO2 formed by the interaction of ●NO and O2 (Eq. 3). It is an oxidizing and nitrating agent, however its formation may only be relevant in the gas phase under conditions of ●NO2 gas exposure, due to the low probability of two ●NO2 colliding with each other.

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8. Nitryl Chloride Under inflammatory conditions, neutrophil-derived MPO catalyzes the oxidation of chloride (Cl-) by H2O2, yielding hypochlorite anion (ClO-), which is in equilibrium with hypochlorous acid (HOCl; pKa ~ 7.5). The reaction between hypochlorous acid and NO2yields nitryl chloride (NO2Cl) (Eq. 10) which in turn may mediate tyrosine oxidation, nitration and chlorination reactions, in addition to modification of DNA bases. However, the biological relevance of nitryl chloride has been questioned, due to the low rate of formation in cells and cell lysates and its consumption by endogenous thiols [46]. H+ HOCl + NO2- → OH- + NO2Cl

(10)

9. Diffusion of Reactive Nitrogen Species in Biological Systems Nitric oxide is a small, uncharged and hydrophobic molecule that can freely diffuse in biological systems, being able to cross biological membranes without need of specific channels or transporters. Its diffusion coefficient in lipid membranes is only two to three times lower than in water (D = 3300 µm2 s-1) [47] and depends on membrane lipid composition [48, 49]; thus, biological membranes do not constitute an effective barrier to ● NO [50]. Moreover, diffusion of ●NO away from the producer cell is the main factor that contributes to the decay of its intracellular concentration, even greater than the loss due to

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the sum of all reactions within the cell. These properties support the role of ●NO as a paracrine rather than autocrine signaling molecule; i.e., it acts mainly in cells different from the cell where it is produced. It can be estimated that, in a biological milieu (i.e., assuming a biological half life of 0.5-5 s), ●NO could travel a mean distance of 50-200 µm from its biological source, a distance comparable to 5-20 times the mean diameter of a cell [49], but this value is highly dependent on the proximity of a blood vessel as discussed below. Auto-oxidation represents the main decomposition route in water, however in biological systems it will only be relevant at very high ●NO concentrations as it depends on the square of its concentration (-d[●NO]/dt = 4 k3 [●NO]2[O2]). For example, considering a ●NO concentration of 100 nM (10 times the concentration needed to activate the muscular enzyme soluble guanylate cyclase, one of the main biological targets) the half-life would be ~ 2 hours. In hydrophobic environments such as in biological membranes, ●NO and O2 concentrations will be 3-4 times higher due to its higher solubility [19] which represent an increase of 50 times in the reaction rate according to the previous equation. In the vasculature, the half-life of ●NO is highly diminished by the presence of oxyhemoglobin (oxyHb). Reaction of ●NO with oxyHb yields methemoglobin (metHb) and NO3- as final products (Eq. 11).

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Hb(Fe+2)O2 + ●NO

Hb(Fe+3) + NO3-

(11)

This reaction occurs with a second order rate constant of 8 x 107 M-1s-1 [51], representing a quantitatively important fate or detoxifying route of vascular ●NO considering the high intravascular hemoglobin concentration. If hemoglobin molecules were free inside vessels instead of being contained within erythrocytes, ●NO half-life should decrease to less than 1 µs, an excessively short half-life to be compatible with its biological role as a signaling molecule. However, studies of Lancaster in the 1990‟s [52] demonstrated that due to the fact that intravascular hemoglobin is encapsulated within erythrocytes the intravascular half-life of ● NO extends to ~ 2 ms, which is in agreement with its function. The fast intravascular consumption of ●NO by oxyhemoglobin generates a deep gradient of ●NO from tissue sources to blood vessels [53]. In addition to red blood cells, another important reaction responsible for ●NO consumption is its reaction with O2●- to yield peroxynitrite (ONOO-/ONOOH) (Eq. 5). Relative to ●NO, peroxynitrite has a shorter half-life in biological systems (between 10-20 ms) due to its reactivity with several biological targets [54]. Considering a diffusion coefficient similar to that of NO3- in aqueous systems (D = 1500 2 -1 µm s ), peroxynitrite can diffuse distances between 5-50 µm which represents 1-2 cellular diameters [55, 56]. However, biological membranes will partially limit peroxynitrite diffusion. Peroxynitrite can cross biological membranes by two different routes: peroxynitrite anion can penetrate cells through anion transporters while peroxynitrous acid can cross through the lipid bilayer by simple diffusion [55] at rates similar to water molecules [57, 58]. Because of its partial limitation to diffuse through biological membranes, target compartmentalization and sites of peroxynitrite formation should be considered in studies of biological peroxynitrite reactivity. Finally, ●NO2, as a neutral and small radical species, can also readily permeate membranes; its solubility in hydrophobic environments is similar to that in aqueous phases [59].

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10. Reactivity of Nitric Oxide and Derived Oxidants with Selected Biomolecules

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10.1. Metal Centers: Nitrosylation and Redox Reactions Interactions of ●NO with metalloproteins constitute a relevant molecular mechanism accounting for signal transduction, cytoxicity and modulation of free radical chemistry. Nitric oxide can interact with different metal centers in proteins including heme iron, iron-sulfur clusters, zinc-sulfur clusters and copper. The reactions typically result in formation of stable nitrosyl-metal complexes or promote redox reactions that may alter protein function.When it interacts with a metal center, it can adopt two different binding geometries, linear or bent, reflecting different metal-NO bonding interactions. The interaction of ●NO with a metal with a linear geometry, the most common bonding mode, consists of both ζ donation from ●NO to metal and η backbonding from occupied d orbitals of proper symmetry on the metal to the η* antibonding orbitals on ●NO. In the bent geometry, there is a “donation” of one electron from the metal to ●NO to form an “NO-” species that binds to the metal in a ζ interaction, leaving an electron pair localized in a sp2 orbital on the nitrogen atom of the ligand. As a consequence of the different bonding interactions in the two different geometries, the ●NO in the linear bonding is formally an NO+ species as it is akin to ●NO donating an electron to the metal prior to bonding, while in the bent geometry the metal can be envisioned to donate first the electron to ●NO, giving NO- which then binds to the metal. Due to the fact that ●NO can coordinate to a metal as either NO+ or NO-, metal-bound nitrosyls can be either electrophilic or nucleophilic [17]. Coordination of ●NO to heme iron in hemoproteins has an important role in the physiological actions of this radical. Nitric oxide is an extremely effective ligand of ferrous heme reacting with second order rate constants typically ~107 M-1 s-1 and, unlike other simple diatomic ligands such as O2 and CO, it can also bind ferric hemes but generally with lower rate constants (102 – 107 M-1 s-1). Hemoproteins with an unoccupied sixth coordination position in heme, such as deoxyhemoglobin or guanylate cyclase, tend to react faster with ● NO than hemoproteins that have all six coordination positions in heme occupied, such as cytochrome c, as ●NO in the latter case must displace the sixth ligand, usually an amino acid. Another kinetic parameter that must be considered when studying ●NO-metal interactions is the dissociation constant: in general, once formed the nitrosyl-heme slowly dissociates, with dissociation usually faster for the ferric-heme nitrosyl complex. The presence of the protein backbone makes the combination reaction of ●NO with heme in proteins slower than with free heme, but more selective, as the relatively apolar environment present in the heme cleft makes diffusion of ●NO much more favorable than that of ●NO-derived species such as NO2-. In summary, kinetics of ●NO binding to iron heme-proteins is highly dependent on both the oxidation state and coordination environment of the iron center. Undoubtedly, a key biological function of ●NO is its action as an activator of the hemecontaining enzyme guanylate cyclase. Other similar and biologically relevant ligands such as O2 or CO also can bind heme in hemoproteins but do not possess the unique biological activity of ●NO. The reason is that contrary to what is observed with other small ligands, ●NO coordination to a ferrous heme-protein, labilizes the trans axial ligand and this modification affects the whole protein structure. In the case of guanylate cyclase, the trans axial ligand is

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an imidazole moiety from a histidine which, after ●NO- assisted release may serve to increase the catalytic activity of the enzyme resulting in production of cGMP (Eq. 12).

(12) In addition to yielding a stable heme-nitrosyl complex, a transient complex may be formed as an intermediate step during ●NO-heme interactions that lead to a redox change in the metal center and formation of NO2- or NO3- (Eqs. 13-14): X-Fe3+ + ●NO → [X-Fe3+….NO]

(13)

[X-Fe3+….NO] + H2O → X-Fe2+ + NO2- + 2 H+

(14)

Formation of the nitrosyl complex may also modify the reactivity of ●NO and make it more available for nitrosylation reactions, as discussed below. A particular example of ●NO reactivity with heme proteins is its reaction with hemoglobin that reflects several possibilities of ●NO interactions with metal centers. Nitric oxide reacts rapidly with oxyhemoglobin (HbFe2+-O2) with a second order rate constant of 8 x 107 M-1 s-1 forming a transient metHb-peroxynitrite complex, which rapidly evolves to metHb and NO3-. This reaction constitutes the main fate of ●NO inside the vascular compartment due to the high oxyhemoglobin concentration therein (Eq. 15):

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Hb (Fe2+)O2 + ●NO

Hb (Fe3+)-OONO

Hb (Fe3+) + NO3ˉ

(15)

At low oxygen concentrations, where a fraction of hemoglobin is deoxygenated, ●NO can also react with deoxyhemoglobin, (HbFe2+) with a fast second order rate constant (2 x 107 M-1 s-1) [60] to yield the stable nitrosyl-heme complex (dissociation constant ~ 10-3 – 10-5 s-1). The latter yields a characteristic electron paramagnetic resonance (EPR) signal that has been used in animal models as a footprint of ●NO overproduction in pathologies such as sepsis [61]. In addition, ●NO may also react with methemoglobin (HbFe3+), especially in conditions of ●NO excess, resulting in the formation of a ferrous-NO+ complex (Eq. 16). The latter may either be hydrolyzed to deoxyhemoglobin (HbFe2+) and NO2- (Eq. 17) or the nitrosyl group may be transferred to nucleophilic compounds (i.e., thiols, Eq. 18). [Hb3+….NO] → Hb2+─NO+

(16)

Hb2+-NO+ + H2O → NO2- + Hb2+

(17)

Hb2+─NO+ + RS- → Hb2+ + RSNO

(18)

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Figure 8. Reactions of nitric oxide and derived species with hemoglobin. Oxyhemoglobin (HbII-O2), the predominant species inside red blood cells at physiological conditions, reacts with ●NO (k = 8 x 107 M-1 s-1) or with ONOO- (k = 1.7 x 104 M-1 s-1) to yield in both cases methemoglobin as final product (HbIII). At low oxygen tensions, deoxygenated hemoglobin (HbII) can interact with ●NO (k = 2.0 x 107 M-1 s-1) to yield a stable iron-nitrosyl hemoglobin. The oxidized hemoglobin HbIII also reacts with ●NO, albeit at significantly slower rates (k = 3 x 103 M-1 s-1); this reaction yields a NO-methemoglobin complex that after a slow reduction step recovers the reduced deoxyhemoglobin (HbII). Nitric oxide can also undergo a one-electron reduction giving nitroxyl (HNO) that reacts with metHb to give iron-nitrosyl hemoglobin, a reaction that has been used for measurement of HNO in biological systems. Deoxyhemoglobin may also act as a nitrite reductase, reducing NO2- to ●NO, a reaction that could represent a pathway for increased ●NO bioavailability in vivo under hypoxic conditions.

While formation of ferrous-heme nitrosyl complexes may in general be detected both by optical spectroscopy and by EPR, ●NO-ferric heme complex is diamagnetic and therefore EPR-silent (Figure 8). The contribution of red blood cells as a source and not merely a sink of intravascular ● NO is another currently investigated issue. Several groups have indicated that deoxyhemoglobin may act as a nitrite reductase, and administration of NO2- at low doses has been reported to cause vasodilation mediated by guanylate-cyclase activation in the vascular smooth muscle cells [62]. Perhaps the most intriguing aspect pertains to how any ●NO molecule generated inside a red blood cell is exported before reacting with the abundant intraerythrocytic oxy- or deoxyhemoglobin, considering its high rate constant of reaction. Other intermediates such as N2O3 or nitrosothiols may be formed as byproducts of ●NO production that could facilitate ●NO-bioactivity exportation [63]. Another biologically-relevant reaction of ●NO with hemoproteins is the inhibition of the complex IV or cytochrome c oxidase of the mitochondrial electron transport chain. The mechanism involves ●NO binding to the sixth coordination position of the heme of cytochrome a3 competing with oxygen binding as the final acceptor. In addition, the binding constant of ●NO is significantly larger than that of O2 resulting in a reversible competitive inhibition of cellular respiration, especially at low oxygen tensions. Nitric oxide has been shown to bind also to iron-sulfur clusters (see Vol. II, Chapter 19) leading to formation of iron-nitrosyl complexes that can progressively promote cluster oxidation and disruption. In early experiments of ●NO-mediated cytotoxicity, it was shown

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that when cells were exposed to ●NO, an EPR signal characteristic of dinitrosyl iron complexes Fe(NO)2(SR)2 was detected and it was attributed to iron removed from susceptible iron-sulfur centers by ●NO, such as that of mitochondrial aconitase. This enzyme participates in the tricarboxylic-acid cycle. In aconitase and other similar dehydratases, the iron-sulfur cluster is of the type [4Fe-4S], with the peculiarity that one of four iron atoms has a free uncoordinated position, known as Feα. This iron is labile and can be attacked, by different oxidants including molecular oxygen and superoxide radical with the concomitant release of the Feα, cluster disruption and enzyme inactivation. It has been proposed that ●NO could reversible react with this Feα to produce an intermediate complex with NO+-like reactivity that could end in cluster disruption after several irreversible steps (Eq. 19).

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(19) However, studies have demonstrated that irreversible ●NO-dependent inactivation of aconitase requires non-physiological ●NO concentrations or, alternatively, formation of other ● NO-derived oxidants such as peroxynitrite to inactivate rapidly the enzyme as discussed below [64, 65]. Peroxynitrite can also react with transition metal centers of proteins, specially metalporphyrin systems such as heme proteins, non-heme iron, copper and manganese ions with high kinetic constants (from 104 to 107 M-1 s-1). In the case of hemoproteins, peroxynitrite oxidations can be one- or two-electron processes depending on the particular metal center and its starting oxidation state. Peroxynitrite reacting as a one-electron oxidant yields nitrogen dioxide (●NO2) in addition with the oxidized metal center, while two-electron redox reactions yield nitrite (NO2-). It has also been established that some metal complexes can catalyze peroxynitrite isomerization to nitrate (NO3-) without affecting the metal center [66] (Eqs. 2022). One-electron oxidation: X-Men + ONOO- X-Men+1=O + ●NO2

(20)

Two-electron oxidation: X-Men + ONOO- X-Men+2=O + NO2-

(21)

Isomerization: X-Men + ONOO-

(22)

X-Men + NO3-

Reactions of peroxynitrite with several hemoproteins such as MPO or cytochrome P450 concomitantly produce, in addition to ●NO2, oxo-metal complexes, which are strong oxidants that promote secondary redox transitions such as one-electron oxidation of nearby tyrosines. Formation of tyrosyl radical, concomitantly with ●NO2 promotes tyrosine nitration reactions (see section 10.3 and Chapter 6). Peroxynitrite can also react with iron-sulfur clusters such as the [4Fe-4S] cluster of mitochondrial aconitase yielding the inactive [3Fe-4S] enzyme. The aconitase iron-sulfur cluster has a net oxidation state of +2, with the local positive charge of the exposed Feα offering an electrostatic attraction for this anionic oxidant. The transference

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of a single electron from the exposed cluster to peroxynitrite can destabilize the cluster, causing the loss of Feα and enzyme inactivation. In the case of zinc-thiolate centers, such as the yeast alcohol dehydrogenase, peroxynitrite reaction results in thiolate oxidation and Zn release with concomitant enzyme inactivation. Peroxynitrite also reacts with the Cu/Zn and Mn centers of superoxide dismutases (SOD; Vol. II, Chapter 5) yielding high oxidant species such as oxo-copper and oxo-manganese plus ●NO2 that also promote metal-catalyzed histidine oxidation and tyrosine nitration (in CuZnSOD and MnSOD, respectively). Nitroxyl can also act as a one-electron reductant of oxidized metalloproteins such as CuZnSOD or methemoglobin. Reaction with SOD results in the reduction of the cupric atom to the cuprous form and concomitant ●NO release while in the case of metHb the ferrous nitrosyl adduct is formed.

10.2. Thiols: S-nitrosylation and Oxidation Reactions Thiols are by far the most abundant nucleophilic species present in vivo, reaching concentrations in the millimolar order. Reactive nitrogen species have been shown to readily react with both proteins and low-molecular-weight thiols (including the amino acid cysteine and the tripeptide glutathione). However, and contrary to common belief, ●NO does not easily undergo reactions with thiols. Indeed, unless high concentrations of ●NO, low oxygen tensions and/or metal traces are present, no thiol S-nitrosation or oxidation is observed. On the other hand, other RNS (such as N2O3 and peroxynitrite) readily react with thiols. In the presence of molecular oxygen, ●NO is oxidized to ●NO2 that can subsequently react with another ●NO molecule to yield N2O3. S-Nitrosation of thiols by N2O3 is facile and results in the formation of S-nitrosothiols [67] (Eq. 23).

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N2O3 + RSH → RSNO + NO2- + H+

(23)

This route may constitute a possible biological mechanism of nitrosothiol formation; however, N2O3 formation is not a fast reaction at least in hydrophilic environments. Thiol nitrosation can also occur via the intermediacy of a ferric heme nitrosyl species that confers ● NO a nitrosonium character; therefore it may perform directly a nucleophilic attack on the thiol to yield an S-nitrosothiol (Eq. 24). Fe2+ NO+ + RSH → Fe2+ + RSNO + H+

(24)

Finally, S-nitrosothiols may be formed as a product of the recombination of thiyl radicals and ●NO (Eq. 25). RS● + ●NO → RSNO

(25)

Thiyl radicals are common intermediates in free radical chemistry, especially in pathological conditions where there is enhanced O2●- and ●NO production, and hence peroxynitrite, as we will discuss later [68]. Moreover, ●NO2 reacts rapidly with thiols by a one-electron oxidation reaction.

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Peroxynitrite directly reacts with thiols by a two-electron process in which thiolates are oxidized to the corresponding sulfenic acid (ROSH). The latter is an unstable species that in the presence of accessible thiols form disulfides (RSSR´). In the case of proteic thiols, disulfides can be formed within the same protein or with low molecular weight thiols leading to mixed disulfide formation (Chapter 4). In addition, peroxynitrite-derived radicals (●OH, ● NO2, CO3●-) can oxidize thiols by one electron yielding the corresponding thiyl radical that can recombine to yield disulfide. More frequently, thiyl radicals also react with oxygen forming the thiylperoxyl radical (RSOO●) or may react with another thiol to form disulfide anion radical (RSSR●-) or recombine with ●NO to yield RSNO. Therefore, products formed from the reaction of ●NO-derived species with thiols will strongly depend on thiol and oxygen concentration as well as the concomitant presence of reactive oxygen species. Reactions of ●NO-derived species can strongly affect cellular signaling. Importantly, formation of S-nitrosothiols may be considered a route of ●NO storage, as they can act as subsequent ●NO donors in the presence of reductants or other thiols. On the other hand, a large number of proteins involved in signaling cascades such as kinases and phosphatases or the transcription factor NF- B (Vol. II, Chapter 12) contain reactive thiols that can be oxidized or S-nitrosylated under conditions of overproduction of RNS, thereby modulating cellular signaling pathways. Lastly, fast reaction of thiols with RNS such as ●NO2 or peroxynitrite may constitute efficient detoxification and antioxidant pathways in biological systems.

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10.3. Tyrosine: Nitration and other Oxidation Reactions Tyrosine is a normal constituent amino acid in proteins (approximately 3.2% on average) which can react with nitric oxide-derived oxidants such as ●NO2 and ONOO-, yielding 3nitrotyrosine (3-NT) and leading to tyrosine nitration, a post-translational modification which has important biological consequences (see also Chapter 6). Protein tyrosine nitration is associated with several pathologies [7, 69-73] such as cardiovascular disease, neurodegeneration, inflammation, and diabetic complications, and serves as a biomarker of oxidative stress in vivo, and a predictor of disease progression and severity [6, 74, 75]. Protein tyrosine nitration may result in dramatic changes in protein structure and function which in turn lead to a loss or a gain of function [7], promote protein aggregation and affect degradation, and alter phosphorylation cascades [76] and immunological responses [7, 77]. One of the most important nitrating agents in vivo is peroxynitrite, and the detection of tyrosine nitration in biological systems was initially defined as a “footprint” of peroxynitrite formation. A well known example of protein tyrosine nitration in vivo, is the inactivation of mitochondrial enzyme MnSOD by peroxynitrite-mediated nitration of Tyr-34 in a process that is catalyzed by the manganese atom. Tyrosine nitration impacts a large number of proteins in vivo with profound biological implications [6, 72, 73]. There is consensus that tyrosine nitration may occur biologically by various routes (e.g. peroxynitrite-or hemeperoxidase-dependent) that are mainly based in free radical chemistry. Indeed, 3–NT as evidenced in vitro and most probably in vivo is the product of at least two consecutive reactions: (1) the intermediate formation of the tyrosyl-phenoxyl radical by oneelectron oxidation of tyrosine and (2) the combination of the phenoxyl radical with ●NO2

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derived from (a) peroxynitrite decomposition [78], (b) hydrogen peroxide (H2O2)-dependent nitrite oxidation catalyzed by hemeperoxidases (e.g. myeloperoxidase) [79], eosinophil peroxidase [25], (c) aerobic oxidation of ●NO [80], or (d) nitrite under acidic conditions, to yield 3-NT (Figure 9). The oxidation of tyrosine to yield tyrosyl radical may be achieved by a number of oxidants such as ●OH, CO3●−, high oxidation states of transition metal complexes (Me (n+1)+ =O; compound I of hemeperoxidases) or ●NO2, which reacts with tyrosine at moderate rates, which may occur in lungs exposed to ●NO/●NO2-contaminated air [82]. In addition, lipidderived peroxyl (LOO●; E°LOOH/LOO● = 1.02 V) radicals were shown to function as possible one-electron oxidants which may participate in the nitration pathway in hydrophobic biocompartments [81, 83]. Indeed, these alternative one-electron oxidants may play important roles in membranes and lipoproteins where, due to the high concentration of unsaturated fatty acids, lipid peroxidation reactions take place and yield large amounts of LOO● radicals (Chapter 7). An alternative mechanism for the formation of 3-NT without the participation of ●NO2 involves the reaction of tyrosyl radical with ●NO to form 3nitrosotyrosine followed by a two-electron oxidation to yield 3-NT with the intermediate formation of iminoxyl radical [7]. This mechanism may be relevant in transition metalcontaining proteins that can oxidize 3-nitrosotyrosine such as prostaglandin H synthase-2 [84]. Additional tyrosine nitration pathways, of uncertain biological significance may involve the following radical-independent electrophilic aromatic nitration mechanisms: 1) the reaction of ONOO- in the presence of transition metal centers through the formation of a complex that operates as a carrier of nitronium cation (NO2+), and 2) reaction with nitryl choride which can participate in tyrosine nitration and chlorination reactions [85]. Concomitant with the formation of 3-NT, other side products can be formed (Figure 9). The combination of two tyrosyl radicals yields the dimerization product 3,3′-dityrosine, a naturally occurring cross-linked amino acid which has also been used as a footprint of oxidative stress.

10.4. Reaction of Reactive Nitrogen Species with other Amino Acids RNS can lead to the modifications of some other amino acids, mainly tryptophan, methionine and histidine. The only amino acids that directly react with peroxynitrite are cysteine (mentioned above), methionine and tryptophan, with second order rate constants of 4.5 x 103 M-1 s-1 [36], 1.8 x 102 M-1 s-1 [86] and 37 M-1 s-1 [87]. Other amino acids do not directly react with peroxynitrite, however they can be modified by the action of peroxynitritederived radicals (●NO2 and ●OH), as happens with tyrosine, phenylalanine and histidine [88]. Peroxynitrite and ●NO2 can lead to free and protein tryptophan nitration (yielding several nitrated products: 1-, 4-, 5-, 6-and 7-nitro-tryptophan) [87], oxidation (hydroxy-tryptophan, N-formyl kynurenine and trytophanyl radical) [89], and nitrosation (nitroso-tryptophan) [90]. In the presence of peroxynitrite, the main nitration products are 1- N-nitroso-tryptophan and 6-NO2-tryptophan [91].

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Figure 9. Tyrosine nitration reactions. Reaction 1 represents tyrosine one-electron oxidation that yields the intermediate tyrosyl radical (A); this reaction reverts to tyrosine in the presence of numerous reductants. In the presence of nitrogen dioxide radical (reaction 2) the main product is 3–nitrotyrosine (B). In reaction 3, nitric oxide reacts with tyrosyl radical leading to the formation of an intermediate species 3–nitrosotyrosine (C) which in turn can be oxidized by two one-electron steps (reaction 4) to yield 3-nitrotyrosine. Tyrosyl radical may dimerize (reaction 5) to yield 3,3‟–dityrosine (D) or may react with other radicals (i.e ·X = ·OH) (reaction 6) yielding the hydroxylated derivative DOPA, then (E). Extracted from (81).

Nitrated tryptophan (and other oxidation products) also results from the reaction of human CuZnSOD and bovine serum albumin (BSA) with peroxynitrite and myeloperoxidase/hydrogen peroxide/nitrite [91]. Nitroso-tryptophan formation has also been reported in BSA [90] and probably implies the intermediacy of N2O3. Usually tryptophan modification in vivo is less abundant than tyrosine modification because tryptophan residues are present in smaller amounts and are normally buried inside proteins. Free methionine can react with peroxynitrite via one- or two electron oxidation pathways, leading to the formation of secondary products such as ethylene in the one-electron oxidation, and methionine sulfoxide and nitrite in the two-electron oxidation where peroxynitrous acid is the main reactive species [86]. Peroxynitrite is capable of oxidizing methionine residues in proteins, as can be found in alpha 1-proteinase inhibitor (alpha 1PI), where peroxynitrite primarly oxidizes the methionine residue [92]. Phenylalanine does not directly react with peroxynitrite; however, its exposure to this oxidant leads to the formation of several products, such as p-, m- and o-tyrosine, as well as nitro-phenylalanine [93].

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Finally, histidine can become oxidized by peroxynitrite derived-radicals to histidinyl radical, 2-oxo-histidine and nitro-histidine [89]. Human CuZnSOD is readily inactivated by peroxynitrite, by reaction with histidine residue in the active site of the enzyme and the intermediate formation of histidinyl radical [94], which in turn by reaction with oxygen can evolve to 2-oxo-histidine, or nitro-histidine in the presence of ●NO2 [95]. Nitric oxide itself does not directly react with amino acids, but can react with the amino acid-derived radicals, in which case ●NO can lead to the formation of transient or permanent nitroso-adducts (e.g. nitroso-tyrosine, [96] and nitroso-tryptophan [90]).

10.5. Lipids Nitrogen dioxide initiates lipid peroxidation by one-electron abstraction on allylic hydrogens in fatty acid double bonds [97] and also promotes cis to trans isomerization [98]. Peroxynitrite initiates lipid peroxidation after homolysis of ONOOH [99]. Both ●NO and ● NO2 can readily react with lipid-derived radicals yielding a myriad of intermediates that lead to the formation of oxidized and nitrated fatty acids [8, 9, 100] (see Chapter 8).

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10.6. Sugars RNS such as peroxynitrite and ●NO2 can react with sugars, which are present in cells as monosaccharides (e.g. glucose) or polysaccharides (e.g. glycosaminoglycans); however, few data regarding these reactions are available. The reaction between peroxynitrite and glucose (and other biologically-relevant alcohols, such as fructose and glycerol) yields nitroso-derivatives. The reaction chemistry is ill-defined but most likely involves the reaction of the sugar with peroxynitrite-derived radicals to yield compounds that have physiological properties similar to those of organic nitrites and nitrates [101]. Regarding the reaction of peroxynitrite with polysaccharides, it has been recently reported that the reaction of peroxynitrite with glycosaminoglycans (GAGs) results in extensive polymer fragmentation, affecting hyaluronan, heparin and chondroitin, dermatan and heparan sulfates [102]. GAGs are large polysaccharides consisting of repeating disaccharide units, either in free form (hyaluronan) or as proteoglycan complexes which are important components of the extracellular matrix and glycocalix. Damage to the extracellular matrix is an important factor in many inflammatory diseases [102], including osteoarthritis where excess ●NO and peroxynitrite may play significant roles. Peroxynitrite also modifies the heparan sulfate perlecan which is an important component of basement membranes and plays a key role in extracellular matrix structure. Peroxynitrite-related structural and functional modifications of perlecan may contribute to the formation of human atherosclerotic lesions [103]. The mechanism of fragmentation depends on the formation of oxidizing radicals, including peroxynitrite-derived ●OH, CO3●- and ●NO2 [104].

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Table 2. Selected biologically-relevant modifications of biomolecules by RNS

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10.7. Nitrogen Bases RNS such as peroxynitrite, nitrogen oxides and nitryl chloride have been implicated in oxidation and nitration of DNA bases, most notably guanine. Under physiological conditions, peroxynitrite readily reacts with guanine yielding two main nitrated products, 8-nitroguanine (8-nitro-G) [105] and 5-nitro-4-guanidinohydantoin (NI) [106], via the intermediate formation of guaninyl radical, in addition to two main oxidation products, 8-oxo-guanine [107] and 2,5-diamino-4H-imidazol-4-one [108]. Importantly, peroxynitrite-derived ●OH may lead to DNA strand breaks [107] (see also Chapter 9). Finally, the reaction of peroxynitrite and other RNS with secondary amines yield N-nitrosamines and N-nitramines [109]. Trioxide of dinitrogen can also directly affect DNA by base deamination, which in the case of guanine:cytosine base pair results in the formation of xanthine and uracil respectively, and therefore in dramatic changes in DNA base composition [45]. Recently, the biological presence of 8-nitro-cGMP, a nitrated guanine nucleotide, (8nitroguanosine 3,5-cyclic monophosphate) as a signaling molecule has been described [110], expanding the breadth of action of nitrated intermediates in biological systems.

10.8. Low Molecular Weight Antioxidants

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RNS, mainly peroxynitrite and ●NO2, can react with either hydrophobic and hydrophilic antioxidants such as - and -tocopherol [111-113], ubiquinol [114], and ascorbic [115] and uric acid [116, 117] which are known to attenuate RNS-dependent oxidation and nitration reactions (Table 2). In summary, RNS can promote a wide variety of nitroxidative modifications in biomolecules (i.e. proteins, lipids, sugars, DNA) and antioxidants, several of which are summarized in Table 2.

Conclusions Nitric oxide (●NO) is a relatively stable free radical mainly formed enzymatically by nitric oxide synthases, and can evolve to a variety of secondary radical and non-radical oxidant species. Nitric oxide is a neutral and hydrophobic molecule which can readily cross biological membranes. Some of the biological effects of ●NO depend on direct reactions with biotargets while others rely on indirect reactions through its derived oxidants. Direct reactions involve i) radical – radical combination reactions, ii) formation of nitrosyl complexes with transition metal centers or nitroso-adducts with biomolecules and iii) redox reactions. Nitric oxide-derived oxidants are also grouped as reactive nitrogen species (RNS) and include nitrogen dioxide (●NO2), peroxynitrite (ONOO-/ONOOH) and dinitrogen trioxide (N2O3). Peroxynitrite, the product of the diffusion-controlled reaction between ●NO and superoxide radical (O2●-), is a reactive peroxide that can participate in two-electron oxidation reactions or evolve to secondary radical intermediates such as ●NO2, hydroxyl radical (●OH) and carbonate radical (CO3●-). Peroxynitrite anion crosses biomembranes through anion channels, while ONOOH freely permeates lipid bilayers.

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Nitrogen dioxide is a good one-electron oxidant and participates in nitration reactions (addition of a -NO2 group); N2O3 participates in both oxidation and nitrosation (addition of a NO-group) reactions. Nitrite (NO2-) and nitrate (NO3-) are typically end products of ●NO catabolism; however, under certain circumstances (acidic pH, reaction with deoxyhemoglobin, salivary reductases) they can lead to the formation of ●NO via NOSindependent reactions. Key reactions of ●NO with biotargets involve the formation of reversible heme-nitrosyl complexes with either guanylate cyclase, leading to enzyme activation, or cytochrome a3 to inhibit mitochondrial respiration. Nitric oxide can also participate in radical chain termination reactions and exert antioxidant effects. Relevant reactions of RNS with amino acids in proteins include thiol S-nitrosation, thiol oxidation and tyrosine nitration; similarly, fatty acids and DNA bases can be nitrated. Due to the short biological half-life of RNS, many of the chemical modifications in biomolecules can be used as "footprints" of their presence in biological systems. In addition, some of the modified biomolecules exhibited altered biological function (e.g. tyrosine nitrated proteins) or represent novel signal transducing agents (e.g. S-nitrosothiols, nitrated fatty acids, nitrocGMP). A variety of low molecular weight antioxidant compounds and even some enzymatic systems (e.g. peroxiredoxins) can readily scavenge or decompose RNS and attenuate or neutralize their biological effects.

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Acknowledgments This work was supported by grants from Agencia Nacional de Investigación e Innovación, Uruguay to SB (FCE_362), NR (FCE_361), and RR (FCE_2486), Comisión Sectorial de Investigación Científica (CSIC, Universidad de la República,Uruguay) and the Howard Hughes Medical Institute to RR. RR is a Howard Hughes International Research Scholar.

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[120] O'Donnell VB, Eiserich JP, Chumley PH, Jablonsky MJ, Krishna NR, Kirk M, et al. Nitration of unsaturated fatty acids by nitric oxide-derived reactive nitrogen species peroxynitrite, nitrous acid, nitrogen dioxide, and nitronium ion. Chem Res Toxicol. 1999 Jan;12(1):83-92. [121] Byun J, Henderson JP, Mueller DM, Heinecke JW. 8-Nitro-2'-deoxyguanosine, a specific marker of oxidation by reactive nitrogen species, is generated by the myeloperoxidase-hydrogen peroxide-nitrite system of activated human phagocytes. Biochemistry. 1999 Feb 23;38(8):2590-600. [122] Suquet C, Warren JJ, Seth N, Hurst JK. Comparative study of HOCl-inflicted damage to bacterial DNA ex vivo and within cells. Arch Biochem Biophys. 2010 Jan 15;493(2):135-42.

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Chapter 4

Sulfur-centered Radicals Christian Schöneich* Department of Pharmaceutical Chemistry, The University of Kansas, 2095 Constant Avenue, Lawrence, KS 66047, U.S.

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1. Introduction Sulfur-centered radicals play an important role in redox biochemistry, where they participate in enzymatic processes [1], the detoxification of carbon- and oxygen-centered radicals [2,3], the post-translational modification of peptides and proteins [4], the cis-transisomerization of unsaturated fatty acids [5], and possibly in long-range electron transfer reactions [6,7]. In biological systems, endogenous sulfur-centered radicals originate primarily from redox processes of the amino acids cysteine (Cys; structure 1) and methionine (Met; structure 2).

The resulting radicals/radical ions possess a rich chemistry, which will be described separately for Cys and Met below. Though the chemistry of sulfur-centered free radicals extends far beyond that of Cys and Met, we will predominantly focus on radicals from Cys and Met due to their biological relevance.

*

Phone: (785) 864 4880; Fax: (785) 864 5736; Email: [email protected]

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(1)

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2. Formation of Thiyl and Perthiyl Radicals The mercapto group of the Cys side chain reacts via hydrogen and electron transfer processes, depending on its ionization state (equilibrium 1). While for free Cys, pKa,RSH 8.2 [8], protein-bound Cys displays pKa values between ca. 5 [9] and 11 [8]. Specifically carbon-centered radicals react with the protonated mercapto group under hydrogen transfer to generate thiyl radicals 3 (reaction 2), while oxidizing radicals (Ox ) preferentially react under electron transfer with the deprotonated sulfhydryl group (reaction 3) [10]. Especially in peptides and proteins containing cystine, i.e. a disulfide bond between two Cys residues, thiyl radicals 3 can be generated via one-electron reduction (reactions 4-6) [11] and homolytic substitution, SH2 (reaction 7) [12]. Reaction 4 generates intermediary twocenter-three-electron-bonded disulfide radical anions, which only have a rather short life-time in the absence of excess thiolate (see reaction 12, below) unless they are incorporated within a rigid protein structure, which would prevent dissociation into thiyl radical and thiolate [1315]. The dissociation of the radical anion is further accelerated through protonation (reaction 6). [Note that the reduction of proteins in the solid state has been observed to yield ultimately perthiyl radicals 4 [16] (for a structure of 4, see reaction 10 below)].

(2)

(3) CysSSCys +

e-

[CysSSCys]-

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(4)

77

Sulfur-centered Radicals [CysSSCys][CysSSCys]-

+ H+

CysSSCys +

R

CysS-

+

CysS

3

(5)

CysSH

+

CysS

3

(6)

CysSR

+

CysS

3

(7)

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(8) For the reaction of disulfides with small aliphatic radicals (i.e., R = CH3 ), theoretical calculations predict that a homolytic substitution such as reaction 7 proceeds like a nucleophilic substitution, SN2, with preferential attack of R from the back side of the disulfide bond and without intermediacy of a sulfuranyl radical [12]. However, if R is a strongly oxidizing radical, i.e. the hydroxyl radical (HO ), homolytic substitution at the disulfide bond is not the only pathway. Pulse radiolysis studies coupled to time-resolved UVand conductivity detection have shown that the reaction of HO radicals with small organic disulfides follows two parallel pathways, homolytic substitution (analogous to reaction 7) and one-electron oxidation, generating disulfide radical cations (reaction 8) [17,18]. In addition, especially for the reaction of HO (and other strongly oxidizing radicals) with cystine, there is the opportunity for hydrogen abstraction from the cystine C-H bond, followed by elimination, yielding dehydroalanine and the Cys perthiyl radical 4, CysSS (reactions 9 and 10) [19]. Cys thiyl radicals are also generated photolytically through the homolytic cleavage of Cys disulfides [20]. Experimental measurements of radical yields during the photolysis of glutathione disulfide indicate negligible formation of Cys perthiyl radicals (< 2%), but when photolysis is carried out with disulfides of -alkyl substituted Cys (e.g., penicillamine disulfide) significant yields of perthiyl radicals are generated [21].

3. Reactions of Thiyl Radicals Thiyl radicals are moderately good one-electron oxidants, which react under electron transfer with a series of biologically relevant electron donors. For example, the reaction of thiyl radicals with ascorbate (AH ) proceeds with k11 = 6x108 M-1s-1 2. Especially the field of radiation biology considers this process an essential part of the radioprotection by thiols [10]: radiation exposure of tissue generates carbon-centered radicals, which would only relatively slowly react with ascorbate. However, these carbon-centered radicals efficiently react with thiols to generate thiyl radicals, which, in turn, are converted back to thiols through reaction with ascorbate. In this process, thiols essentially serve as catalysts for the reduction of weakly oxidizing organic radicals by ascorbate.

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Christian Schöneich

CysS•

+

AH

CysS

+

A• +

H+

(11)

Thiyl radicals efficiently complex with thiolate to form disulfide radical anions (reaction 12). These disulfide radical anions have reducing properties and react efficiently with oxygen to yield superoxide radical anion (reaction 13) [2].

[CysS-SR] +

O2

Cys-SR +

O2

(13)

Hence, the association with thiolate converts the oxidizing thiyl radical into a reducing radical complex, where the relative yields of oxidizing and reducing species are essentially a function of concentration and pKa of the available thiol/thiolate, the pH of the medium, and K12.

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Thiyl radicals reversibly add to the double bonds of unsaturated fatty acids (reaction 14), generating an intermediary -alkylthio-substituted radical, which can ultimately result in cistrans-isomerization [5]. Note that for isolated double bonds, thiyl radical elimination from the -alkylthio-substituted radical 5 is very fast, ensuring that the equilibria in reaction 14 are normally located on the side of the reactants. Within membrane structures such thiyl radicalinduced cis-trans-isomerization can only proceed for thiyl radicals, which can enter the lipophilic membrane layer; one biological candidate for such a species would be HS [23]. The HS radical possesses a pKa = 6.9 for equilibrium 15, so that a significant fraction of HS exists in the protonated state at physiological pH.

HS can be generated through oxidation of H2S, which can be enzymatically produced from homocysteine through cystathionine -synthase (CBS) and cystathionine -lyase (CSE), and, chemically, through -elimination from Cys. The exposure of tissue to ionizing radiation may lead to another biologically relevant lipophilic thiyl radical, CH3S , which can be generated through the action of H on free or protein-bound Met (reactions 16-18) [24]. Here, the addition of H to the Met sulfur generates the intermediary sulfuranyl radical 6. In reaction sequence 17-18, the sulfuranyl radical is shown to decompose via cleavage of the Met C-S bond, generating CH3SH, which subsequently transfers an H -atom to the carboncentered radical, yielding CH3S [in well oxygenated solution, oxygen addition to the carboncentered radical would compete with such H -atom transfer, ultimately generating homoserine]. Of course, any Cys thiyl radical generated on a membrane protein may access unsaturated fatty acids, creating the potential for protein-bound thiyl radicals to induce cistrans-isomerization of unsaturated fatty acids.

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Thiyl radicals will react with polyunsaturated fatty acids via cis-trans-isomerization, as described below, and via H -atom abstraction from bis-allylic methylene groups (reaction 19) [25]. The rate-constants for the latter increase with the number of bis-allylic methylene groups in the fatty acid molecule. As for cis-trans-isomerization, such H -atom abstraction from membrane-bound fatty acids would require the presence of a lipophilic thiyl radical such as HS or CH3S .

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Much emphasis has also been placed on hydrogen abstraction reactions by thiyl radicals from carbohydrates and amino acids. Initial radiation chemical studies provided evidence for the reversibility of H -atom transfer (reaction 20) between radicals from model alcohols and ethers [26,27] as models for 2-deoxyribose, the building block of nucleotides. Subsequently, a kinetic NMR method was employed to measure rate constants for the H -atom transfer from various carbohydrates to thiyl radicals [28]. This reaction is not only of potential biological concern during oxidative stress, but represents an important step in the enzymatic generation of 2-deoxyribose for nucleotide synthesis by ribonucleotide reductases [1]. In general, equilibrium 20 is located on the left hand side with k20 103-104 M-1s-1 and k-20= 106-108 M-1s1 , depending on the nature of the thiyl radical, alcohol, ether and/or carbohydrate.

Especially, the C-H bonds of peptides are susceptible to H -atom abstraction by thiyl radicals (shown for N-acetyl amino acid amides in reaction 21) due to the captodative stabilization of the generated carbon-centered radicals. Rate constants for the intermolecular reaction of thiyl radicals with N-acetyl amino acid amides (and diketopiperazines) are in the order of k21 = 103-105 M-1s-1 (rate constants for the reverse reaction, k-21 have not been obtained) [29], while the anionic forms of amino acids, i.e. structures in which the free amino group provides particular good stabilization of the forming carbon-centered radical, react with k22> 105 M-1s-1 [30]. The resulting -amino type radical is a reducing species, which would reduce molecular oxygen to superoxide.

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A reversible intramolecular H•-atom transfer process, analogous to that of reaction 22, has been observed for thiyl radicals of glutathione (Vol. II, Chapter 1), which react with the C-H bond of the N-terminal -Glu residue, again generating an -amino-type radical (reaction 23) [2,30]. The location of equilibrium 23 depends on the protonation state of the N-

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terminal -Glu residue, where the N-terminal -amino-type radical would again have the opportunity to react withmolecular oxygen. Hence, through equilibrium 23, glutathione thiyl radicals may generate superoxide.

Pulse radiolysis and laser flash studies have provided kinetic information for the intramolecular reversible H -atom transfer between Cys thiyl radicals and C-H bonds in various model peptides (reaction 24), where k24 and k-24 depend on the nature of the amino acid (AA) target within the model peptides [31]. For example, for AA = Gly, k24 105 s-1 and k-24 106 s-1 in the model peptides N-Ac-Cys-(Gly)6 and N-Ac-Cys-(Gly)2Asp(Gly)3, and for AA = Ala, k24 104 s-1 and k-24 105 s-1 for the model peptide N-Ac-Cys-(Ala)2Asp(Ala)3. Hence, the equilibrium constant K24 0.1 for both N-Ac-Cys-(Gly)2Asp(Gly)3 and N-Ac-Cys(Ala)2Asp(Ala)3,which would be consistent with comparable calculated C-H bond energies for Gly and Ala [32]. On the other hand, the absolute values of the rate constants for both peptides vary by approximately one order of magnitude. Further experimental evidence for

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H -atom transfer between Cys thiyl radicals and amino acid residues in model peptides as well as a protein (insulin) come from covalent H/D-exchange studies, leading to the incorporation of deuterium into original C-H bonds of the target amino acids when experiments are carried out in D2O (reactions 25-27) [33-35]. Here, dissolution of a starting disulfide-containing peptide in D2O first leads to H/D exchange of exchangeable protons such as those of the peptide bonds, amides, hydroxyl, carboxyl and amino groups. Photolysis of the disulfide leads to thiyl radicals, which abstract an H -atom from a nearby amino acid, yielding thiol and a carbon-centered radical. The sulfhydryl group undergoes H/D exchange and transfers a D -atom back to the intermediary carbon-centered radical, leading to covalent Dincorporation, which, in the final products, can be detected and quantitated by mass spectrometry. For peptides containing amino acids different from Gly, these reversible H atom transfer reactions also provide the possibility for epimerization, i.e. conversion of Linto D-amino acids, and such a reaction was experimentally demonstrated for Ala [35]. Especially the L- to D-amino acid conversion, but also the formation of protein carboncentered radicals, which can react with molecular oxygen, represent potential pathways to protein damage initiated by thiyl radicals [4] (see also Chapter 6). Such reactions must, therefore, be taken into account when evaluating thiols as protein targets for free radicals and reactive oxygen and nitrogen species: the oxidation of a protein thiol may not necessarily mandate that the ultimate damage to the protein is located at this specific thiol residue, but the intermediary thiyl radical may catalyze protein damage elsewhere.

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Thiyl radicals react reversibly with molecular oxygen (reaction 28) [2,36], leading to thiyl peroxyl radicals, where k28 = 2.2x109 M-1s-1 and k-28 = 6.2x105 s-1 . Because of the rapid dissociation of thiyl peroxyl radicals (reaction 28), these species do not represent an efficient sink for thiyl radicals in biological environment. Thiyl peroxyl radicals appear to be weaker one-electron oxidants than thiyl radicals, and undergo a relatively slow intramolecular rearrangement to sulfonyl radicals, RSO2 (k 2x 103s-1) [36] (reaction 29). Because of the overall inefficient reaction with molecular oxygen, and the slow rearrangement to sulfonyl radicals, physiological tissue concentrations of oxygen are not expected to compete efficiently with intramolecular H -atom transfer processes such as shown in reactions 24 and 25. A more efficient competition would be complexation with thiolate (reaction 12) or reaction with ascorbate (reaction 11) if these reactants can access a thiyl radical, for example in the interior of a protein. Whether a thiyl radical will be able to initiate protein damage will, therefore, depend on the location of the thiyl radical within the three-dimensional structure of a protein, defining accessibility and the proximity of potential target C-H bonds for H -atom abstraction.

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4. Reactions of Perthiyl Radicals There is considerably less information available on the reactions of perthiyl radicals as compared to thiyl radicals. Perthiyl radicals react under electron transfer with ascorbate, albeit with a significantly lower rate constant (k30 = 4.1x106 M-1s-1) [21] and add molecular oxygen (k31 = 5.1x106 M-1s-1) [21]. RSS•

+

AH

RSS•

+

O2 → RSSOO•

→ RSS

+

A• +

H+

(30) (31)

They are involved in reversible H•-atom transfer reactions with activated C-H bonds, e.g., of 2-propanol (reaction 32). However, equilibrium 32 is located far more to the left hand side as compared with the analogous equilibrium of thiyl radicals, i.e K32< K20 (for 2-propanol).

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5. Formation of Met Radical Cations Met radical cations are generated through one-electron oxidation of Met by strong oxidants such as, for example, HO•. The detailed mechanism involves addition of HO• to the sulfur, generating an initial hydroxyl sulfuranyl radical 7 (reaction33) [38,39]. HO

+

Met

Met(>S -OH) 7

(33)

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The hydroxyl sulfuranyl radical 7 can decompose via various pathways, of which here only the biologically relevant pathways are discussed. When Met is part of a peptide sequence, i.e. does not contain free amino and carboxyl groups, there are three documented pathways leading to sulfide radical cations [39,40], the unimolecular dissociation of 7 into radical cation 8 and HO (reaction 34), the elimination of HO assisted through formation of an (S O)-three-electron bonded radical cation (reaction 35), and an intramolecular protontransfer from the N-terminal amide bond leading to the elimination of water under formation of an (S N)-three-electron-bonded radical (reaction 36). Evidence for reaction 35 has been obtained through time-resolved pulse radiolysis studies with N-acetyl Met amide [39], several linear Met-containing peptides [39], and model compounds, in which the Met structure was incorporated into a rigid norbornane framework such as, for example, structures 9a and 10 [41].

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Here, the formation of (S O)-three-electron-bonded radical cations is supported by timeresolved conductivity measurements in addition to time-resolved UV-spectroscopy (in contrast, no spectral evidence for an (S O)-three-electron bond was obtained for structure 9b, in which the amide ligand is in exo-orientation to the thioether). On the other hand, evidence for reaction 36 comes from pulse radiolysis of Met incorporated into a diketopiperazine system, i.e. cyclo-Met-Met (structure 11) [40]. Moreover, pulse radiolysis data on the HO•-radical-dependent oxidation of Ca2+-calmodulin show transient absorption spectra which closer resemble those of (S N)- rather than (S O)-three-electron bonded radicals [42]. In contrast to peptide-bound Met, the reaction of HO -radicals with free Met nearly exclusively yields a short-lived (S N)-bonded intermediate [38], which subsequently decarboxylates. In oxygenated aqueous solution, the resulting -amino radicals convert to 3methylthiopropionaldehyde (methional) [24]. At this point, it is not clear which parameters would control whether reaction 35 or 36 prevails. However, the general tendency to stabilize sulfide radical cations through complexation with electron-rich functional groups can provide a kinetic and/or thermodynamic driving force for the formation of such radical cations. This may be highly relevant for the oxidation of -amyloid peptide ( -AP) through Cu(II), leading to intermediary Cu(I), and, ultimately, the formation of H2O2[43]. Based on studies with several full length and truncated -AP peptides, it was suggested that the electron donor in -AP is Met-35[44]. The most recently measured reduction potential for Cu(II)- AP is Eo‟ = 0.28 V vs. NHE [45], i.e. a value considerably less positive than that for free Met, for which a peak potential of Ep 1.5-1.7 V vs. NHE has been measured for irreversible electrochemical oxidation [46]. Hence, from a comparison of these values it appears that an electron-transfer from Met to Cu(II) in -AP is highly endothermic. On the other hand, any

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means to significantly lower the peak potential for Met oxidation would enhance the potential for such an electron transfer. The complexation of a sulfide radical cation with an amide bond provides such an approach, which, for example, for structure 9a lowers Ep by 0.33V compared to structure 9b (which has no opportunity to form an intramolecularly (S O)bonded radical cation) [41]. It remains to be shown whether incorporation of Met into specific peptide sequences will allow for a further reduction of Ep. These electrochemical measurements stand in some contrast to theoretical calculations, which predicted [47] that such a reduction of Ep could not be achieved.

6. Reactions of Met Radical Cations Methionine sulfide radical cations are strong one-electron oxidants, which are expected to oxidize most biologically relevant one-electron donors. Here, we will focus on a few reactions with the potential to cause irreversibly protein damage. Met sulfide radical cations deprotonate to yield -(alkylthio)alkyl radicals (reaction 37) [39], which can add oxygen to generate peroxyl radicals. However, especially for -(alkylthio)alkyl radicals generated from cyclo-Met-Met (structure 11), a decomposition process was observed, which depended linearly on the concentration of H+ [40]. Together with parallel conductivity data, this feature was rationalized by a potential formation of amide radical cations, which subsequently underwent hydrolysis, electron transfer and decarboxylation. Only the latter part of this still speculative, but intriguing, pathway is displayed in reactions 38-40. Also the decomposition of (S O)-bonded radical cations of structure 9a could not be completely accounted for through well established reactions such as deprotonation (analogous to reaction 37) [41]. This feature was rationalized by a potential addition of water, followed by elimination of NH3 and CO2. Overall, reactions such as 37-40 have the potential to cause irreversible damage to Metcontaining sequences in peptides and proteins through initial one-electron oxidation of Met. The ensuing products are not amenable to repair (unlike the common two-electron oxidation

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product Met sulfoxide, which can be reduced back to Met by methionine sulfoxide reductases).

Conclusions The current Chapter focuses on the formation and reactions of radicals from Cys and Met, and summarizes only those mechanisms which may be of relevance under physiological conditions. Additional work must be performed on the potential protein damage inflicted by thiyl radicals and/or the potential hydrolysis reactions of Met-derived radical cations. Ultimately, these reactions may lead to protein fragmentation, a feature frequently observed under conditions of oxidative stress.

Acknowledgment Financial support provided by the NIH (PO1AG12993) is gratefully acknowledged.

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[10] [11]

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[12] [13]

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[17]

[18]

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[19] Elliot AJ, McEachern RJ, Armstrong DA Oxidation of amino-containing disulfides by Br2-• and •OH. A pulse radiolysis study. J Phys Chem 1981; 85: 68-75. [20] Kolano C, Helbing J, Bucher G, Sander W, Hamm P.Intramolecular disulfide bridges as a phototrigger to monitor the dynamics of small cyclic peptides. J Phys Chem B 2007; 111:11297-11302. [21] Everett SA, Schöneich C, Stewart JH, Asmus K-D. Perthiyl radicals, trisulfide radical ions, and sulfate formation. A combined photolysis and radiolysis study on redox processes with organic di- and trisulfides. J Phys Chem 1992; 96: 306-314. [22] Chatgilialoglu C, Altieri A, Fischer H. The kinetics of thiyl radical-induced reactions of monounsaturated fatty acid esters. J Am Chem Soc 2002; 124:12816-12823. [23] Lykakis IN, Ferreri C, Chatgilialoglu C.The sulfhydryl radical (HS•/S•-): a contender for the isomerization of double bonds in membrane lipids. Angew Chem Int Ed Engl 2007; 46: 1914-1916. [24] Barata-Vallejo S, Ferreri C, Postigo A, Chatgilialoglu C. Radiation chemical studies of methionine in aqueous solution: understanding the role of molecular oxygen. Chem Res Toxicol 2010; 23: 258-263. [25] Schöneich C, Dillinger U, von Bruchhausen F, Asmus KD. Oxidation of polyunsaturated fatty acids and lipids through thiyl and sulfonyl radicals: reaction kinetics, and influence of oxygen and structure of thiyl radicals. Arch Biochem Biophys 1992; 292: 456-467. [26] Akhlaq MS, Schuchmann HP, von Sonntag C The reverse of the 'repair' reaction of thiols: H-abstraction at carbon by thiyl radicals. Int J Radiat Biol Relat Stud Phys Chem Med 1987; 51: 91-102. [27] Schöneich C, Bonifačić M, Asmus K-D. Determination of absolute rate constants for the reversible hydrogen-atom transfer between thiyl radicals and alcohols or ethers. J Chem Soc Faraday Trans 1995; 91: 1923-1930. [28] Pogocki D, Schöneich C Thiyl radicals abstract hydrogen atoms from carbohydrates: reactivity and selectivity. Free Radic Biol Med 2001;31: 98-107. [29] Nauser T, Schöneich C Thiyl radicals abstract hydrogen atoms from the (alpha)C-H bonds in model peptides: absolute rate constants and effect of amino acid structure. J Am Chem Soc 2003;125: 2042-2043. [30] Zhao R, Lind J, Merenyi G, Eriksen TE. Kinetics of one-electron oxidation of thiols and hydrogen abstraction by thiyl radicals from -amino C-H bonds. J Am Chem Soc 1994; 116: 12010-12015. [31] Nauser T, Casi G, Koppenol WH, Schöneich C Reversible intramolecular hydrogen transfer between cysteine thiyl radicals and glycine and alanine in model peptides: absolute rate constants derived from pulse radiolysis and laser flash photolysis. J Phys Chem B 2008;112: 15034-15044. [32] Rauk A, Yu D, Taylor J, Shustov GV, Block DA, Armstrong DA. Effects of structure on alpha C-H bond enthalpies of amino acid residues: relevance to H transfers in enzyme mechanisms and in protein oxidation. Biochemistry 1999; 38: 9089-9096. [33] Mozziconacci O, Sharov V, Williams TD, Kerwin BA, Schöneich C. Peptide cysteine thiyl radicals abstract hydrogen atoms from surrounding amino acids: the photolysis of a cystine containing model peptide. J Phys Chem B 2008; 112: 92509257.

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[34] Mozziconacci O, Williams TD, Kerwin BA, Schöneich C. Reversible intramolecular hydrogen transfer between protein cysteine thiyl radicals and alpha C-H bonds in insulin: control of selectivity by secondary structure. J Phys Chem B 2008; 112: 15921-15932. [35] Mozziconacci O, Kerwin BA, Schöneich C Reversible hydrogen transfer between cysteine thiyl radical and glycine and alanine in model peptides: covalent H/D exchange, radical-radical reactions, and L- to D-Ala conversion. J Phys Chem B 2010; 114: 6751-6762. [36] Zhang X, Zhang N, Schuchmann H-P, von Sonntag C. Pulse radiolysis of 2mercaptoethanol in oxygenated aqueous solution. Generation and reactions of the thiylperoxyl radical. J Phys Chem 1994; 98: 6541-6547. [37] Everett SA, Folkes LK, Wardman P, Asmus KD Free-radical repair by a novel perthiol: reversible hydrogen transfer and perthiyl radical formation. Free Radic Res 1994; 20: 387-400. [38] Hiller K-O, Masloch B, Göbl M, Asmus K-D. Mechanism of the OH• radical induced oxidation of methionine in aqueous solution. J Am Chem Soc 1981; 103: 2734-2743. [39] Schöneich C, Pogocki D, Hug GL, Bobrowski K. Free radical reactions of methionine in peptides: mechanisms relevant to beta-amyloid oxidation and Alzheimer's disease. J Am Chem Soc 2003;125: 13700-13713. [40] Bobrowski K, Hug GL, Pogocki D, Marciniak B, Schöneich C Stabilization of sulfide radical cations through complexation with the peptide bond: mechanisms relevant to oxidation of proteins containing multiple methionine residues. J Phys Chem B 2007; 111: 9608-9620. [41] Glass RS, Hug GL, Schöneich C, Wilson GS, Kuznetsova L, Lee TM, Ammam M, Lorance E, Nauser T, Nichol GS, Yamamoto T. Neighboring amide participation in thioether oxidation: relevance to biological oxidation. J Am Chem Soc 2009; 131: 13791-13805. [42] Nauser T, Jacoby M, Koppenol WH, Squier TC, Schöneich C. Calmodulin methionine residues are targets for one-electron oxidation by hydroxyl radicals: formation of S N three-electron bonded radical complexes. Chem Commun 2005; 587-589. [43] Hewitt N, Rauk A. Mechanism of hydrogen peroxide production by copper-bound amyloid beta peptide: a theoretical study. J Phys Chem B 2009; 113: 1202-1209. [44] Butterfield DA, Boyd-Kimball D. The critical role of methionine 35 in Alzheimer's amyloid beta-peptide (1-42)-induced oxidative stress and neurotoxicity. Biochim Biophys Acta 2005; 1703: 149-156. [45] Jiang D, Men L, Wang J, Zhang Y, Chickenyen S, Wang Y, Zhou F. Redox reactions of copper complexes formed with different beta-amyloid peptides and their neuropathological relevance. Biochemistry 2007; 46: 9270-9282. [46] Sanaullah, Wilson S, Glass RS The effect of pH and complexation of amino acid functionality on the redox chemistry of methionine and X-ray structure of [Co(en)2(L-Met)](ClO4)2•H2O. J Inorg Biochem 1994; 55: 87-99. [47] Brunelle P, Rauk A. One-electron oxidation of methionine in peptide environments: the effect of three-electron bonding on the reduction potential of the radical cation.J Phys Chem A 2004; 108: 11032–11041.

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Chapter 5

Redox-Active Metals: Iron and Copper Willem H. Koppenol* and Patricia L. Bounds Institute of Inorganic Chemistry, Department of Chemistry and Applied Biosciences Eidgenössische Technische Hochschule, CH-8093 Zürich, Switzerland

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1. Introduction Dioxygen† [1, 2] (O2) is simultaneously necessary for life and a source of potential harm [3, 4]. Harmful reactions, in which partially reduced oxygen species (PROS)‡ such as superoxide (O2 ) and hydrogen peroxide (H2O2) may be formed, are, in part, mediated by redox-active metals, the most physiologically relevant of which are iron and copper. These metals are essential for the transport, storage, and activation of O2, and for electron transfer, e.g., during respiration (see also Vol. II, Chapters 19-20). Along the reductive pathway of O2 to water, partially reduced and potentially reactive intermediates are sequentially made and managed by metalloproteins [5]. Although the redox reactivity of the metals in these proteins *

Email: [email protected] Nomenclature. Formula, systematic name and still allowed trivial name in italics: O 2, dioxygen, oxygen; O2 , dioxide(•1 ) or dioxidanidyl, superoxide; HO2 , hydridodioxygen(•), dioxidanyl, or hydrogen dioxide (hydroperoxyl or perhydroxyl are obsolete); H2O2, dioxidane, hydrogen peroxide; HO , hydridooxygen(•) or oxidanyl, hydroxyl; O , oxide(• ) or oxidanidyl; OCl , oxidochlorate(1 ), hypochlorite; HOCl, hydroxidochlorine, hypochlorous acid; NO , oxidonitrogen(•) or nitrogen monoxide (nitric oxide is obsolete); ONOO , oxidodioxidonitrate(1 ), peroxynitrite; ONOOH, hydroperoxidooxidonitrogen, peroxynitrous acid; Fe3+ or Fe(III), iron(3+) or iron(III) (ferric is obsolete); Fe2+ or Fe(II), iron(2+) or iron(II) (ferrous is obsolete); Cu+ or Cu(I), copper(+) or copper(I) (cuprous is obsolete); Cu2+ or Cu(II), copper(2+) or copper(II) (cupric is obsolete); FeO2+, oxidoiron(2+) or oxidoiron(IV) (ferryl is obsolete)[1,2]. The abbreviations edta, dtpa and nta, and atp refer to the metal chelators ethylenediaminetetraacetate, diethylenetriaminepentaacetate, nitrilotriacetate and adenosinetriposphate, respectively; these abbreviations are not capitalised.[2] NTBI is non-transferrinbound iron; cp20 is 3-hydroxy-1,2-dimethylpyridin-4(1H)-one, and icl670, 4-[3,5-bis(2hydroxyphenyl)-1H-1,2,4-triazol-1-yl]benzoic acid. ‡ We use the acronym PROS rather than ROS (for “reactive oxygen species”); PROS is not generally meant to include O2, which, strictly speaking, is also a reactive species. Analogously, we apply the acronym PONS for “partially oxidized nitrogen species” instead of RNS for “reactive nitrogen species”. †

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is harnessed and directed by the protein and/or co-factor to which it is bound, there can be some non-productive release of O2 or H2O2. One approach that nature uses to circumvent non-productive reactions of oxygen is to employ more than one redox-active metal ion, sometimes stabilized by one or more non-redox-active metal ions, for the transfer of multiple electrons over a short time frame, e.g. a copper and an iron ion for the reduction of O2 by cytochrome c oxidase [6], and four manganese ions for the oxidation of water in photosystem II [7]. Even so, oxyhemoglobin [8, 9] and oxymyoglobin [10] reportedly release O2 , and xanthine oxidase [11], and cytochromes P450 [12] are known to produce H2O2. Another source of O2 in vivo is the reduction of O2 by electrons that leak from the mitochondrial electron transport chain; in fact, 1 – 4% of all O2 consumed during respiration may be reduced to O2 in this fashion [13, 14] (Vol. II, Chapter 15). It is now well established that PROSs, including peroxynitrite (ONOO ), play roles in a stunning array of diseases [15, 16]. One function of the protein environment of a metalloprotein is to modulate the reactivity of the metal ion; the potential for harmful reactivity likely to be catalyzed by free metal ions such as iron and copper outside of the protein milieu is the topic of this review. In vivo, excess free metal ions are sequestered in specialized storage proteins ― iron in ferritin [17] (Vol. II, Chapter 19) and copper possibly in metallothioneins (Vol. II, Chapter 20), while circulating iron is captured by transferrin [18] and copper by ceruloplasmin [19]. However, transfer of metals from storage to functional metalloproteins and the turnover of these proteins likely gives rise to low concentrations of low-molecular weight metal ion complexes [20, 21]; this pool of iron and copper may increase as a consequence of oxidative stress.

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2. Redox Active Iron and Copper In Vivo The role played by iron and copper in diseases of ageing and oxidative stress has been recently reviewed [22]. Both iron and copper ions “redox cycle”, i.e. shuttle between two oxidation states via transfer of a single electron (Figure 1), as part of their normal functions in vivo. Redox cycling is also widely believed to be involved in the generation of damaging PROS; “free” iron and copper, actually free in solution or non-specifically bound to albumin or lowmolecular-weight ligands, are invoked as being particularly damaging. However, the link between pools of free transition metals, PROS, and disease is tenuous, and the evidence in the literature is largely circumstantial. Further, the speciation, that is the ligands, of “free” iron and copper ions involved in disease states have not been precisely identified. Diseases in which excess iron plays a role include hemochromatosis and iron-loading anemias, such as thalassemia, sickle cell disease and myelodysplasia [23] (Vol. III, Chapter 2). When normal mechanisms of iron intake and elimination become imbalanced, the binding capacity of ferritin and transferrin are exceeded, and pools of free, or non-transferrin-bound, iron (NTBI) increase (Vol. II, Chapter 19). The NTBI, which has been detected in the serum of hemochromatosis [24], sickle cell anemia and thalassemia patients [25], may consist in part of iron(III)-citrate complexes [26]. Although citrate has been identified as a ligand in NTBI, the exact composition of the complex or complexes existing in vivo remain elusive [27]. Citrate is present in the serum at a concentration of 0.1 mM, and the clinical range of NTBI concentrations varies between 0 to 10 M [28]. The aqueous speciation of iron(III) citrate is highly dependent on the iron/citrate molar ratio and, under the clinical conditions described,

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several complexes have been shown to be possible [29]. The mononuclear Fe(cit)25 complex predominates over the clinically observed iron/citrate molar ratios, and, additionally, the multinuclear complexes Fe3(cit)33 and Fe3(cit)47 become relevant at higher NTBI concentrations [30]. Thus, the term NTBI may encompass several different species, of which one or more could be redox-active. Altered concentrations of copper, zinc and iron ions in the brain are a feature of Alzheimer‟s disease, and copper is widely thought to contribute to the oxidative stress model of Alzheimer pathophysiology [31] (see also Vol. III, Chapter 10). Copper has been shown to bind tightly to amyloid- peptide (K = 10–11 M), and the copper-amyloid- complex oxidizes cholesterol to cytotoxic products [32, 33]. Copper also plays a role in Parkinson‟s disease, atherosclerosis, diabetes, and other age-related diseases, as reviewed by Brewer [22].

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3. Biological Oxidants Reactive species formed in vivo that are thought to be physiologically relevant include O2 , nitrogen monoxide (NO ), the hydroxyl radical (HO ), the trioxocarbonyl radical (CO3 ), nitrogen dioxide (NO2 ), H2O2, ONOO , and hypochlorite (OCl ) (see also Chapters 2-4). The first four species in the list each possess an unpaired electron and are, therefore, called radicals; H2O2, ONOO and OCl have no unpaired electrons and are, therefore, not radicals§. It is important to recognise that radical species are also functionally produced in vivo, e.g., the nitric oxide synthases generate NO , a diatomic radical species that plays vital roles in neuronal cell communication, endothelial vasodilation, and immune response [34, 35]. Of all the biological oxidants listed above, HO is the most destructive: HO is a promiscuous oxidant that reacts with small organic molecules, proteins, nucleic acids, and lipids by abstraction of hydrogen or addition to a double bond to form a radical species that can undergo further transformations. The HO has a high positive electrode potential|| (E° (HO ,H+/H2O) = +2.31 V at pH 7) [36], and the second-order rate constants (kobs = 109 – 1010 M 1s 1) [37] are in a range that indicates diffusion-controlled reactions. In contrast, H2O2 does not react directly with most organic compounds, and any toxicity attributed to H2O2 §

||

The term “radical”, which was used in the past to refer to a group that is part of an organic molecule, e.g., “methyl radical,” has been replaced with substituent; in former usage, the term “free radical” referred to an unattached group, usually containing an unpaired electron. In current usage, the word “radical” indicates that the group possesses an unpaired electron, and the word “free” is no longer included. The values given are relative to the normal hydrogen electrode, a platinum electrode in a solution that is 1 molal in H+ (1 gram H+ per kg of water, essentially the same as 1 molar) and in equilibrium with an atmosphere of 100 kPa of H2, that, by definition, has a value of 0 V. If we use O 2 and O2 as an example, the electrode potential of the couple O2/O2 refers to the reaction: O2 + e O2 , with a value of 0.35 V, relative to the normal hydrogen electrode. The electrode potential was formerly called a reduction potential. This value would be measured if one could connect a stable solution of O 2 (1 molal) in equilibrium with an atmosphere of O2 (100 kPa) with the normal hydrogen electrode by way of two platinum electrodes connected to a volt meter, and a salt bridge between the solutions to allow ions to flow between the solutions. Electrons would flow from the negatively charged electrode in the O2/O2 solution – producing O2 – to the positive electrode in the H+/H2 solution where H2 would be formed. It should be clear, though, that a stable solution of O2 does not exist; instead, the electrode potential is measured by indirect means. An oxidation potential refers to the reaction O2 O2 + e and the sign is reversed: +0.35 V.

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would depend on the presence of redox-active transition metal ions, iron and copper, the topic of this Chapter. It is known that NO , O2 , and H2O2 are produced enzymatically, and the remaining species listed are formed from these (Figure 2). In phagocytic cells, i.e., neutrophils and macrophages, O2 generated by NADPH oxidase reacts to produce H2O2 (Reaction 1) and ONOO (Reaction 2) [38]. Also in neutrophils, H2O2 is a substrate in the reaction of myeloperoxidase to generate OCl (Reaction 3). The reactive ONOO and OCl synthesized in phagocytic cells are part of the host organism‟s defence arsenal to combat pathogenic organisms [39] and destroy foreign objects [40]. Reaction 4, the reaction of ONOO with carbon dioxide (CO2), yields mostly NO3 and CO2, but also the oxidizing radicals CO3 and NO2 [41, 42]. 2 O2 + 2H+

O2 + H2O2

(1)

O2 + NO

ONOO

(2)

H2O2 + Cl

OCl + H2O

(3)

ONOO + CO2

CO3 + NO2

(4)

Hasc–

asc – + H+

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NTBI-Fe3+

NTBI-Fe2+

H2O2 + H+

H2O + HO Figure 1. Redox cycling by iron.

+

L-Arginine + 1.5 (NADPH + H ) + 2 O2

PROS

+

L-Citrulline + 1.5 (NADP ) + 2 H2O

NOS

CO2

NO

CO3 – + NO2

ONOOCO2–

ONOO–

CO2 + NO3–

NADPH + e– NADP– + H+

O2

NOX

O2

–

HO2 H

H2O2

SOD

+

O2

–

O2

Figure 2. Generation and interconnectedness of PROS.

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OCl–

MPO

Cl–

H2 O

Redox-Active Metals: Iron and Copper

95

The electrode potential of O2 , E°(O2/O2 ) = –(0.18 + 0.02) V, or –(0.35 + 0.02) V (pO2 = 0.100 Mpa) indicates that O2 is only mildly reducing; O2 is, indeed, not very damaging [43]. Reaction 1, the disproportionation of O2 is catalyzed by superoxide dismutases [44] (SODs; see Vol. II, Chapter 5). At pH 7, O2 is 0.5% protonated (HO2 ), and the protonated form also acts as an oxidant: E° (HO2 ,H+/H2O2) = 1.04 V [45]. The electrode potentials indicate that the oxidation of O2 – by HO2 is thermodynamically likely, and the reaction indeed proceeds rapidly (k = 1.0 • 108 M 1s 1) [46] even in the absence of SOD. The importance of NO as a biological signaling molecule [47] has emerged in the past two decades. In our view, NO itself is not damaging per se aside from its affinity for iron(II) which may lead to inhibition of hemoproteins with an open coordination site. The very rapid reactions of NO with oxymyoglobin and oxyhemoglobin (k = 4.4 • 107 M 1s 1 and 8.9 • 107 M 1s 1, respectively, relative to heme concentration) produce the corresponding metmyoglobin and methemoglobin (and nitrate) [48] in which the heme iron is oxidized and cannot bind O2. In theory, NO2 can be formed via Reaction 5, the autoxidation of NO ; this reaction, however, requires two NO and one O2 [49-53], and the mechanism is complex [54]. Reaction 5 is likely to be slow (hours to days) in vivo, given the estimated physiological concentrations of NO and O2 [55]. In contrast, NO diffuses from tissue to the nearest red blood cell, where it is scavenged by hemoglobin more rapidly than it can react with O2 [56]. Thus, the lifetime of NO in vivo is determined by reactions with O2 and hemoglobin, and Reaction 5 can be neglected.

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2 NO + O2

2NO2

(5)

The reaction of NO with O2 to form ONOO– (Reaction 2) is diffusion-controlled (k = 1.6 • 1010 M 1s 1) [57], and ONOO– is recognized as an important biological oxidant. At physiological pH, ONOO– is largely protonated (pKa = 6.8) [58]; ONOOH reacts much more slowly and selectively than HO [59] with biological targets. Evidence for formation of ONOOH in vivo is largely based on immunodetection of nitrotyrosine residues [60]; it should, however be noted that nitrotyrosine may also be formed by peroxidase-catalyzed reactions of nitrite and H2O2. Tyrosine nitration blocks phosphorylation and, thus, impacts on the signaling functions of tyrosine residues [61]. Furthermore, nitrotyrosine can, at least in vitro, accept an electron from ascorbate and then reduce O2 [62], thereby contributing to the formation of O2 –. The main reaction of ONOOH is isomerisation to NO3–, but it is widely believed that ONOOH undergoes homolysis to at least some extent to yield NO2 and HO [63-65]. Estimates of the yield of homolysis range from as low as ca. 1% [66] to as high as ca. 40% [67]. We have sought but failed to find evidence to support a mechanism in which homolysis accounts for more than a very low percentage of the decay of ONOOH [68-72]. The radicals produced in the reaction of ONOO– with CO2 [73], NO2 and CO3 [41, 42] are strongly oxidizing species: (E° (NO2 /NO2 ) = +1.04 V) [74] and (E° (CO3 /CO32 ) = +1.58 V) [75, 76]. These radicals have been implicated in ONOO–-mediated nucleic acid oxidation [77] and tyrosine nitration [78]. Finally, the OCl– produced by myeloperoxidase in activated phagocytic cells reacts with many biomolecules, including nucleotides, DNA and low-molecular-weight thiols [79], as well as protein thiol [80] and amino residues [81].

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4. Redox Chemistry and Biochemistry of Iron: The Historical Context The involvement of transition metal ions in oxidative stress pathologies is often attributed to “Fenton chemistry”, and the Fenton reaction, reduction of H2O2 by iron(II) to generate HO (Reaction 6), has long been of interest to the free radical community. H. J. H. Fenton discovered the reaction that bears his name serendipitously [82]. While still an undergraduate student, Fenton was shown a violet coloured solution that had been obtained by a fellow student who was mixing reagents, including tartaric acid, at random, and later reproduced the reaction and published it as a test for tartaric acid in 1876 [83]. A full account, including the structure of the oxidized product, 2,3-dihydroxymaleic acid, was published in 1896 [84]. Although Fenton used iron(II) and H2O2 to modify organic compounds, he was not aware that HO• is formed in the reaction. Two years after Fenton‟s death, Fritz Haber and Richard M. Willstätter [85] published a paper on radical chain reactions in organic chemistry and biochemistry, in which they attributed the action of catalase on H2O2 to the initiation of a radical reaction (Reaction 6, the Fenton reaction) [86]. Later, Haber and his assistant Joseph Weiss [87, 88] investigated the decay of H2O2 by iron salts at low pH and concluded that, since more than 1 H2O2 per 2 Fe2+ is consumed when H2O2 is present in excess, the mechanism involves a radical chain reaction (Reactions 7 and 8), with chain termination by Reaction 9. Fe2+ + H2O2

Fe3+ + HO + HO•

(6)

HO• + H2O2

H2O + O2• + H+

(7)

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O2• + H+ + H2O2 Fe2+ + HO• + H+

O2 + HO• + H2O Fe3+ + H2O

(8) (9)

By the late 1940s, the mechanism proposed by Haber and Weiss for the iron-catalyzed decomposition of H2O2 had been criticised by Philip George, who showed that O2•– does not react with H2O2 [89]. An alternative mechanism at low pH proposed by George and co-workers in 1949 [90] on the basis of product analyses is summarised in Table 1. Note that O2 occurs in these reactions as HO2 (pKa = 4.8) [43]. More extensive proposals for the mechanism were described in two publications in 1951 [91, 92], the first of which contains a reference to Fenton's first full report of the oxidation of tartaric acid [93]. The rate constants were determined later. The mechanism published by George and coworkers was corroborated in 1985 [94]. It should be noted that all of these investigations of iron-catalyzed decomposition of H2O2 were carried out at low pH and are generally not physiologically significant. A new field of study, free radical biochemistry, was created in the wake of the discovery in 1969 of an enzymatic function for hemocuprein [95], namely catalysis of the disproportionation of O2 (Reaction 1) [96]. The rate constant for the copper/zinc SOD (CuZnSOD) reaction, which is governed by electrostatic guidance of O2 to the active site [97, 98], indicates that the reaction is close to diffusion-controlled (kcat 1 2 109 M 1s 1) [99, 100]. The majority of

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CuZnSOD, ca. 5–10 M, is found in the cytosol of the cell; some is also found in the mitochondrial intermembrane space [101]. In 1973, Fe- and Mn-containing SODs were discovered [102, 103]: the mitochondrial matrix contains manganese SOD (MnSOD), and bacteria contain a structurally related Fe-enzyme. A tetrameric CuZnSOD manages O2 in the extracellular space [104]. Given that Reaction 1 uncatalyzed is rapid (k = ca. 106 M 1s 1 at neutral pH), and that SODs are widespread, the conclusion is warranted that O2 is reactive and, therefore, must be dangerous. Unfortunately, George‟s work from the 1940s was not consulted, and, in the 1970s, reduction of H2O2 by O2 (Reaction 8) from the Haber-Wilstätter mechanism was proposed [105] as a possible harmful reaction from which the organism needed to be protected. At this point in history, Reaction 8, in which the relatively innocuous O2 is converted to the far more reactive HO radical, became known as the Haber-Weiss reaction#. Again, it had to be pointed out in other studies that Reaction 8 is too slow to be significant [106-111]. When it was realized that O2 does not reduce H2O2, iron complexes were invoked to act as catalysts, as in the mechanism of George et al., and Reactions 6 and 9 became known as the “Fenton-catalyzed Haber-Weiss reaction.” Later, it was realised and accepted that other reductants, e.g., monohydrogen ascorbate, are more likely candidates to reduce Fe3+ complexes in vivo.

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5. Biological Relevance of the Fenton Reaction The toxicity of iron, and, by implication, copper, originates from its ability to reduce peroxides, via Fenton chemistry. Although George and coworkers provided a mechanism for the decay of H2O2 catalyzed by iron at low pH, it is not clear that the mechanism has any meaning at neutral pH where aqueous Fe3+ ions are not available for reaction because they have formed hydroxide, phosphate, or other complexes. If such Fe3+ complexes are present in solution, are they reduced by O2 , or monohydrogen ascorbate, and are the corresponding Fe2+ complexes likely to be oxidized by H2O2 or organic peroxides? Table 1. Rate constants for the reactions in the mechanism of iron-catalyzed decomposition of H2O2 at low pH [89] Reactions Fe2+ + H2O2 + H+ Fe3+ + H2O + HO• HO• + H2O2 H2O + HO2• Fe2+ + HO• + H+ Fe3+ + H2O 2+ + Fe + HO2 + H Fe3+ + H2O2 Fe3+ + HO2 Fe2+ + O2 + H+

#

Reaction No. 9 7 10 11 12

k (M 1s 1) 41.5 2.7 • 107 4.3 • 108 1.2 • 106 2.0 • 104 (pH 1)

It is not clear why Reactions 7 and 8 have become known as the Haber-Weiss cycle rather than the HaberWillstätter cycle; it may be because the often-quoted paper by Haber and Weiss is in English and, thus, more accessible to the international research community, while the language of the original paper of Haber and Willstätter is German. Haber and Willstätter 87 is cited by Haber and Weiss 88 but neither paper refers to Fenton 84, although Reaction 6 is the Fenton reaction.

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The thermodynamic requirements for a metal complex in aqueous solution at neutral pH to redox-cycle are simple: when O2 – is the reductant, the metal-chelate must have a electrode potential between E°‟(O2/O2 –), –0.35 V and E°‟(H2O2/ OH,H2O), +0.39 V [5, 112]. The window of redox-opportunity (0.74 V) is therefore wide. For both iron and copper, redox cycling involves transfer of a single electron; thus, only one-electron electrode potentials may be considered. Potentials for two-electron oxidations, e.g., two-electron electrode potential of the NAD+, H+/NADH couple (–0.32 V) [113, 114], should never be considered in discussions of redox-cycling. Such a reaction is kinetically not feasible, as it would require the simultaneous collision of three reactants, NADH and two iron complexes, which is unlikely under the dilute concentrations found physiologically. Although O2 – has been shown to support hydroxylation of benzoic acid catalyzed by iron chelates [115], these studies were carried out with chelating agents that, with the exception of citrate [116], are not physiologically relevant. It is also important to recognize that, in most tissue compartments, the steady-state concentration of O2 – is maintained at a vanishingly low level by SODs [117]. The rate constant for reduction of iron(III)-edta† by O2 –, determined directly by pulse radiolysis, is ca. 1 • 106 M 1s 1 at physiological pH [118], three orders of magnitude smaller than the catalytic rate constant of CuZnSOD; rate constants for the reaction of iron(III)-dtpa† and -atp were much less than 1 • 106 M 1s 1 and could not be determined. Not surprisingly, it is now felt that monohydrogen ascorbate is a more likely physiological reductant of iron(III) complexes. Given that the electrode potential of the asc /Hasc couple is +0.28 V [119], the redox window for redox cycling involving ascorbate is much smaller, i.e., 0.11 V. However, for a typical serum concentration of 50 M monohydrogen ascorbate, the concentration of the ascorbyl radical is likely to be in the nanomolar range. Thus, the electrode potential for the ascorbyl/monohydrogen ascorbate couple is, according to the Nernst equation, ca. 0.18 V less than +0.28 V, which would widen the redox window to 0.29 V. Similar considerations of the H2O2,H+/HO , H2O couple increases the electrode potential from +0.39 V to ca. +0.9 V, an adjustment that results in a much larger window of 0.8 V, from +0.1 V to +0.9 V [120]. A number of studies with iron complexed to citrate, atp or aminopolycarbonates, such as edta, dtpa and nta†, suggest that the Fenton reaction, and its equivalent with organic peroxides, is thermodynamically feasible [121] and does proceed. Rate constants of the order of 102 to 105 M 1s 1 have been reported for the Fenton reactions of these various iron(II) complexes [122131]; there is, however, no good agreement between the published rate constants for a given complex. The reaction, which requires binding of H2O2 to iron(II), generally proceeds faster when more exchangeable water molecules are bound to iron(II) [132], i.e., when more ligand sites on the iron are available. However, all these reactions are much slower than that of H2O2 with catalase, and, thus, not likely to be physiologically significant.**

**

It is not sufficient to simply compare rate constants; the relative concentrations of the Fe2+ complex and catalase must be considered as well. It is the product of the concentration of the iron complexes or catalase with the respective rate constants for the reaction with H2O2 that must be compared. Catalase is present at ca. 2 M, and the concentration of redox-active iron is, at most, a few M. In the following, the concentration of the Fe2+ complex 5 M is used as an example to demonstrate that the Fenton reaction with Fe2+ complexes is outcompeted by catalase. This example is, of course, valid only for homogeneous solution: kFenton[Fe2+] = (1 • 104 M 1s 1)(5 • 10 6 M) = 0.05 s 1

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99

In vitro studies performed under physiological conditions indicate that HO• is not the only oxidant formed during the Fenton reaction [121, 133-135]. Depending on the conditions, the oxidant may be (1) oxidoiron(IV), a higher oxidation state of iron, as suggested during the 1930s (Reaction 13) [136], (2) H2O2 bound to iron(III), or (3) HO•. The pattern of hydroxylation of salicylate by iron(II)-edta and H2O2 compared to that by HO• generated by radiolysis of water suggests that HO• formation is likely [137]. Isotope/ESR studies established that the oxygen in either the HO• or oxidoiron(IV) is from H2O2 [138].

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Fe2+ + H2O2

FeO2+ + H2O

(13)

What is the likelihood that the Fenton reaction occurs in vivo? Assuming that monohydrogen ascorbate is the source of electrons in vivo, there are number of problems with the hypothesis that iron is a catalyst in oxidative processes: (1) We do not know the composition/structure of the iron complex (or complexes) in vivo reduced by monohydrogen ascorbate and subsequently oxidized during the Fenton reaction, nor do we know any rate constants for the reactions. (2) The rate constants for the Fenton reaction (102 105 M 1s 1) that have been determined are remarkably unimpressive when compared to that for the reaction of catalase with H2O2 (3.5 107 M 1s 1) [139]. Additionally, H2O2 is scavenged by glutathione peroxidase [140] and peroxiredoxin [141]. (3) The Fenton reaction is even slower in the presence the NO , which is present in vivo and which binds to iron(II) [142]. (4) The hypothesis does not include a protective role for SOD, but only for proteins that remove H2O2, which is contrary to experimental observations that SOD alone can protect tissues from oxyradical damage. Given that most biomolecules are unreactive toward O2 –, concentrations of ca. 5–10 M CuZnSOD in the cytosol would seem higher than necessary to protect against damage from O2 –. Clearly, SOD does protect, but not from Fenton chemistry. Beckman et al. [143] wrote in 1990 that “generation of strong oxidants by the iron-catalyzed Haber-Weiss reaction is not an entirely satisfactory explanation for SOD-inhibitable injury in vivo” and proposed that NO reacts with O2 – to form ONOO–, and that formation of ONOOH is kinetically far more feasible than the “iron-catalyzed Haber-Weiss reaction.” Earlier in this Chapter, we described ONOO as an important and selective oxidizing and nitrating species. Thus, SOD protects by eliminating O2 – and prevents, thereby, formation of ONOO . The diffusion-controlled rate constant for the reaction of O2 – with NO [57] necessitates the micromolar concentration of SOD observed in vivo. Where, then, may the Fenton reaction play a role? In the acidic environment of the lysosome, where ferritin is decomposed, the pH is between 4 and 5; here, the concentration of chelatable iron is relatively high (ca. 15 M), catalase is absent, the coordination of iron is altered and the iron may be reduced due to the presence of cysteine. Under conditions of oxidative stress, lysosomes may be damaged and ruptured, which would release relatively high concentrations of redox-active iron [144, 145] into the cytosol (Vol. II, Chapter 16). Ferritins released into the bloodstream during inflammation and autoimmune diseases may contribute to locally high iron concentrations that lead to apoptosis [146]. Fenton chemistry-derived oxidative damage can be prevented by chelation of the iron with desferrioxamine (dfo) [146-148].

kcatalase[catalase]

= (3.5 • 107 M 1s 1)(2 • 10 6 M) = 70 s

1

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The extracellular NTBI levels may contribute to intracellular labile iron pool (LIP) levels (Vol. II, Chapter 19) and play a role in oxyradical damage [149], or be a symptom of iron overload. Any form of mobile iron poses a threat to the liver and the heart, where excess iron accumulates, and iron chelation therapy is employed to promote excretion. The „wish list‟ for an effective iron chelator has been described as: „stable in vivo, non-toxic, rapidly excreted, preferably orally-active, and produced with a low synthetic cost‟ [150]. Toxicity is ascribed to Fenton chemistry, thus, the chelate ought not be redox-active: the three chelating agents used clinically, dfo, cp20 and icl670, bind iron(III) far more tightly than iron(II), which results in very negative standard electrode potentials ( 0.45 to 0.60 V [120, 151-153]) well outside the window of redox opportunity, such that monohydrogen ascorbate cannot reduce these iron(III) complexes. The low electrode potentials also imply that dfo, cp20 and icl670 bind iron(II) at micromolar concentrations only in part or not at all, which has been experimentally verified for cp20 [154]: when iron(II) is oxidized, it is complexed by cp20, thereby preventing reduction and redox cycling. Although copper is less ubiquitous than iron in living systems, copper and iron proteins have similar functions, including electron transfer and O2 binding and activation. In humans, copper is used for electron transfer reactions in the enzymes CuZnSOD and cytochrome c oxidase. During the 1980s, it was recognized and demonstrated that free copper ions can, in theory, redox cycle and promote reactions analogous to those catalyzed by free iron [155158]. In in vitro studies under acidic conditions, copper(I) was shown to form an intermediate Cu+H2O2, which reacts with organic scavengers, however, not by a Fenton mechanism [159]. Other investigators [160] concluded on the basis of pH and substrate concentration dependence experiments that no HO is formed during the reaction of copper(I) with H2O2. Studies with red blood cells show that monohydrogen ascorbate can reduce copper complexes to initiate reactions similar to those described for iron [161]. However, the concentration of free copper ions in the cell is extremely low, only ca. 1 copper ion per cell [162], and the possibility that copper participates in oxyradical damage reactions in vivo is vanishingly small. The disproportionation of O2 – catalyzed by free copper ions is faster by a factor of 4 than that catalyzed by SOD [163, 164]; the inclusion of edta to bind free copper(II) prevents the reaction with O2 –. It is important to note that edta, nta and dtpa, common metal chelation agents that are often used as models for physiological low-molecular-weight iron complexes in biochemical studies, may enhance the Fenton activity of iron at neutral pH. Depending on the complex, one or more coordination sites of iron are occupied by water, which may give way to H2O2 [132]. In the case of copper, the complexes formed with these ligands are essentially inert at physiological pH, and redox chemistry is prevented. Thus, addition of chelators can produce differential effects on redox behavior depending on the target metal, e.g., edta and nta enhance iron-mediated damage to chromatin but protect against copper-mediated damage [165]. Similar differential effects of edta on oxidative reactivity in cerebrospinal fluid and brain tissue was recently reported [166], and these authors correctly point out that different chelation strategies are required for sequestration of iron – “the main characteristics of an iron sequestrant should be its ability to compete with metal ligands naturally present in CSF” – and copper – “copper requires application of a strong and efficient chelator”.

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6. Cautionary Remarks Given the low micromolar concentrations of iron and H2O2 found under normal physiological conditions and the rate constants for Fenton reactions with iron bound to physiologically relevant ligands, it is kinetically unlikely that iron causes damage, and the same may hold for copper. Iron and copper may contribute to oxidative processes under conditions of higher local concentrations of these ions. The destruction of ferritin (and by extension ceruloplasmin) in lysosomes may provide such concentrations. NTBI in blood is an indicator of excess iron; this form of iron does not appear to be very redox-active, but can be considered dangerous since it is a form of mobile iron that may be deposited in the liver or in the heart. To determine the rate constant for a reaction of physiological interest, it is usually necessary to use reagents at concentrations that are significantly higher than physiological conditions; the rate constant thus determined can be extrapolated to the dilute conditions found in vivo to estimate how fast the process is in reality. However, a rate constant determined in vitro is valid for homogeneous solution, whereas the many reactions going on inside the cell take place under more heterogeneous conditions. Thus, the observation that common HO∙ scavengers fail to protect against oxidative damage in vivo has been attributed to formation of HO∙ at locations within cellular components that are poorly accessible to the scavenger. Alternatively, formation of oxidoiron(IV), which might react less rapidly than HO∙ with scavengers, has been postulated. Results obtained from experiments performed with cells to which relatively high concentrations of metal ions and/or H2O2 have been added cannot be realistically extended to prove that oxidative damage occurs under normal physiological conditions, where the antioxidant defence system consisting of monohydrogen ascorbate, glutathione, vitamin E, SODs, catalase, glutathione peroxidase, peroxiredoxin, etc. is intact. Some in vitro experiments can be additionally criticized for having used simple iron salt solutions: e.g., iron(III) salt solutions are very acidic, and, at neutral pH, colloids of iron(III) hydroxide/phosphate and iron(III)oxide, which may simply be toxic rather than catalysts of redox reactions, are formed. Iron(II) autoxidizes rapidly at pH 7 to generate O2∙– and H2O2, which are likely to complicate the interpretation of findings. In short, the speciation of a metal ion determines its thermodynamic and kinetic properties and, by extension, its reactivity. One obstacle to progress in the field of redox metal toxicity is the inability to follow oxidative stress in cells in a time-resolved fashion. Although widely used fluorescence techniques have been touted to do so, the addition of the fluorescent indicator itself may lead to the release of PROS [167, 168].

Conclusions Are redox-active metal ions causative agents in disease, or do diseases cause redox-active metal ions to accumulate? Although metals, especially iron and copper, are clearly implicated in a variety of oxidative-stress-related pathologies, it has not yet been definitively demonstrated whether low-molecular-weight metal ion complexes act to initiate damage or

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are simply liberated as a sequela of oxyradical damage, and it is possible that metals, once released as a result of an oxidative insult, act in a cascade fashion to propagate injury. It is the findings from in vivo studies that most clearly support a role for involvement of redox-active metals in oxidative stress-related pathologies; results from in vitro mechanistic studies based on kinetics have sometimes called those findings into question. Both approaches to the study of free radical biology and medicine have merit and contribute to the ultimate goal of elucidating mechanisms that make sense at the molecular, cellular and organismic levels alike.

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[112] Koppenol WH. Thermodynamics of Fenton-driven Haber-Weiss and related reactions. In: Cohen G, Greenwald RA, eds. Oxy Radicals and Their Scavenging Systems. Vol. I. Molecular Aspects. New York: Elsevier Biomedical, 1983: 84-88. [113] Pierre JL, Fontecave M. Iron and activated oxygen species in biology: The basic chemistry. BioMetals 1999; 12:195-199. [114] Pierre JL, Fontecave M, Crichton RR. Chemistry for an essential biological process: the reduction of ferric ion. BioMetals 2002; 15:341-346. [115] Baker MS, Gebicki JM. The effect of pH on the conversion of superoxide to hydroxyl free radicals. Arch Biochem Biophys 1984; 234:258-264. [116] Baker MS, Gebicki JM. The effect of pH on yields of hydroxyl radicals produced from superoxide by potential biological iron chelators. Arch Biochem Biophys 1986; 246:581-588. [117] Koppenol WH, Bounds PL. Hormesis. Science 1989; 246:311. [118] Butler J, Halliwell B. Reaction of iron-EDTA chelates with the superoxide radical. Arch Biochem Biophys 1982; 218:174-178. [119] Williams MH, Yandell JK. Outer-sphere electron transfer reactions of ascorbate anions. Aust J Chem 1982; 35:1133-1144. [120] Merkofer M, Kissner R, Hider RC, Brunk UT, Koppenol WH. Fenton chemistry and iron chelation under physiological relevant conditions: Electrochemistry and kinetics. Chem Res Toxicol 2006; 19:1263-1269. [121] Koppenol WH. Chemistry of iron and copper in radical reactions. In: Rice-Evans CA, Burdon RH, eds. Free Radical Damage and its Control. Amsterdam: Elsevier Science B.V., 1994: 3-24. [122] Sutton HC, Winterbourn CC. Chelated iron-catalyzed OH. formation from paraquat radicals and H2O2:mechanism of formate oxidation. Arch Biochem Biophys 1984; 235:106-115. [123] Rahhal S, Richter HW. Reduction of hydrogen peroxide by the ferrous iron chelate of diethylenetriamine-N,N,N',N",N"-pentaacetate. J Am Chem Soc 1988; 110:31263133. [124] Wink DA, Nims RW, Desrosiers MF, Ford PF, Keefer LK. A kinetic investigation of intermediates formed during the Fenton reagent mediated degradation of Nnitrosodimethylamine: Evidence for an oxidative pathway not involving hydroxyl radical. Chem Res Toxicol 1991; 4:510-512. [125] Yamazaki I, Piette LH. ESR spin-trapping studies on the reaction of Fe2+ ions with H2O2-reactive species in oxygen toxicity in biology. J Biol Chem 1990; 265:1358913594. [126] Rush JD, Koppenol WH. The reaction between ferrous aminopolycarboxylate complexes and hydrogen peroxide. An investigation of the reaction intermediates by stopped-flow spectrophotometry. J Inorg Biochem 1987; 29:199-215. [127] Rush JD, Koppenol WH. Reactions of FeIInta and FeIIIedda with hydrogen peroxide. J Am Chem Soc 1988; 110:4957-4963. [128] Rush JD, Maskos Z, Koppenol WH. Reactions of iron(II)nucleotide complexes with hydrogen peroxide. FEBS Lett 1990; 261:121-123. [129] Gilbert BC, Jeff M. Oxidative damage and radical repair: One-electron transfer reactions involving metal ions, peroxides and free radicals. In: Rice-Evans C,

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[130]

[131] [132] [133]

[134]

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[138]

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[139] [140]

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Dormandy T, eds. Free Radicals: Chemistry, Pathology and Medicine. London: The Richelieu Press, 1988: 25-49. Croft S, Gilbert BC, Lindsay Smith JR, Whitwood AC. An E.S.R. investigation of the reactive intermediate generated in the reaction between Fe(II) and hydrogen peroxide in aqueous solution. Direct evidence for the formation of the hydroxyl radical. Free Radic Res Commun 1992; 17:21-39. Rush JD, Koppenol WH. Reactions of Fe(II)-ATP and Fe(II)-citrate complexes with tert-butyl hydroperoxide and cumylhydroperoxide. FEBS Lett 1990; 275:114-116. Graf E, Mahoney JR, Bryant RG, Eaton JW. Iron-catalyzed hydroxyl radical formation. J Biol Chem 1984; 259:3620-3624. Rush JD, Maskos Z, Koppenol WH. Distinction between hydroxyl radical and ferryl species. In: Packer L, Glazer AN, eds. Methods in Enzymology, Vol. 186. San Diego: Academic Press, 1990: 148-156. Yamazaki I, Piette LH. EPR spin-trapping study on the oxidizing species formed in the reaction of the ferrous ion with hydrogen peroxide. J Am Chem Soc 1991; 113:7588-7593. Wardman P, Candeias LP. Fenton chemistry: An introduction. Radiat Res 1996; 145:523-531. Bray WC, Gorin MH. Ferryl ion, a compound of tetravalent iron. J Am Chem Soc 1932; 54:2124-2125. Maskos Z, Rush JD, Koppenol WH. The hydroxylation of the salicylate anion by a Fenton reaction and gamma-radiolysis: A consideration of the respective mechanisms. Free Radic Biol Med 1990; 8:153-162. Lloyd RV, Hanna PM, Mason RP. The origin of the hydroxyl radical oxygen in the Fenton reaction. Free Radic Biol Med 1997; 22:885-888. Chance B, Greenstein DS, Roughton FJW. The mechanism of catalase action. I. Steady state analysis. Arch Biochem Biophys 1952; 37:301-321. Esworthy RS, Chu F-F, Geiger P, Girotti AW, Doroshow JH. Reactivity of plasma glutathione peroxidase with hydroperoxide substrates and glutathione. Arch Biochem Biophys 1993; 307:29-34. Peskin AV, Low FM, Paton LN, Maghzal GJ, Hampton MB, Winterbourn CC. The high reactivity of peroxiredoxin 2 with H2O2 is not reflected in its reaction with other oxidants and thiol reagents. J Biol Chem 2007; 282:11885-11892. Lu C, Koppenol WH. Inhibition of the Fenton reaction by nitrogen monoxide. J Biol Inorg Chem 2005; 10:732-738. Beckman JS, Beckman TW, Chen J, Marshall PA, Freeman BA. Apparent hydroxyl radical production by peroxynitrite: Implications for endothelial injury from nitric oxide and superoxide. Proc Natl Acad Sci USA 1990; 87:1620-1624. Yu ZQ, Persson HL, Eaton JW, Brunk UT. Intra-lysosomal iron: A major determinant of oxidant-induced cell death. Free Radic Biol Med 2003; 34:12431252. Persson HL, Yu ZQ, Tirosh O, Eaton JW, Brunk UT. Prevention of oxidant-induced lysosomal cell death by lysosomotropic iron-chelators. Free Radic Biol Med 2003; 34:1295-1305.

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[146] Bresgen N, Jaksch H, Lacher H, Ohlenschläger I, Uchida K, Eckl PM. Iron-mediated oxidative stress plays an essential role in ferritin-induced cell death. Free Radic Biol Med 2010; 48(10):1347-1357. [147] Kurz T, Leake A, Von Zglinicki T, Brunk UT. Relocalized redox-active lysosomal iron is an important mediator of oxidative-stress-induced DNA damage. Biochem J 2004; 378(3):1039-1045. [148] Kurz T, Gustafsson B, Brunk UT. Intralysosomal iron chelation protects against oxidative stress-induced cellular damage. FEBS J 2006; 273:3106-3117. [149] Pietrangelo A. Iron, firend or foe? "Freedom" makes the difference. J Hepatol 2000; 32:862-864. [150] O'Sullivan B, Xu J, Raymond KN. New multidentate chelators for iron. In: Badman DG, Bergeron RJ, Brittenham GM, eds. Iron Chelators. New Development Strategies. Ponte Vedra Beach, FL: The Saratoga Group, 2000: 177-208. [151] Cooper SR, McArdle JV, Raymond KN. Siderophore electrochemistry: Relation to intracellular iron release mechanism. Proc Natl Acad Sci USA 1978; 75:3551-3554. [152] Steinhauser S, Heinz U, Bartholomä M, Weyhermüller T, Nick H, Hegetschweiler K. Complex formation of ICL670 and related ligands with Fe(III) and Fe(II). Eur J Inorg Chem 2004;4177-4192. [153] Merkofer M, Kissner R, Koppenol WH. Redox properties of the iron complexes of CP20, CP502, CP509 and ICL670. Helv Chim Acta 2004; 87:3021-3034. [154] Merkofer M, Domazou AS, Nauser T, Koppenol WH. Dissociation of CP20 from iron(II)(cp20)3; A pulse radiolysis study. Eur J Inorg Chem 2006;671-675. [155] Marshall LE, Graham DR, Riech KA, Sigman DS. Cleavage of deoxyribonucleic acid by the 1,10-phenanthroline-cuprous complex. Hydrogen peroxide requirement and primary and secondary structure specificity. Biochemistry 1981; 20:244-250. [156] Gutteridge JMC. Tissue damage by oxy-radicals: The possible involvement of iron and copper complexes. Med Biol 1984; 62:101-104. [157] Aust SD, Morehouse LA, Thomas CE. Role of metals in oxygen radical reactions. J Free Radic Biol Med 1985; 1:3-25. [158] Stoewe R, Prütz WA. Copper-catalyzed DNA damage by ascorbate and hydrogen peroxide: kinetics and yield. J Free Radic Biol Med 1987; 3:97-105. [159] Masarwa M, Cohen H, Meyerstein D, Hickman DL, Bakac A, Espenson JH. Reactions of low-valent transition-metal complexes with hydrogen peroxide. Are they "Fenton-like" or not? 1. The case of Cu+aq and Cr2+aq. J Am Chem Soc 1988; 110:4293-4297. [160] Johnson GRA, Nazhat NB, Saadalla-Nazhat RA. Reaction of aquacopper(I) ion with hydrogen peroxide. J Chem Soc Faraday Trans I 1988; 84:501-510. [161] Shinar E, Rachmilewitz EA, Shifter A, Rahamim E, Saltman P. Oxidative damage to human red cells induced by copper and iron complexes in the presence of ascorbate. Biochim Biophys Acta 1989; 1014:66-72. [162] Rae TD, Schmidt PJ, Pufahl RA, Culotta VC, O'Halloran TV. Undetectable intracellular free copper: The requirement of a copper chaperone for superoxide dismutase. Science 1999; 284:805-808. [163] Klug-Roth D, Rabani J. Pulse radiolytic studies on reactions of aqueous superoxide radicals with copper(II) complexes. J Phys Chem 1976; 80:588-591.

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[164] Butler J, Koppenol WH, Margoliash E. Kinetics and mechanisms of the reduction of ferricytochrome c by the superoxide anion. J Biol Chem 1982; 257:10747-10750. [165] Dizdaroglu M, Rao G, Halliwell B, Gajewski E. Damage to the DNA bases in mammalian chromatin by hydrogen peroxide in the presence of ferric and cupric ions. Arch Biochem Biophys 1991; 285:317-324. [166] Spasojevic I, Mojovic M, Stevic Z, Spasic SD, Jones DR, Morina A et al. Bioavaliability and catalytic properties of copper and iron for Fenton chemistry in human cerebrospinal fluid. Redox Rep 2010; 15:29-35. [167] Zielonka J, Kalyanaraman B. "ROS-generating mitochondrial DNA mutations can regulate tumor cell metastasis"-a critical commentary. Free Radic Biol Med 2008; 45(9):1217-1219. [168] Muller FL. A critical evaluation of cpYFP as a probe for superoxide. Free Radical Biol Med 2009; 47:1779-1780.

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 6

Protein Oxidation Tilman Grune* Institute of Biological Chemistry and Nutrition, University of Hohenheim, Stuttgart, Germany

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1. Introduction: Why does Protein Damage Matter? Reactive oxygen species (ROS) are generated constantly within cells at low concentrations under physiological conditions (Chapter 2). Cellular production of ROS occurs from both enzymatic and non-enzymatic sources. Any electron-transferring protein or enzymatic system can result in the formation of ROS as “by-products” of electron transfer reactions, usually to O2. An enhanced ROS production might occur as the outcome of acute cell stresses. Cellular oxidative damage is the result of an imbalance between ROS-generating and ROS-scavenging systems [1]. Proteins are among the main targets for oxidants as a result of their abundance in biological systems, and the high rate constants for several reactions of proteins with ROS. Most protein damage is non-repairable, and has deleterious consequences on protein structure and function [2]. Oxidative protein damage in vivo is important because it can impair the functioning of receptors, antibodies, signal transduction, transport proteins and enzymes. The dysfunctional proteins can also lead to secondary damage to other biomolecules. Thus, oxidative damage that functionally inactivates DNA repair enzymes may result in oxidative DNA damage (Chapter 9) and increased mutation frequency, while oxidation of DNA polymerases may decrease their fidelity in replicating DNA. Furthermore, oxidized proteins may be recognized as „foreign‟ by the immune system [3], triggering antibody formation and perhaps autoimmunity. Most oxidized proteins are *

Correspondence to: Tilman Grune. Institute of Nutrition, Department of Nutritional Toxicology, Friedrich Schiller University, Dornburger Str. 24, 07742 Jena, Germany. Phone: +49 3641 949670; Fax: +49 3641 9496 72; Email: [email protected]

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catabolized by proteasomal and lysosomal pathways, but some proteins appear to be poorly degraded and, therefore, accumulate within cells. This accumulation may contribute to a range of human pathologies.

2. How does Damage Occur? Damage to proteins can occur directly by attack of reactive species (ROS or RNS), or by secondary damage involving the attack of products of lipid peroxidation, such as 4hydroxynonenal (4-HNE), malondialdehyde (MDA) or acrolein. Oxidation can occur at both the protein backbone and at amino acid side-chains. The side of attack depends on numerous factors. Furthermore, proteins can also be damaged by glycoxidation, a non-enzymatic chemical reaction between reducing carbohydrates and proteins involving oxygen. Some oxidants have a very specific damage pattern and react with a limited number of amino acid residues, whereas other ROS, such as the hydroxyl radical (HO•), lead to a widespread and relatively non-specific damage. Some protein damage is reversible, such as peroxiredoxin inactivation, disulfide formation, methionine sulfoxide formation, destruction of Fe-S clusters by O2 - and, possibly, nitration. Other damage, like oxidation of side-chains to carbonyl residues, appears irreversible and the protein has to be degraded and replaced. This Chapter concentrates on the reactions of oxidants with proteins as the major component of most biological systems. It provides an outline of key oxidation pathways and amino acid modifications.

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3. Chemistry of Protein Oxidation The chemistry of protein oxidation is thought to be more complex than that of oxidative DNA damage, because instead of four bases and one sugar there are 20 amino acids, each able to form several oxidation products. The amino acid side-chains provide ample sites for reaction with oxidants and yield a large array of products. There is no generally accepted scheme for classification of the oxidative modifications. For ease of discussion, these have been classified into categories which focus on the most important modifications.

3.1. Backbone Damage The reaction rate of most non-radical oxidants with the protein backbone is slow and, therefore, little damage is observed. On the other hand, radicals react rapidly with the protein backbone. This occurs primarily via hydrogen atom abstraction at the -carbon site (reaction (1) [4]), to give a carbon-centered radical.

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These species seem to have two major fates: I.

Reaction with another radical, which is in most situations a limited process due to low radical fluxes and steric interactions. II. Reaction with O2 to give a peroxyl radical (ROO•; reaction (2) [4])

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The fate of peroxyl radicals formed on proteins is incompletely understood. One main decomposition pathway appears to be via the release of a hydroperoxyl radical (HO2• ). This in turn generates an imine, which afterwards undergoes hydrolysis, eventually resulting in backbone fragmentation (reaction (3) [5]).

An alternative pathway involves hydrogen abstraction from another molecule and the formation of a hydroperoxide (reaction (4) [6]). Subsequent decomposition can also result in backbone fragmentation. Both mechanisms require oxygen. Under anaerobic conditions, little backbone fragmentation is detected, underscoring the key role of peroxyl radical formation in the fragmentation process [5].

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3.2. Aliphatic Residues

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The majority of radical-mediated reactions with aliphatic residues are again based on hydrogen atom abstraction which leads to carbon-centered radicals. With reactive species such as •OH, reaction can occur at all available sites of a protein, which results in the formation of a range of radicals [4]. There are three major fates for the generated carboncentered radical (Figure 1): I.

Dimerization with another radical. In most situations, this is a limited process due to low radical fluxes and steric hindrances. In the presence of O2 dimerization is uncommon, but in an anoxic environment, this reaction might play a major role in cross-linking within and between proteins. II. Carbon-centered radicals can be repaired by thiols with resulting formation of a thiyl radical (see also Chapter 4). This reaction seems to have a slow reaction rate [7]. III. The major pathway is a reaction with O2 to generate peroxyl radicals (Chapter 2). The fate of peroxyl radicals is incompletely understood. It is known that such species can undergo an array of radical-radical termination reactions that can generate alcohols and carbonyl-compounds. Also, peroxyl radicals can abstract hydrogen atoms and form hydroperoxides. Such hydroperoxides might be long-lived and persist in protein molecules [8]. Residues of leucine, valine, proline, and isoleucin are most susceptible to form stable hydroperoxides, having a tertiary carbon and two adjacent methylene groups. Other hydroperoxide-forming residues (partly non-aliphatic) are those of alanine, tyrosine, and arginine, while asparagine, aspartic acid, cysteine, methionine, glycine, histidine, phenylalanine, serine, and threonine do not form stable products or hydroperoxides at all [9]. A number of studies have quantified the yield of hydroperoxide groups formed on oxidized

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amino acids and proteins and it is evident that these modifications are main products formed via the oxidation of many amino acids and a wide range of oxidants [8]. Another notable reaction is the two-electron transfer from hypohalous acids (HOBr, HOCl) to give the corresponding unstable bromamines/bromamides and chloramines/ chloramides [10]. Their decomposition can give rise to carbonyl groups, especially with lysine and arginine [11].

3.3. Carbonylated Proteins

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Protein carbonylation is a non-enzymatic modification which can be produced by a multitude of oxidative pathways. These modifications appear irreversible and, therefore, the protein can be destroyed and become dysfunctional. ROS can react directly with lysine, arginine, proline, and threonine side chains, particularly via metal-catalyzed oxidation [12] engendering 2-aminoadipate semialdehyde from lysyl residue [13] (reaction (5)), glutamate semialdehyde from arginyl and prolyl residue [13] (reaction (6)), 2-pyrrolidone from prolyl residue [14] (reaction (7)), and 2-amino-3-ketobutyric acid from the threonyl residue [14] (reaction (8)) .

Figure 1. Major fates of carbon-centered radicals.

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Direct oxidation of proteins by ROS can yield carbonyl derivatives resulting from the cleavage of protein backbone by the -amidation pathway or by oxidation of side chains. In addition, carbonyl groups may be introduced into proteins by reactions with aldehydes (4HNE, MDA) produced during lipid peroxidation [15] (Chapter 7). Finally, carbonyl groups can be generated with reactive carbonyl derivatives (ketoamines, ketoaldehydes, deoxyosones) generated as a consequence of the reaction of reducing sugars or their oxidation products with lysine residues of proteins (glycation and glycoxidation products) [14].

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3.4. Oxidation of Sulfur Centers The susceptibility to sulfur-oxidation renders cysteine and methionine residues major sites of oxidation within proteins. Oxidation of cysteine residues is facile with a very wide range of oxidants. With most oxidants (one-radical process) thiyl radicals (RS•) are formed. The fate of these species is multifaceted with dimerization to give disulfide (RSSR, cystine) and generation of peroxyl radicals as a result of reaction with O2 [16]. Reaction of an initial thiyl radical with another radical can also occur with numerous other molecules, so mixed dimers are prevalent products (Chapter 4). As this reaction involves the interaction of two radicals, the rate of this process is situation-dependent and affected by steric factors when a thiyl radical is formed on a protein [4].

Figure 2. Modifications of cysteine.

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Reactions of cysteine residues with two-electron oxidants result in the initial formation of adduct species (e.g. RSNO species with NO+, RSCl species with HOCl or Cl2). Most of these species are short-lived and undergo hydrolysis reactions to give oxyacids (RSO2H-sulfinic acid, RSO3H-sulfonic acid) by a complex and incompletely understood chemistry, at least for proteins in a cellular system [16]. Furthermore, reactions with other thiol groups may yield dimer species (mixed dimers or cystine). Such disulfides are one of few oxidative lesions that can be repaired by the disulfide isomerase systems to maintain the cysteine residues in their reduced form [17]. There is considerable evidence that the regulated oxidation of cysteine and reduction of cystine represent a redox switching mechanism that controls the behavior of many critical proteins [18]. The repair of sulfinic and sulfonic acids in general is not possible, although the common consensus that sulfinic acid cannot be reduced to sulfenic acid or thiol has been challenged. In April 2003, several papers presented compelling evidence that formation of cysteine sulfinic acid in proteins might be a reversible process in vivo. This was shown for peroxiredoxins, but the component reducing the sulfinic acid remains unknown [19, 20]. Methionine residues are also readily oxidized by a large number of species due to their low oxidation potential. The main oxidation product is the corresponding sulfoxide, which may, to a lesser extent, be further oxidized to sulfone (reaction (9)) [21].

S H2C

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R4

R6

CH2

CH

C

NH

O

R5

O

oxidation

H2C R4

S

R6

CH2

O

further oxidation

CH

C

NH

O

R5

H2C R4

S

R6 O

CH2

(9)

CH

C

NH

O

R5

Sulfoxides are formed with a broad spectrum of one- and two-electron oxidants, but the formation mechanisms are obviously different. It is important to note that two stereoisomers are formed (S- (D-) and R- (L-) form) [22]. Interestingly, both stereoisomers of methionine sulfoxide can be repaired. A set of enzymes (methionine sulfoxide reductases) evolved, which act in different compartments and which are stereospecific. It has been suggested that methionine residues act as „drain‟ for reactive species, as the resulting sulfoxide can be readily repaired. Also, the reversible oxidation of methionine plays a role in redox signaling events [23].

3.5. Oxidation of Aromatic Residues The main reaction of most oxidants with aromatic residues is addition. Also electron transfer reactions to give transient radical-cations occur with strong oxidants like SO4-•. These cations usually react rapidly with water and give rise to hydroxylated products, resulting partly in protein fragmentation [24]. Furthermore, substitution reactions are known, like reaction of HOCl with histidine, which leads to short-lived N-chlorinated forms [25]. Reaction of various oxidants with tyrosine gives rise to tyrosine phenoxyl radical. One of the possible fates of this radical is dimerization, with the formation of dityrosine [17]. This can

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lead to intra- or inter-molecular cross-links in proteins [26]. Reaction with chlorinating species (Cl2 or HOCl) gives rise to chlorinated aromatics such as 3-chlorotyrosine, and reaction with nitrating species gives rise to nitrated products, like 3-nitrotyrosine (for further information see section 4.1) [17]. Most of these reactions are unaffected by the presence of O2, in contrast to modifications of protein backbone and aliphatic residues; therefore, a similar spectrum of products can be observed under oxygenated or anoxic conditions.

4. Oxidative Modifications of Amino Acid Residues due to RNS The presence of •NO in biological systems leads to the formation of reactive nitrogen species (RNS; see also Chapter 3). One of the potentially most significant reactions of •NO is with superoxide: •

NO + O2 - → ONOO-

The product is peroxynitrite a strong oxidant and the corresponding base to peroxynitrous acid, ONOOH. The reaction rate for the formation of peroxynitrite has been determined to be three times faster than scavenging of O2 - by superoxide dismutase (SOD) [27]. Nitric oxide is, therefore, the only known biological molecule produced under pathological conditions in sufficient concentrations to outcompete SOD for O2 -.

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Table 1. Main products formed on oxidation of aromatic amino acids and protein side-chains

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One important reaction of peroxynitrite is catalyzed by transition metals, including those within the reactive core of SOD. In this mechanism, peroxynitrite would first form a complex with the transition metal to yield a polarized carrier of nitronium cation (NO2δ+-Oδ--Me(n-1)+). Thereafter, mostly the polarized carrier of NO2+ may attack tyrosine as a two-electron acceptor to yield a nitroarenium ion intermediate, which then evolves to 3-nitrotyrosine (NO2-Tyrosine) and a proton [28]. The polarized peroxynitrite-transition metal-complex may decompose to free NO2+ by heterolysis, but the half-life of NO2+ is too short in aqueous systems and, therefore, the metal-bound NO2+ complex would be the main reactant [28].

4.1. 3-Nitrotyrosine

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Tyrosine nitration is a suitable marker for the production of reactive nitrogen-centered oxidants. It is not necessarily due to the formation of peroxynitrite, but the latter remains the most likely source in vivo. Other reactive species such as nitrogen dioxide can form nitrotyrosine in simple solutions. Acidified nitrite can also produce nitrotyrosine if left for several days in contact with a protein. However, the amounts of nitrogen dioxide or nitrite present in vivo are far lower than necessary to yield high levels of nitration in vivo. Addition of a NO2-group to tyrosine lowers the pKa of its phenolic -OH and adds a bulky substituent. Therefore, if relevant tyrosines are targeted, nitration may alter protein conformation and function and also inhibit tyrosine phosphorylation [28]. A notable loss of enzyme activity linked to nitration in vivo occurs with Mn-SOD [29]. Another scenario is a gain-of-function modification, in which case a small fraction of nitrated protein can elicit a substantive biological signal. This has been shown in a few proteins such as cytochrome c, which acquires a strong peroxidase activity after nitration [30].

5. Reactions of Relevant Oxidants 5.1. Hydroxyl Radical The hydroxyl radical reacts with all amino acid residues in proteins. However, amino acid residues having reducing properties, like thiol, thioether, methionyl, aromatic and heterocyclic moieties, are more susceptible to •OH attack [31]. The reaction starts with cleavage of chemical bonds and the production of the amino acid radical. Such radicals may interact with neighboring amino acid residues in proteins, effecting a free-electron migration within the molecule. As a result, the final reaction product does not have to correspond to the site of primary attack [32]. The transient radicals generated by •OH in amino acid backbones tend to create new chemical bonds, producing aggregation of protein molecules, especially by new -SS-bond and dityrosine bridge formation. Generally, the •OH interaction with proteins in the presence of oxygen and O2 - produces polypeptide fragmentation rather than aggregation, while in the absence of oxygen, aggregation predominates [33].

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5.2. Superoxide Anion Radical and Hydrogen Peroxide O2 - and H2O2 are relatively inert oxygen species. The superoxide anion radical reacts with proteins at physiological conditions much more slowly than the hydroxyl radical and is not considered critical for protein damage in vivo. However, there are some exceptions: superoxide radicals react rapidly with iron-sulfur clusters and inactivate proteins containing these clusters. The reaction rate constant for the inactivation of mitochondrial and cytosolic aconitase is very high, and other Fe-S proteins are also inactivated by O2 - [34, 35]. Reactions of H2O2 with proteins are slower compared to O2 -. Most reactions of H2O2 are mediated by the hydroxyl radical. For example, SOD treated with H2O2 produces 2-oxohistidine from histidine-118, which is located in the active site of the enzyme [36].

5.3. Hypochlorite

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Hypochlorite is another strong oxidant and its lifetime is long enough to react selectively with preferable electron donors. Such donors are thiols and thioethers, which compete for hypochlorite and act as HOCl scavengers, protecting less active reductants from oxidation. This protective function of thiol groups within the same polypeptide chain has been amply documented; for example, treatment of albumin with HOCl first induces oxidation of all available –SH groups [37]. The other amino acid residue that is particularly susceptible to oxidation is the indole moiety of tryptophan. HOCl oxidizes indole residues at a pH range of 2-11. Acting together with chloramines the reaction product is the 2-oxoindole derivative. This oxoindole formation is visible by spectral changes at 250 and 280 nm. At pH values between 3 and 5, oxoindole residues tend to spontaneously hydrolyze peptide bonds and undergo cyclization to iminolactone, effecting breakage of polypeptide structures [38].

Figure 3. Different stages of protein oxidation caused by increasing oxidative damage (according to Jung et al. [45]).

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5.4. Chloramines Chloramines exhibit a particular affinity for protein surface-oriented thiol and thioether groups. Almost exclusively, the surface-exposed methionine residues are attacked by chloramines, yielding methionine sulfoxide. Buried methionyl and cysteinyl residues remain unaffected or appear to be oxidized only partially [39].

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6. Protein Aggregation due to Oxidation The degree of protein oxidation caused by oxidants depends on many factors, including the nature and flux rate of the oxidant, location and the presence of antioxidants. Most of the oxidative modifications coincide with a malfunction of the protein. This might be a result of a direct modification in one of the protein domains required for function, or due to a partial unfolding of the protein (all the above mentioned modifications might lead to partially unfolded proteins). If a native protein in its active form is exposed to increasing amounts of oxidants, more and more amino acid side chains are oxidized and as a result the protein loses activity. The amino acid side chain oxidation is accompanied by an increased unfolding of the protein and due to that process hydrophobic amino acids, normally hidden in the protein core, are exposed to the surface. These unfolded protein structures constitute a degradation signal and can be recognized by the proteasomal system [40, 41] (see „Degradation of oxidatively damaged proteins‟). However, if these proteins are not degraded at this stage, the exposed hydrophobic structures tend to form thermodynamically driven aggregates (by hydrophobic and electrostatic interactions), in order to minimize the contact surface between these hydrophobic residues and the hydrophilic solvent [42]. With time, such aggregates cross-link covalently due to reactions between carbon-, oxygen-, and nitrogen-centered radicals of amino acid side chains and due to the action of cross-linking molecules like bifunctional aldehydes (4-HNE and MDA), products of lipid peroxidation [43]. In general, such material is referred to as lipofuscin, lipofuscin-like pigments, ceroid or age pigment [44, 45] (see also Vol. II, Chapter 16). Such protein aggregates are essentially insoluble and metabolically stable under normal physiological conditions. Aggregates are unrelated to the original function of the protein, but may introduce a new toxic element into cellular metabolism. The occurrence of protein aggregates in cells may trigger a multitude of intracellular reactions. Most protein aggregates are ubiquitinated, a process that may culminate in cell cycle arrest. Heavily-oxidized and cross-linked proteins are also poor substrates for the proteasome and may, in fact, inhibit proteasomal activity [46].

7. Repair of Protein Damage After oxidation of proteins, living cells try to rescue the defective polypeptides and restore their function. This process is energetically more advantageous than re-synthesis. However, repair mechanisms evolved only for the most frequent and easy reparable protein modifications and most oxidized proteins undergo selective proteolysis. Disulfide bonds and

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Protein Oxidation methionine sulfoxides can be repaired enzymatically by [47], thioredoxin(Trx)/thioredoxin reductase system reductases (Msr) [49]. Furthermore, there are a number stress proteins, with the ability to reconstitute native unfolding.

protein disulfide isomerase (PDI) [48], and methionine sulfoxide of proteins, such as heat shock or protein structure after oxidative

7.1. Methione Sulfoxide Reductase Methionine sulfoxide reductases (Msr) play a key role in the repair of methionine residues. Methionine sulfoxide has R- and S-stereoisomers, with MsrA (also called MsrS) specific for S-isomers and MsrB (MsrR) for R-isomers. In animal cells it appears that there is one gene that codes for MsrA and three genes coding for MsrB proteins. MsrA localizes primarily in the mitochondria or cytoplasm, whereas the three MsrB proteins have different subcellular localizations. MsrB1, a selenoprotein, is mainly deployed in the nucleus and cytoplasm, MsrB2 in the mitochondria and MsrB3 within mitochondria and the endoplasmic reticulum. The reducing power for MsrA and MsrB is supplied by thioredoxin which may replaced by dithiothreitol in laboratory assays often [23, 49].

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7.2. Thioredoxin/thioredoxin Reductase System Thioredoxins (Trx), polypeptides with a relative molecular mass of about 12 kDa, have been isolated and characterized in prokaryotes, yeast, plants and mammalian cells (see Vol. II, Chapter 9). Several different thioredoxins have been described, for example the cytosolic Trx1 and the mitochondrial Trx2. Thioredoxins are classically defined by their ability to reduce disulfides (S-S bond) to thiols (two –SH groups) and in this process the thioredoxin is oxidized to a disulfide. The thioredoxin disulfide is then cycled back to its active form by the enzyme thioredoxin reductase, using NADPH as electron donor (reactions 10-11) [48, 50]. Trx-(SH)2 + protein-S2 → Trx-S2 + protein-(SH)2

(10)

Trx-S2 + NADPH+H+ ↔ Trx-(SH)2 + NADP+

(11)

7.3. Protein Disulfide Isomerase Protein disulfide isomerase (PDI) is an essential folding catalyst and chaperone primarily found in the endoplasmic reticulum, but also associated with other membranes. It is found in a wide range of species, including fungi, plants and animals. It is constitutively expressed in most tissues and organs and its cellular abundance is due to both its high expression and unusual stability. This 55 kDa protein is a member of a large family of dithiol/disulfide oxidoreductases, the thioredoxin superfamily. Its structure contains two double-cysteine redox-active sites, each within domains with high sequence similarity to thioredoxin, separated by two thioredoxin-related domains lacking reactive cysteines (Figure 4). PDI is the

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most versatile member of the superfamily, capable of catalyzing oxidation, reduction and disulfide isomerization. The two cysteines in the active centre can either exist in the dithiol (reduced) form or form an intramolecular disulfide (oxidized PDI). The redox state of the active site is generally maintained by gluthatione (GSH) and its disulfide (GSSG) [47].

8. Degradation of Oxidatively Damaged Proteins For the most part, oxidatively modified proteins are not repaired and must be removed by proteolytic degradation. To maintain the cellular function and integrity it is essential to have systems available that are able to recognize and degrade misfolded or damaged proteins in an efficient way, to prevent their aggregation and cross-linking. To prevent such accumulation of protein aggregates, several protein-degrading systems and enzymes have been evolved. This includes the lysosomal system and, most importantly, the proteasome.

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8.1. The Proteasomal System The proteasome is responsible for the degradation of misfolded, abnormal, and oxidized proteins. The proteasomal system also participates in a multitude of cellular processes like signal transduction, antigen processing, transcription, apoptosis and cell cycle progression. Considering the array of processes in which this system is involved, it is not surprising that the proteasome has a low specificity and a broad substrate spectrum. Eukaryotes have developed a complicated proteasomal system. The simplest form is the 20S „core proteasome‟ which plays also a central role in other proteasomal constructs. The 20S proteasome has a total molecular weight of 670-700 kDa and consists of two sets of 14 subunits forming a barrel-shaped structure with seven monomers per ring. Each of the subunits has a molecular weight ranging between 18 to 35 kDa. The two outer rings contain seven different -subunits, while the two inner rings are constructed out of seven different -subunits [51].

Figure 4. Oxidation of reduced PDI by glutathione disulfide. The reduced form of the active site cysteines of PDI react with glutathione disulfide (GSSG) to form the intramolecular disulfide and two molecules of reduced glutathione (according to Wilkinson et al. [47]).

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Figure 5. Structure of the mammalian 20S „„core‟‟ proteasome. The figure shows different perspectives of a simplified model structure of the subunit arrangement of the 20S „„core‟‟ proteasome. The a-rings are blue, the b-rings red. The whole proteasome has a diameter of about 100 Å and is 160 Å in height.

Three of these -subunits within each ring are catalytically active, the 1, 2 and 5. Each of the three subunits shows a different proteolytic activity: 1 exhibits a peptidylglutamyl-peptide-hydrolysing-activity (cleavage after acidic amino acids), a trypsin-like (after basic amino acids) activity at 2 and a chymotrypsin-like activity (after neutral amino acids) at the 5-subunit. The products of proteasomal degradation exhibit lengths between 1 and 35 amino acids, the average length of the products being 8–12 amino acids [52]. In eukaryotic cells the recognition and substrate entrance to the proteolytic inner chamber is only possible after contact with the -rings followed by conformational change that “open up” the gate formed by the 2, 3 and 4 subunits. The N-terminal ends of these subunits in the inactivated proteasome are turned to the axis of the proteasome, blocking the channel to the proteolytic chamber built by the -rings. The exposure of hydrophobic structures of misfolded or damaged unfolded proteins is assumed to „„activate‟‟ the proteasome by inducing of conformational changes in the gating structures of the -rings [53]. Another possibility is the attachment of regulatory subunits (like 11S or 19S) to the proteasomal -rings, resulting in increased activity and changed substrate specificity, caused by a maximal opening of the gated substrate channel by conformational changes of the blocking N-terminal ends. The 11S as well as the 19S regulator can be attached on one or on both sides of the 20S proteasome. Furthermore, a hybrid form with the 11S regulator on the one side and the 19S regulator on the other side has been identified. The attachment of the 19S regulator to each of the ends of the 20S proteasome forms the 26S proteasome [54]. Generally the proteasome acts in the following way: the damaged protein is recognized by the proteasome and enters through the opening in the middle of the -ring. Afterwards, the protein is processed in the catalytic chamber between the -rings and the resulting small polypeptides leave the proteasome through the opening of the second -ring [53]. While the 20S core proteasome is able to degrade proteins in an ATP- and ubiquitinindependent manner, the 26S proteasome is responsible for degradation of polyubiquitinated substrates. On this occasion the ATPase activity of the 19S regulator is used to facilitate unfolding of the ubiquitinated proteins and their translocation into the opening of the proteasome.

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Numerous studies in vitro as well as in vivo have been performed in order to investigate the influence of oxidation on the proteasomal protein turnover. In summary, all available experimental data suggest that the proteasome plays an important role in the removal of oxidized proteins from the cellular environment.

8.2. The Lysosomal System Lysosomes were first isolated from Christian de Duve in 1955 [55]. They were initially referred to as “lytic bodies”, being the equivalent to vacuoles in plants. The lysosomal system consists of numerous acidic vesicles (pH ~4-5) that constantly fuse and divide. Many different degrading enzymes, such as phosphatases, proteases, polysaccharidases, oligosaccharidases and lipid-hydrolysing enzymes are found in lysosomes, enabling the degradation of almost any cellular structure. Several proteins from mitochondria and both the endoplasmic/sarcoplasmic reticulum as well as foreign proteins from outside are degraded within lysosomes by a group of proteases called cathepsins, a protease family consisting of at least 16 members. Cathepsins are synthesized as inactive preproenzymes, glycosylated posttranslationally and directed towards the lysosomal compartment. However, during recent years it has become clear that cathepsins can also be released from the lysosomes where they can catalyze substrate cleavage [56]. Besides protein turnover, the cathepsins are also involved in a number of biological processes such as immune response, antigen processing and apoptosis [57]. An overview of different lysosmal cathepsins and their properties is shown in Table 2. Table 2. Lysosomal cathepsins and their properties [61-73]

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Cathepsin

*

Category*

Protease activity

Optimum pH * *

Occurence

A

serine protea se

exopeptidase

4.5-5.0

4.5-5.0

B

cysteine protea se

endopeptidase/exopeptidase

4.5-5.5

ubiquitous

C

cysteine protea se

exopetidase

5.0

ubiquitous

D

a spa rta te protease

endopeptidase

3.5-5.0

ubiquitous

E

a spa rta te protease

endopeptidase

3.0-4.0

cells of immune system (not in resting Blymphocytes), osteoclasts, gastric epithelial cells

F

cysteine protea se

endopeptidase

5.2-6.8

ubiquitous

H

cysteine protea se

endopeptidase/exopeptidase

5.4-6.8

ubiquitous

K

cysteine protea se

endopeptidase

5.5

osteoclasts, ovary

L

cysteine protea se

endopeptidase

4.5-5.5

ubiquitous

O

cysteine protea se

endopeptidase

acidic (lack of detailed information)

ubiquitous

S

cysteine protea se

endopeptidase

7.0

a ntigen-presenting cells

V

cysteine protea se

endopeptidase

4.0-5.7

thymus, testis

W

cysteine protea se

unknown

-

NK cells; CD8 + cytotoxic T cells

X

cysteine protea se

exopeptidase

5.0-6.0

ubiquitous

Subdivision according tothe the amino acidconfers that catalytic confersactivity catalytic activity. cording to amino acid that Substrate dependent. ** Substrate dependent

**

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In general, lysosomes are responsible for the digestion of macromolecules from phagocytosis, endocytosis and autophagy. Three different mechanisms of autophagy have been described for mammalian cells: macroautophagy, microautophagy and chaperonemediated autophagy [58-60] (see also Vol. II, Chapter 16). Macroautophagy involves the sequestration of organelles and soluble proteins in a double-membrane vesicle, called autophagosome. The outer membrane of the autophagosome fuses with a lysosome to form an autolysosome/autophagolysosome where the contents are degraded via lysosomal degrading enzymes. Macroautophagy is induced early in starvation to generate essential amino acids. Microautophagy is the transfer of cytosolic components into the lysosome by direct invagination of the lysosomal membrane into the lysosomal lumen. In chaperone-mediated autophagy only those proteins that have a consensus peptide sequence related to the pentapeptide KFERQ get recognized by the binding of an hsc70-containing chaperone/cochaperone complex. After unfolding, the substrate proteins are translocated into the lysosomal lumen for degradation with the assistance of a lysosomal chaperone (lys-hsc70) [74]. Although lysosomes are known to be major components of the cellular proteolytic machinery, the ability of lysosomal proteases to selectively recognize and degrade oxidatively damaged proteins is limited and the lysosomal protein degradation is mainly a nonselective process (only chaperone-mediated autophagy is very selective for substrates). However, there are some hints that monoubiquitination directs membrane proteins, like receptors, to the endosomal compartment and into lysosomes [75]. Lysosomes have also been viewed as storage places for waste products, such as lipofuscin (Vol. II, Chapter 16). Since lipofuscin is nondegradable, the transfer of active cathepsins and other hydrolases to lipofuscin-loaded lysosomes means a waste of enzymes in the futile attempt to degrade the pigment. In the event that lipofuscin-loaded lysosomes become plentiful, they may deplete lysosomes of sufficient enzymes and create a lack of degrading capacity [76].

9. Detection of Protein Damage A brief overview of procedures for the analysis of popular indices of protein oxidation is given here. Detailed information can be found in relevant literature.

9.1. Protein Carbonyls One of the most common methods to measure protein oxidation is the detection of protein carbonyls. Protein carbonyl content (PCC) is the most widely used marker of oxidative modification of proteins. Protein carbonyls can react specifically with 2,4-dinitrophenyl hydrazine (DNPH) and form the corresponding hydrazones (DNP), which have a peak absorbance around 360 nm. Although the hydrazone derivative of the protein bound carbonyl itself is colored, it is much more sensitive to detect those derivates with antibodies (antiDNP), e.g. to exclude interference by proteins containing chromophores that also have absorbance near 360 nm. To identify specific proteins that are carbonylated, DNPH-treated proteins can further be separated by gel electrophoresis. After being transferred from the gel

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to a membrane, protein-bound DNP in individual proteins can again be explored by the use of anti-DNP antibodies. By using PCC as a marker, it could be demonstrated that oxidative damage to proteins correlates well with aging and the severity of certain diseases. In view of the fact that the formation of protein carbonyl groups is orders of magnitude greater than other oxidative modifications, the level of protein carbonyl groups has often been employed for the quantification of generalized protein oxidation [77].

9.2. Protein Thiol Groups To identify specific proteins exhibiting loss of thiol groups due to oxidative damage, the thiols can be labeled with various compounds, e.g. DTNB (Ellmann‟s reagent) [78] or [14C] iodoacetamide [79]. All methods are based on the measurement of the difference between the normal signal and the (lower) signal in the oxidized sample which reflects loss of thiol groups.

9.3. Protein-bound Nitrotyrosine

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The availability of anti-nitrotyrosine antibodies provides a sensitive method for detection of protein-bound nitrotyrosine, e.g. immunoblots, immunohistochemistry or enzyme-linked immunosorbent assay (ELISA). Although, these methods are widely used, their reliability depends on many factors, including the quality of the antibody [80-82].

Conclusions Considering the manifold possibilities of protein primary or secondary oxidation and further modification by reactive molecules, different lines of antioxidant defense exist to protect cells from oxidative proteotoxic stress. Since oxidative protein modification is an inevitable by-product of cellular metabolism, several systems developed during evolution that are able to reduce or repair the damage, or to recognize and remove damaged proteins. The first line of defense comprises antioxidant mechanisms which reduce the probability of oxidative protein modification. The second line consists of enzymatic repair systems including the thioredoxin/thioredoxin reductase, the glutaredoxin-/glutathione-/glutathione reductase-system and the methionine sulfoxide reductases. Since cysteine and methionine are the only two amino acids that can be enzymatically reduced, strong and efficient systems are needed to recognize and remove oxidatively-modified proteins and thereby preserve cellular functionality. This third line of defense contains the two most important machineries for the degradation of oxidized proteins, the lysosomal and the proteasomal system (with the latter responsible for clearing about 90% of damaged cellular proteins).

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[59] Todde V, Veenhuis M, van dK, I. Autophagy: principles and significance in health and disease. Biochim Biophys Acta 2009; 1792(1):3-13. [60] Cuervo AM, Bergamini E, Brunk UT, Droge W, Ffrench M, Terman A. Autophagy and aging: the importance of maintaining "clean" cells. Autophagy 2005; 1(3):131140. [61] Chikuma T, Ogura Y, Kasamatsu M et al. High-performance liquid chromatographic-fluorimetric assay for cathepsin A (lysosomal protective protein) activity. J Chromatogr B Biomed Sci Appl 1999; 728(1):59-65. [62] Zavasnik-Bergant T, Turk B. Cysteine cathepsins in the immune response. Tissue Antigens 2006; 67(5):349-355. [63] Wex T, Buhling F, Wex H et al. Human cathepsin W, a cysteine protease predominantly expressed in NK cells, is mainly localized in the endoplasmic reticulum. J Immunol 2001; 167(4):2172-2178. [64] Linebaugh BE, Sameni M, Day NA, Sloane BF, Keppler D. Exocytosis of active cathepsin B enzyme activity at pH 7.0, inhibition and molecular mass. Eur J Biochem 1999; 264(1):100-109. [65] Vanha-Perttula T, Hopsu VK, Sonninen V, Glenner GG. Cathepsin C activity as related to some histochemical substrates. Histochemie 1965; 5(2):170-181. [66] Simon DI, Ezratty AM, Loscalzo J. The fibrin(ogen)olytic properties of cathepsin D. Biochemistry 1994; 33(21):6555-6563. [67] Kageyama T. Procathepsin E and cathepsin E. Methods Enzymol 1995; 248:120-136. [68] Wang B, Shi GP, Yao PM, Li Z, Chapman HA, Bromme D. Human cathepsin F. Molecular cloning, functional expression, tissue localization, and enzymatic characterization. J Biol Chem 1998; 273(48):32000-32008. [69] Bossard MJ, Tomaszek TA, Thompson SK et al. Proteolytic activity of human osteoclast cathepsin K. Expression, purification, activation, and substrate identification. J Biol Chem 1996; 271(21):12517-12524. [70] Klemencic I, Carmona AK, Cezari MH et al. Biochemical characterization of human cathepsin X revealed that the enzyme is an exopeptidase, acting as carboxymonopeptidase or carboxydipeptidase. Eur J Biochem 2000; 267(17):54045412. [71] Schwartz WN, Barrett AJ. Human cathepsin H. Biochem J 1980; 191(2):487-497. [72] Pangkey H. Purification and characterization of cathepsin S from hepatopancreas of carp Cyprinus carpio. Fisheries Science 2000; 66(6):1130-1137. [73] Bromme D, Li Z, Barnes M, Mehler E. Human cathepsin V functional expression, tissue distribution, electrostatic surface potential, enzymatic characterization, and chromosomal localization. Biochemistry 1999; 38(8):2377-2385. [74] Majeski AE, Dice JF. Mechanisms of chaperone-mediated autophagy. Int J Biochem Cell Biol 2004; 36(12):2435-2444. [75] Hofmann K, Falquet L. A ubiquitin-interacting motif conserved in components of the proteasomal and lysosomal protein degradation systems. Trends Biochem Sci 2001; 26(6):347-350. [76] Kurz T, Terman A, Gustafsson B, Brunk UT. Lysosomes and oxidative stress in aging and apoptosis. Biochim Biophys Acta 2008; 1780(11):1291-1303.

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[77] Buss H, Chan TP, Sluis KB, Domigan NM, Winterbourn CC. Protein carbonyl measurement by a sensitive ELISA method. Free Radic Biol Med 1997; 23(3):361366. [78] Ellman GL. Tissue sulfhydryl groups. Arch Biochem Biophys 1959; 82(1):70-77. [79] Sethuraman M, McComb ME, Heibeck T, Costello CE, Cohen RA. Isotope-coded affinity tag approach to identify and quantify oxidant-sensitive protein thiols. Mol Cell Proteomics 2004; 3(3):273-278. [80] ter Steege JC, Koster-Kamphuis L, van Straaten EA, Forget PP, Buurman WA. Nitrotyrosine in plasma of celiac disease patients as detected by a new sandwich ELISA. Free Radic Biol Med 1998; 25(8):953-963. [81] Inoue H, Hisamatsu K, Ando K, Ajisaka R, Kumagai N. Determination of nitrotyrosine and related compounds in biological specimens by competitive enzyme immunoassay. Nitric Oxide 2002; 7(1):11-17. [82] Herce-Pagliai C, Kotecha S, Shuker DE. Analytical methods for 3-nitrotyrosine as a marker of exposure to reactive nitrogen species: a review. Nitric Oxide 1998; 2(5):324-336.

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 7

Lipid Peroxidation Angel Catalá* Instituto de Investigaciones Fisicoquímicas Teóricas y Aplicadas, (INIFTA-CCT La Plata-CONICET), Facultad de Ciencias Exactas, Universidad Nacional de La Plata, Argentina

Abbreviations

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PUFAs, polyunsaturated fatty acids; C18:1n-9, oleic acid; C18:2n-6, linoleic acid; C18:3n-3, -linolenic acid; C20:4n-6, arachidonic acid; C22:6n-3 docosahexaenoic acid; LH, lipid molecules; R●, radical; L●, lipid radical; LOO●, lipid peroxyl radical; LO●, lipid alkoxyl radical; LOOH, lipid hydroperoxide; Me, metal ions; NRP, non-radical products; 4-HNE, 4-hydroxy-2-nonenal; 4-HHE, 4-hydroxy-2-hexenal; MDA, malondialdehyde; TBARS, thiobarbituric reactive species; IsoPs, isoprostanes; *

Corresponding author. Prof. Dr. Angel Catalá. INIFTA, CONICET, CIC. . CC 16, Sucursal 4. 1900- La Plata, Argentina.. Tel 54-221-425-7430 54-221-425-7291 (Ext. 105);. Fax 54-221-425- 4642;. Email: [email protected] The author is Member of Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET) Argentina.

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Angel Catalá IsoFs, isofurans; NeuroPs, neuroprostanes; ROS, reactive oxygen species; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PI, phosphatidylinositol; PS, phosphatidylserine; COX, cyclooxygenase.

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1. Introduction The onset of lipid peroxidation within cellular membranes is associated with changes in their physicochemical properties and functional inactivation of proteins located in the membrane environment. Polyunsaturated fatty acids (PUFAs) and their metabolites have diverse physiological roles, among others in the context of energy metabolism, membrane structure, cell signaling and regulation of gene expression. Lipids containing PUFAs are prone to free radical–initiated oxidation and may participate in chain reactions that increase injury to biomolecules. Lipid peroxidation, which often leads to generation of lipid hydroperoxides, is a result of oxidative stress. Hydroperoxides are generally reduced to alcohols by glutathione peroxidases. Nevertheless, these enzymes are diminished in certain diseases resulting in a transitory augmentation of lipid hydroperoxides that favors their decomposition into several products, including hydroxy-alkenals. The best characterized are 4-hydroxy-2-nonenal (4-HNE) and 4-hydroxy-2-hexenal (4-HHE), which originate from lipid peroxidation of n-6 and n-3 fatty acids, respectively. Compared to free radicals, these aldehydes are moderately stable and can diffuse within the cell or be extruded extracellularly to attack distant targets. They display high reactivity with biomolecules, such as proteins, DNA and phospholipids, generating various intra- and intermolecular covalent adducts. At the membrane level, proteins and amino lipids can be covalently modified by hydoxyalkenals, damaging membrane structure. In addition, these aldehydes can act as bioactive molecules in physiological and/or pathological events. This Chapter provides an outline on the intracellular production of hydroxy-alkenals and oxidized phospholipids, their activity as intracellular messengers and the mechanisms by which they modulate cell signaling and affect transcription factors and gene expression.

2. Origin and Physiological Role of PUFAs PUFAs and their metabolites have various structural, metabolic, signaling and regulatory functions. Mammalian tissue requirements for PUFAs cannot be met solely by de novo biosynthesis. Animals are absolutely dependent on plants for providing the two major precursors of the n-6 and n-3 fatty acids, C18:2n-6; linoleic and C18:3n-3; -linolenic acids, and convert them to fatty acids containing three to six double bonds [1]. Some of the animal daily needs for long chain PUFAs are covered by the diet. However, most of the long chain PUFAs found in animal tissues are synthesized by conversion of essential plant-derived fatty

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acid precursors (C18:2n-6 and C18:3n-3) to their respective C20- and C22-carbon polyenoic products via elongations and desaturations. Unsaturation of a fatty acid chain is a major determinant of the melting temperature of triglycerides, as well as the fluidity of biological membranes that are made of a phospholipid bilayer. Thus, fatty acid desaturases, which introduce a double bond into a long-chain fatty acid, are evolutionarily conserved across species [2]. Long chain PUFAs such as arachidonic acid (C20:4n-6) and docosahexaenoic acid (C22:6n-3) are also crucial for various biological functions [3, 4]. They confer fluidity, flexibility and selective permeability to cellular membranes in higher eukaryotes, and affect other cellular and physiological processes in both plants and animals. Animal biosynthesis of high polyunsaturated fatty acids from linoleic, -linolenic and oleic acids is mainly modulated by the delta6 and delta5 desaturases through dietary and hormonally-stimulated mechanisms [5].

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3. Lipid Peroxidation in Cellular Membranes All cellular membranes are especially vulnerable to oxidation due to the presence of abundant PUFAs [6]. The dynamic growth of this field over the past years is illustrated in many excellent reviews [4, 7-18]. Lipid peroxidation of PUFAs, which can be enzymatic or non-enzymatic, is a major mechanism of cell injury during oxidative stress. Non-enzymatic lipid peroxidation can be initiated in vitro by adding ascorbate in the presence of oxygen and Fe2+ or Fe3+ ions to tissue preparations such as whole homogenates, mitochondria, microsomes or nuclei [19-21]. Among cellular macromolecules, PUFAs exhibit the highest sensitivity to oxidative damage. Their sensitivity grows exponentially with increasing numbers of double bonds per fatty acid molecule [22]. Membrane phospholipids are particularly vulnerable to oxidation not only because of their high PUFA content, but also due to their close proximity to non-enzymatic and enzymatic systems generating reactive oxygen species (ROS) and other oxidants. Figure 1 depicts the mechanism for non-enzymatic lipid peroxidation of docosahexaenoic acid 22:6 n-3, a major dietary n-3 PUFA and a major structural lipid of retinal photoreceptor outer segment membranes.

4. Mechanisms of Lipid Peroxidation Oxidative stress resulting from an imbalance between prooxidant/antioxidant systems, promotes damage to cellular biomolecules such as proteins (Chapter 6), nucleic acids (Chapter 9), structural carbohydrates and lipids [23]. Among these targets, the peroxidation of lipids is particularly damaging because of its propensity to propagate free radical reactions. The general process of lipid peroxidation consists of three stages: initiation, propagation, and termination [4, 19]. The initiation phase includes hydrogen atom abstraction. Several species can abstract the first hydrogen atom, including the radicals hydroxyl (HO●), alkoxyl (RO●), peroxyl (ROO●) and possibly HO2●, but not H2O2 or O2−● [24].

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Figure 1. Mechanism of lipid peroxidation of phospholipids containing docosahexaenoic acid (22:6 n-3). R1 = fatty acid, R2 = fragmentation products of fatty acid oxidation.

Membrane lipids, mainly phospholipids, containing PUFAs are highly susceptible to peroxidation because abstraction of a hydrogen atom from a methylene (-CH2-) group, leaves an unpaired electron in the vicinal carbon (-CH●-). The presence of a double bond in the fatty acid weakens the C–H bonds in the carbon neighboring the double bond and thus, facilitates the abstraction of H●. The initial reaction of HO● with PUFAs produces a lipid radical (L●), which in turn reacts with molecular oxygen to form a lipid peroxyl radical (LOO●). The LOO● can abstract hydrogen from an adjacent fatty acid to produce a lipid hydroperoxide (LOOH) and a second lipid radical [4, 19]. The LOOH may undergo reductive cleavage by reduced metals, such as Fe++, producing lipid alkoxyl radical (LO●). Both alkoxyl and peroxyl radicals stimulate the chain reaction of lipid peroxidation by abstracting additional hydrogen atoms [25]. Peroxidation of lipids can damage the membrane architecture, causing changes in fluidity and permeability, alterations in ion transport and inhibition of metabolic processes [16]. Injury to mitochondria induced by lipid peroxidation can promote further ROS generation [26]. Furthermore, LOOH can decompose to reactive aldehyde products, including

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malondialdehyde (MDA), 4-HNE, 4-HHE and acrolein [27-29] (Figure 1); these reactions are favored by the presence of reduced metals or ascorbate. In fact, such oxidation products and a series of hydroxyl-alkenals have been detected by gas chromatography-mass spectrometry (GC-MS) from low-density lipoprotein undergoing copper-catalyzed oxidation [30]. Another publication reported additional products derived from the oxidation of erythrocyte membrane in vitro [31]. Assays to measure lipid peroxidation are commonly employed for the assessment and quantification of radicalinduced damage [15, 32-37]. There are two broad outcomes to lipid peroxidation: structural damage to membranes and generation of secondary products. Membrane damage is caused by fragmentation of fatty acyl chains, lipid-lipid and lipid-protein cross-linking, and endocyclization reactions producing isoprostanes and neuroprostanes. Thus, peroxidation of long chain PUFAs such as arachidonic (C20:4n-6) and docosahexaenoic (C22:6n-3) acids produce isoprostanes and neuroprostanes, respectively. These processes promote changes in the biophysical properties of membranes that may have profound effects on the activity of membrane-bound proteins. The consequences of PUFA peroxidation are severe: damage of membranes, inactivation of proteins, impairment of cellular division, etc [38-40].

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5. Enzymatic Lipid Peroxidation by Lipoxygenases Lipoxygenases comprise a family of lipid peroxidation enzymes that oxygenate free and esterified PUFAs to the corresponding hydroperoxy derivatives. They are widely distributed in both plant and animal kingdoms [41, 42] and classified into three major isoforms with respect to positional specificity of arachidonic acid oxygenation: 5-, 12- or 15-lipoxygenases. Arachidonic acid is oxidatively metabolized by rat liver microsomes at a rate of approximately 5 nmol per min per mg of protein at 25 C. This reaction is dependent on the presence of NADPH and oxygen. Studies with various inhibitors indicate a role for membrane-bound cytochrome P-450 in the transformation of arachidonic acid to a mixture of hydroxy acid derivatives. The stoichiometry of the reaction conforms to that of a monooxygenase reaction i.e., one molecule of NADPH is oxidized per molecule of consumed oxygen, suggesting a reaction mechanism different from that proposed for lipid peroxidation reactions. No evidence for formation of prostaglandin-like metabolites has been obtained. The diene character of some of the resulting metabolites suggests another role for cytochrome P450, the participation in hydrogen abstraction reactions for substrate activation [43].

6. Non-enzymatic Lipid Peroxidation Non-enzymatic lipid peroxidation is a free radical-driven chain reaction in which one radical can induce the oxidation of a large number of lipid molecules (LH) mainly phospholipids containing PUFAs [46]. The chain reaction is initiated by the abstraction of a hydrogen atom from a cis-interrupted methylene group of a PUFA residue [44]. Monounsaturated and saturated fatty acids are much less reactive and usually do not participate in lipid peroxidation. Thus, fatty acids with one or no double bonds can undergo

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oxidation but not a chain lipid peroxidation process; e.g., oleic acid with 18 carbon atoms and 1 double bond (C18:1 n9).

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Figure 2. Non-enzymatic lipid peroxidation.

Initiation of non-enzymatic lipid peroxidation (Figure 2) is usually performed by a radical (R●) of sufficient reactivity (a). Molecular oxygen rapidly adds to the carbon-centered lipid radical (L●) formed in this process, yielding lipid peroxyl radical (LOO●) (b), which, in turn, can abstract hydrogen from another PUFA (c). Reaction (c) is known as a “propagation reaction”, implying that one initiating hit can result in the conversion of several PUFA molecules to lipid hydroperoxides. Lipid hydroperoxide (LOOH) is the first, relatively stable, product of the lipid peroxidation reaction [45]., A “termination reaction” limits the extent of this process, yielding non-radical products (NRP) (d). LOOH may give rise to radicals capable of reinitiating lipid peroxidation by redox-cycling of transition metal ions [(e) and (f)]. Lipid hydroperoxides may yield a large variety of products, including short and long chain aldehydes and phospholipid and cholesterol ester core aldehydes (many of which can be utilized for assays to quantify lipid peroxidation [46]).

7. Peroxidation Products of Membrane Phospholipids Peroxidation of fatty acyl groups occurs mostly in membrane phospholipids and alters the physicochemical properties of the lipid bilayers, resulting in cellular dysfunction. A variety of lipid byproducts are produced as a result of lipid peroxidation (Table 1), which may have either adverse or beneficial biological effects.

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Table 1. Compounds generated during lipid peroxidation of membrane phospholipids

Several in vitro markers of oxidative stress are available; however most of them are of limited value because they lack sufficient sensitivity and/or specificity, or require invasive methods in vivo. Isoprostanes (IsoPs) are prostaglandin (PG)-like compounds produced independently of cyclooxygenase (COX) enzymes, primarily by free radical-induced peroxidation of arachidonic acid. F(2)-IsoPs are a group of 64 compounds isomeric in structure to COX-derived PGF(2 alpha). Other products of the IsoP pathway are also formed in vivo by rearrangement of labile PGH(2)-like IsoP intermediates, including E(2)-, D(2)IsoPs, cyclopentenone-A(2)-, J(2)-IsoPs and highly reactive acyclic-ketoaldehydes (isoketals). Measurement of F(2)-IsoPs is a reliable approach to assess oxidative stress status in vivo, providing an important tool to explore the role of oxidative stress in the pathogenesis of human disease. Moreover, F(2)-IsoPs and other products of the IsoP pathway exert potent biological actions via receptor-dependent and/or independent mechanisms, and therefore may be pathophysiological mediators of disease. The formation of highly reactive cyclopentenone isoprostane compounds (A3/J3-isoprostanes) in vivo from eicosapentaenoic acid was described recently [47]. Oxidation of docosahexaenoic acid, an abundant unsaturated fatty acid in the central nervous system, results in the formation of IsoP-like compounds, termed neuroprostanes. Aldehydic molecules generated during lipid peroxidation trigger several biological effects. Compared to free radicals, these molecules are relatively stable and can diffuse both within cells and extracellularly to attack distal targets. Aldehydes are highly reactive towards bio-molecules, such as proteins, DNA, and phospholipids, generating a host of intra- and intermolecular covalent adducts. Moreover, they can also act as bioactive molecules under physiological and/or pathological conditions. At very low and nontoxic concentrations, aldehydes affect and regulate several cellular functions, including signal transduction, gene expression and cell proliferation [13]. Ketoaldehydes, including MDA and glyoxal, comprise an important group of reactive aldehydes originating from lipid peroxidation. MDA is highly abundant and has been shown to disturb aminophopholipid organization in the membrane bilayer of erythrocytes [48].

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8. Generation of Oxidized Phospholipids by Lipid Peroxidation Biomembranes contain different phospholipid classes (head group heterogeneity), subclasses (acyl, alkyl chains) and species (chain length and unsaturation degree). Phosphatidylcholine (PC) is the main phospholipid in all mammalian cells (40–50%) and thus, most oxidized phospholipids detected in mammalian tissues possess a choline moiety. However, oxidized phosphatidylethanolamine (PE) was recently found in the retina, a tissue abundant in ethanolamine lipids [49] and enriched for docosahexaenoic acid [50]. The surface of apoptotic cells contains oxidized phosphatidylserine (PS) [51]. In eukaryotic phospholipids, the sn-1 position is either linked to an acyl residue via an ester bond or to an alkyl residue via an ether bond, whereas the sn-2 position almost exclusively contains acyl residues. The highly oxidizable n-3 and n-6 PUFAs are preferably bound to the sn-2 position of glycerophospholipids. Thus, most of the oxidized phospholipids are modified at this position. The sn-1 position of glycerol is usually occupied by a saturated fatty acid. By contrast, plasmalogens (alkenylacylglycerophospholipids) contain a vinyl ether bond in position sn-1 and, as a consequence, are susceptible to oxidative modifications. The differences in chemical structure of various types of phospholipids determine the physical properties of the membrane. PC tends to form bilayers with little curvature, while PE imposes a negative curvature on lipid bilayers [52]. Conversely, introduction of the micelleforming LPC into a PC membrane results in a positive curvature. In addition to the polar head groups, the polarity, length and unsaturation of the phospholipid acyl chains also have an impact on physical membrane properties. Thus, phospholipid oxidation products (Figure 2) are very likely to modify the properties of biological membranes because their polarity and shape may differ significantly from those of their parent molecules. Thus, they may alter lipid–lipid and lipid–protein interactions and, consequently, also membrane protein functions. When the sn-2 fatty acids of phospholipids are oxidized by radicals, several different types of oxidative products are formed. These include phospholipids containing fatty acid oxidation products (usually referred to as oxidized phospholipids), lysophospholipids, and fragmentation products of fatty acid oxidation. Some of these products (lysophosphatidic acid or lysophospholipids) can be formed both enzymatically and non-enzymatically. Work from many laboratories has identified a variety of phospholipid oxidation products and confirmed bioactivities of these phospholipids on vascular endothelial cells, leukocytes and platelets. Despite the large number of fatty acid oxidation products that have already been identified, it is almost certain that many other bioactive oxidized phospholipids remain to be discovered. There are several important issues in the field of bioactive phospholipids, including preparation, identification, quantification and biological testing. Mass spectrometry (MS) and tandem mass spectrometry (MS/MS) using the soft ionization methods (electrospray and matrix-assisted laser desorption ionization) are among the finest approaches for the study of oxidized phospholipids. Product ions of oxidized phospholipids, allow detection of changes in the fatty acyl chain and specific features such as presence of new functional groups in the molecule and their location within the fatty acyl chain [53].

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9. Generation of Hydroxy-alkenals by Lipid Peroxidation of n-3 and n-6 PUFAs Lipids containing PUFAs are susceptible to free radical–initiated oxidation and can contribute in chain reactions that amplify damage to biomolecules, as described above. As a consequence of oxidative stress, various aldehydes are formed during decomposition of lipid hydroperoxides in vivo. Some of them, such as 4-HNE and 4-HHE (Figure 3), are highly reactive and may be considered as “second toxic messengers”, which disseminate and augment initial free radical events. 4-HNE is the main aldehyde formed during lipid peroxidation of n-6 polyunsaturated fatty acids, such as linoleic acid C18:2 n-6 and arachidonic acid C20:4 n-6. It was identified three decades ago as a cytotoxic aldehyde formed during the NADPH/Fe++-induced peroxidation of liver microsomal lipids [54]. 4-HNE has been associated with several pathological conditions and serves as a marker of oxidative stress. Lipid peroxidation of n-3 polyunsaturated fatty acids such as -linolenic acid C18:3 n-3 and docosahexaenoic acid C22:6 n-3 generates 4-HHE, a closely related molecule, which is a potential mediator of mitochondrial permeability transition [55].

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10. Biomarkers of Lipid Peroxidation: Isoprostanes, Isofurans and Neuroprostanes As discussed above, lipid peroxidation results in the breakdown of PUFAs into smaller fragments. On the basis of structural features, the reactive short-chain aldehydes generated during this process can be classified into three families: 2-alkenals, 4-hydroxy-2-alkenals, and ketoaldehydes (for details see [13]). Some of the chemically and metabolically stable oxidation products are useful as in vivo biomarkers of lipid peroxidation. These include the isoprostanes (IsoPs) and isofurans (IsoFs), derived from arachidonic acid C20:4 n-6, and neuroprostanes (NeuroPs), derived from docosahexaenoic acid C22:6 n-3 [56]. IsoPs are formed in situ by peroxidation of C2:4n-6 bound to phospholipids. Two major structural isomers of IsoPs are formed from a common endoperoxide intermediate: F2-IsoPs by reduction and D2/E2-IsoPs by isomerization. The ratio of F-ring to D/E-ring compounds reflects the reducing environment in which IsoPs form; the greater the reducing equivalents the larger the F- to D/E-ring ratio [57]. Fessel et al. [58] demonstrated an oxygen insertion step that diverts intermediates in the IsoP pathway to form tetrahydrofuran ring-containing compounds, termed IsoFs. Like IsoPs, IsoFs are chemically and metabolically stable and thus qualify as in vivo biomarkers. In the mammalian retina, 15-F2t-isoprostanes and its metabolite (2,3-dinor-5,6-dihydro15-F2t-IsoP) evoke vasoconstriction by stimulating TXA2 formation from neurovascular cells [59]. Equivalent inhibition of the constrictor effect of 8-iso-PGF2α by the PLA2 blocker OPPC, the COX inhibitor indomethacin, and the thromboxane synthase inhibitor CGS-12970 suggests that 8-iso-PGF2α acts on the synthesis of thromboxane, rather than on its receptors, by stimulating the release of arachidonic acid, which is metabolized by COX into thromboxane-dominated prostanoids. The data further suggest that 8-iso-PGF2α elicits retinal

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vasoconstriction by releasing endothelin and the prostanoid thromboxane from retinal parenchymal and endothelial cells in response to Ca2+ entry [59, 60]. The technology for measuring IsoPs has been extended to include quantification of oxidation products of C22:6 n-3 that have been named NeuroPs. The same nomenclature for NeuroPs and IsoPs has been adopted; for example, the reduced form has an F-ring containing four unsaturated bonds in the side chains, or F4-NeuroPs [61]. In contrast to C20:4 n-6, that is evenly distributed in all cell types and areas of the brain, C22:6 n-3 is highly concentrated in neuronal membranes [62]. Thus, NeuroPs provide a unique molecular tool to quantify oxidative damage in these membranes, which may be important in the context of neurological diseases.

Figure 3. Reactive hydroxy-alkenals generated during lipid peroxidation of n-3 and n-6 polyunsaturated fatty acids.

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Table 2. Analytical methods for studying lipid peroxidation in membranes

11. Analytical Methods for Studying Lipid Peroxidation in Membranes Many methods are available to quantify each of the three stages of lipid peroxidation (initiation, propagation, and termination [63]) and some of them are summarized in Table 2.

12. Covalent Modification of Proteins and Aminophospholipids by Lipid Peroxidation Products Oxidative stress is implicated in many pathophysiological states, such as aging (Vol. II, Chapter 23), atherosclerosis, diabetes, and neurodegenerative disorders (Vol. III, Chapters 810). ROS attack and modify diverse biological substrates, including PUFAs, leading to lipid

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hydroperoxide formation [45]. Hydroperoxides are normally reduced to alcohols by glutathione peroxidases [76] (see also Vol. II, Chapter 8). Nevertheless, the activities of these enzymes decrease in aging [77] and diabetes [78, 79], resulting in temporary accumulation of lipid hydroperoxides; these eventually undergo degradation into several products, including the hydroxy-alkenals 4-HNE [27] and 4-HHE [80].

Figure 4. Covalent modifications of amino-phospholipids and proteins by hydroxy-alkenals during lipid peroxidation of biological membranes.

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Lipid decomposition products can covalently modify proteins and amino lipids (Figure 4). Aliphatic aldehydes (alkanals), such as 1-hexanal or 1-nonanal, may attack the Nε-amino groups of lysine residues in proteins to form Schiff bases, while α,β-unsaturated aldehydes (alkenals), such as acrolein or 4-HNE, react with lysine, cysteine and histidine residues through Michael-type addition [27, 81]. Lipid hydroperoxides [82] and keto fatty acids (products of lipoxygenase-catalyzed reactions) [42], may also react with proteins [83, 84]. In addition, pyrrole compounds from long chain epoxides and lysine have been identified [85]. Nevertheless, the molecular mechanisms of protein modification by lipid hydroperoxides remains incompletely understood. Recent work provided evidence that ethanolamine phospholipids may form adducts with hydroxy-alkenals in situ [86].

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13. Damage of Cellular Membranes by Lipid Peroxidation Oxidative-stress induced peroxidation of membrane lipids modifies the physical properties of cellular membranes [87]. These include permeability to diverse solutes and the packing of lipids and proteins, which are crucial for proper membrane function. Considering that damage of cellular membranes is associated with aging and disease [88], many studies have focused on understanding structural and functional implications of lipid peroxidation. The complexity of biological membranes often necessitates the employment of model systems, such as lipid vesicles (liposomes) for lipid peroxidation studies. This approach is powerful and has yielded valuable insights but also has limitations, because peroxidation of liposomes may differ from that of biological membranes. As discussed earlier, PUFAs of membrane phospholipids are particularly susceptible to peroxidation and undergo significant modifications, including the rearrangement or loss of double bonds and, in some cases, the reductive degradation of lipid acyl side chains [68, 89, 90]. Lipid hydroperoxides also accumulate in the bilayer and further contribute to changes in the structural organization and packing of membrane lipid components [75]. Many biophysical consequences of these structural modifications have been well characterized and include changes in membrane fluidity [91-93] increased membrane permeability [94-96] alteration of membrane thermotropic phase properties [97-99] and changes in membrane protein activity [100-107].

14. In Vivo Reactivity of Hydroxy-Alkenals Aldehydic molecules generated during lipid peroxidation are moderately stable, highly reactive and can cause cell damage [27]. Because of a conjugated double bond between the α and β carbons, the γ carbon of these aldehydes is electron-deficient and reacts readily with nucleophilic molecules such as thiols and amines. The central nervous system is particularly vulnerable to oxidative stress due to the high rate of oxygen consumption and the presence of high levels of PUFAs (see Vol. III, Chapter 10). Thus, there has been much focus on possible roles of lipid peroxidation-derived

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aldehydes in contributing to neuronal dysfunction in neurodegenerative diseases associated with oxidative stress. Among such aldehydes, 4-HNE appears to have an important pathophysiological action. At modest concentrations, 4-HNE induces apoptosis [108, 109] impairs proteasomal function [110, 111] and impacts signal transduction by modulating adenylate cyclase, JNK, protein kinase C (PKC), and caspase 3. However, at lower concentrations, 4-HNE appears to promote cell proliferation [13, 112]. All biological activities attributed to 4-HNE reflect its capacity to bind covalently to protein targets and modulate their activity [113]; many such targets remain to be elucidated.

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15. 4-HNE Regulates Mitochondrial Uncoupling Mitochondria are the main intracellular producers of ROS in most cells and, in addition, important targets for their damaging effects (see Vol. II, Chapter 15). Mitochondrial oxidative damage may contribute to various pathological conditions, including neurodegenerative diseases, ischaemia/reperfusion injury and inflammatory disorders [27]. At least part of mitochondrial damage during oxidative stress is attributed to lipid peroxidation via superoxide (O2.-), involving generation of 4-HNE (but also 4-HHE and MDA) [27, 114116]. Mitochondrial proteins are targets of 4-HNE adduct formation, that leads to inactivation of the 2-oxoglutarate dehydrogenase and pyruvate dehydrogenase complexes, cytochrome c oxidase and NADH-linked respiration [117-119]. A model for activation of mitochondrial carriers (UCPs and ANT) by superoxide, through initiation of lipid peroxidation, has been proposed [120, 121]. According to this model,, 4-HNE induces mitochondrial uncoupling by specific interactions with the uncoupling proteins UCP1, UCP2 and UCP3, and with the adenine nucleotide translocase (ANT). These proteins are members of a large family of at least 35 anion carriers present in the mitochondrial inner membrane [122]. Mild uncoupling decreases mitochondrial production of ROS, which can subsequently engender 4-HNE production [123]. In this negative feedback loop, 4-HNE-dependent signaling decreases ROS production via uncoupling, which may serve to regulate mitochondrial production of superoxide. The employment of phenylbutylnitrone, a mitochondrially-targeted spin trap that reacts rapidly with carbon-centered radicals but not superoxide or lipid peroxidation products, prevented activation of UCPs by superoxide but did not block activation by 4-HNE. This suggests that superoxide and lipid peroxidation products share a common pathway for the activation of UCPs [124].

16. Hydroxy-alkenals as Second Messengers An increasing body of evidence supports a function of ROS as second messengers in signal transduction and gene regulatory pathways [125] (see also Vol. II, Chapters 12-14). Likewise, it appears that lipid peroxidation products, such as 4-HNE, also possess signaling capacity. Under physiological conditions, lipid peroxidation predominantly occurs in cells that are not rapidly proliferating, and steady-state cellular levels of 4-HNE range from 0.1– 0.3 mM. However, in response to oxidative insults, 4-HNE accumulates in membranes to

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reach concentrations approaching 5 mM [13]. At concentrations akin to those “physiologically” found in normal cells and plasma, 4-HNE appears to modulate cellular functions, gene expression and biochemical pathways, without cytotoxic effects [112] and for this reason, it is viewed by many researchers as an intracellular signaling intermediary rather than a waste byproduct of lipid peroxidation [13, 126]. Earlier results demonstrated that 4-HNE inhibits proliferation and promotes differentiation of leukemic cells [127, 128], while it elicits antiproliferative and proapoptotic responses in other cell types [129]. Nuclear receptors control gene expression by various mechanisms involving both activation and repression of DNA transcription [130-132]. Peroxisome proliferator-activated receptors (PPARs) are ligand-activated transcription factors belonging to the nuclear hormone receptor super family [133]. Following the initial isolation of PPARα as the receptor mediating peroxisome proliferation in rodent hepatocytes [134], two related isotypes, PPARβ and PPARγ have been characterized [135]. PPARs are sensors that modulate gene expression by integrating various lipid signals [136]. Pizzimenti et al [137], uncovered a connection between 4-HNE and PPAR pathways in leukemic cell growth and differentiation, providing early evidence for the involvement of a lipid peroxidation product in signaling. More recently, Cerbone et al [138] showed that 4-HNE and PPAR ligands affect proliferation, differentiation, and apoptosis in colon cancer cells. 4-HNE was also reported to modulate expression of the oncogenes c-myc and c-jun in K562 and HL-60 cells[ 140, 141]; both c-myc and c-jun regulate cell proliferation and survival, further reinforcing the concept for a signaling function of hydroxy-alkenals.

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Conclusions This Chapter provides an outline of pathways of lipid peroxidation and their biological implications. Membrane phospholipids containing PUFAs are particularly susceptible to oxidation and can contribute to chain reactions that amplify damage to biomolecules. Lipid peroxidation often occurs in response to oxidative stress, and a great diversity of phospholipid oxidation products and aldehydes are formed from the decomposition of lipid hydroperoxides. Bioactivities of these products on vascular endothelial cells, leukocytes, and platelets have been described. Some of the resulting aldehydes, such as 4-HNE, are highly reactive and may covalently modify proteins and amino lipids and damage biological membranes. At lower concentrations, however, 4-HNE and possibly other hydroxy-alkenals generated under physiological conditions may function as second messengers and modulate signal transduction and gene expression.

Acknowledgments Studies in the author‟s laboratory were supported by Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), PIP 2008-0157, Agencia Nacional de Promoción Científica y Tecnológica (ANPCyT) PICT-13399 and bilateral Grant HU/PA03BI/008.

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[109] Ji C, Amarnath V, Pietenpol JA, et al. 4-hydroxynonenal induces apoptosis via caspase-3 activation and cytochrome c release. Chem Res Toxicol 2001; 14: 10901096. [110] Friguet B, Szweda LI. Inhibition of the multicatalytic proteinase (proteasome) by 4hydroxy-2-nonenal cross-linked protein. FEBS Lett 1997; 405: 21-25. [111] Awasthi YC, Ansari GA, Awasthi S. Regulation of 4-hydroxynonenal mediated signaling by glutathione S-transferases. Methods Enzymol 2005; 401: 379-407. [112] Awasthi YC, Yang Y, Tiwari NK, et al. Regulation of 4-hydroxynonenal-mediated signaling by glutathione S-transferases. Free Radic Biol Med 2004; 37: 607-619. [113] Kutuk O, Basaga H. Apoptosis signalling by 4-hydroxynonenal: a role for JNK-cJun/AP-1 pathway. Redox Rep 2007; 12: 30-34. [114] Uchida K, Stadtman ER. Modification of histidine residues in proteins by reaction with 4-hydroxynonenal. Proc Natl Acad Sci USA 1992; 89: 4544-4548. [115] Nadkarni DV, Sayre LM. Structural definition of early lysine and histidine adduction chemistry of 4-hydroxynonenal. Chem Res Toxicol 1995; 8: 284-291. [116] Cohn JA, Tsai L, Friguet B, et al. Chemical characterization of a protein-4-hydroxy2-nonenal cross-link: immunochemical detection in mitochondria exposed to oxidative stress. Arch Biochem Biophys 1996; 328: 158-164. [117] Humphries KM, Szweda LI. Selective inactivation of alpha-ketoglutarate dehydrogenase and pyruvate dehydrogenase: reaction of lipoic acid with 4-hydroxy2-nonenal. Biochemistry 1998; 37: 15835-15841. [118] Humphries KM, Yoo Y, Szweda LI. Inhibition of NADH-linked mitochondrial respiration by 4-hydroxy-2-nonenal. Biochemistry 1998; 37: 552-557. [119] Musatov A, Carroll CA, Liu YC, et al. Identification of bovine heart cytochrome c oxidase subunits modified by the lipid peroxidation product 4-hydroxy-2-nonenal. Biochemistry 2002; 41: 8212-8220. [120] Echtay KS, Esteves TC, Pakay JL, et al. A signalling role for 4-hydroxy-2-nonenal in regulation of mitochondrial uncoupling. EMBO J 2003; 22: 4103-4110. [121] Brand MD, Affourtit C, Esteves TC, et al. Mitochondrial superoxide: production, biological effects, and activation of uncoupling proteins. Free Radic Biol Med 2004; 37: 755-767. [122] Bouillaud F, Couplan E, Pecqueur C, et al.Homologues of the uncoupling protein from brown adipose tissue (UCP1): UCP2, UCP3, BMCP1 and UCP4. Biochim Biophys Acta. 2001; 1504: 107-119. Review [123] Papa S, Skulachev VP, Reactive oxygen species, mitochondria, apoptosis and aging. Mol Cell Biochem 1997; 174: 305-319. [124] Murphy MP, Echtay KS, Blaikie FH, et al. Superoxide activates uncoupling proteins by generating carbon-centered radicals and initiating lipid peroxidation: studies using a mitochondria-targeted spin trap derived from alpha-phenyl-N-tertbutylnitrone. J Biol Chem 2003; 278: 48534-48545. [125] Gamaley IA, Klyubin IV. Roles of reactive oxygen species: signaling and regulation of cellular functions. Int Rev Cytol 1999; 188: 203-255. [126] Yang Y, Sharma R, Sharma A, et al. Lipid peroxidation and cell cycle signaling: 4hydroxynonenal, a key molecule in stress mediated signaling. Acta Biochim. Pol 2003; 50: 319-336.

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[127] Barrera G, Di Mauro C, Muraca R, et al. Induction of differentiation in human HL60 cells by 4-hydroxynonenal, a product of lipid peroxidation. Exp Cell Res 1991; 197: 148-152. [128] Barrera G, Pizzimenti S, Muraca R, et al. Effect of 4-hydroxynonenal on cell cycle progression and expression of differentiation-associated antigens in HL-60 cells. Free Radic Biol Med 1996; 20: 455-462. [129] Awasthi YC, Sharma R, Cheng JZ, et al. Role of 4-hydroxynonenal in stressmediated apoptosis signaling. Mol Aspects Med 2003; 24: 219-230. [130] Germain P, Staels B, Dacquet C, et al. Overview of nomenclature of nuclear receptors. Pharmacol Rev 2006; 58: 685-704. [131] Nettles KW, Greene GL. Ligand control of coregulator recruitment to nuclear receptors. Annu Rev Physiol 2005; 67: 309-333. [132] Bain DL, Heneghan AF, Connaghan-Jones KD, et al. Nuclear receptor structure: implications for function. Annu Rev Physiol 2007; 69: 201-220. [133] Nuclear Receptor Nomenclature Committee, A unified nomenclature system for the nuclear receptor superfamily. Cell 1999; 97: 161-163. [134] Issemann I, Green S. Activation of a member of the steroid hormone receptor superfamily by peroxisome proliferators. Nature 1990; 347: 645-650. [135] Dreyer C, Krey G, Keller H, et al. Control of the peroxisomal beta-oxidation pathway by a novel family of nuclear hormone receptors. Cell 1992; 68: 879-887. [136] Ricote M, Glass CK. PPARs and molecular mechanisms of transrepression. Biochim Biophys Acta 2007; 1771: 926-935. [137] Pizzimenti S, Laurora S, Briatore F, et al. Synergistic effect of 4-hydroxynonenal and PPAR ligands in controlling human leukemic cell growth and differentiation. Free Radic Biol Med 2002; 32: 233-245. [138] Cerbone A, Toaldo C, Laurora S, et al. 4-Hydroxynonenal and PPARgamma ligands affect proliferation, differentiation, and apoptosis in colon cancer cells. Free Radic Biol Med 2007; 42: 1661-1670. [139] Fazio VM, Barrera G, Martinotti S, et al. 4-Hydroxynonenal, a product of cellular lipid peroxidation, which modulates c-myc and globin gene expression in K562 erythroleukemic cells. Cancer Res 1992; 52: 4866-4871. [140] Barrera G, Pizzimenti S, Laurora S, et al. 4-hydroxynonenal and cell cycle. Biofactors 2005; 24: 151-157.

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Chapter 8

Lipid Nitration Lucía Bonilla Cal and Homero Rubbo* Departamento de Bioquímica, Facultad de Medicina, Universidad de la República. Montevideo, Uruguay

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Introduction Nitration of aliphatic compounds has been a subject of interest in organic chemistry since the early 19th century. Nonetheless, it was only recently linked to the pathobiological conditions associated to inflammation and nitro-oxidative stress. Pioneer work by Pryor et al. during the 1980s shed light over the radical pathways involved in the synthesis of nitro-fatty acids [1, 2]. During this period, the sole sources for nitration were considered to be toxins normally present in polluted air and/or cigarette smoke, particularly nitrogen dioxide (•NO2). A few years later and alongside with the breakthrough of nitric oxide (•NO) identification as the endothelial derived relaxing factor (EDRF) came the acknowledgment that endogenous synthesis of •NO-derived reactive species was involved in physiological processes involving modification of cellular targets. Hence, lipid nitration gained in the 1990s biological relevance as a potential pathway for nitro-oxidative modification of biological molecules [3]. In 2001 and 2002 several in vitro studies reported the ability of nitro-fatty acids to promote vasodilation, inhibit platelet activation and down-regulate the pro-inflammatory profile in human neutrophils [4-7]. In 2002, for the first time, nitrated derivatives of linoleic acid were detected in the plasma of healthy individuals [8]. Moreover, increased concentrations of these nitro-derivatives were found in hyperlipidemic donors, thus reinforcing their biological relevance and positioning nitro-fatty acids as potential footprints for nitro-oxidative damage in human disease. At this time, the starting hypothesis that nitrated lipids could have deleterious cellular effects rapidly switched to our actual conception that nitro-fatty acids represent redox signaling mediators that are able to modulate a variety of cell signaling pathways by interaction with specific cellular targets. *

Email: [email protected]

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This Chapter reviews the major chemical and biological aspects of nitrolipids, highlighting their roles as novel endogenously produced anti-inflammatory cell signaling mediators.

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2. Nitration of Biomolecules Nitration of biomolecules represents a key biologically-relevant redox signaling and injury event. There is a wide array of chemical reactions that might lead to nitration of targets including proteins, lipids and DNA, linked to overproduction of reactive nitrogen species (RNS) derived from the interaction of nitric oxide (•NO) with reactive oxygen species (ROS). Since production of RNS is intimately connected with oxidative stress, this condition is referred to by some authors as nitro-oxidative stress. Reactions involving synthesis of RNS are very relevant to pathophysiology since they lead to removal of the endothelial-derived relaxing factor - •NO - from the vasculature, critical for regulation of vascular tone. In the last decade, biological nitration has also been reported to modulate lipid activity in a physiologically relevant way. Nitration is traditionally classified into ionic, radical ion and free radical reactions [9], where present evidence suggests that radical (homolytic) nitration should occur in vivo. Nitration is classically defined as the reaction between an organic compound and a nitrating agent to introduce a nitro group (-NO2) on to a carbon atom (C-Nitration) or to produce nitrates (O-Nitration) or nitramines (N-Nitration). The nitro group most frequently substitutes a hydrogen atom [9]. A good candidate as nitrating agent in vivo is nitrogen dioxide (•NO2), which reacts with several biological targets such as thiols, ascorbate and urate (k~107 M-1s-1), as well as unsaturated fatty acids, tryptophan and tyrosine residues (k~105 M-1s-1). Sources for • NO2 formation in vivo include a) •NO autoxidation in cellular hydrophobic compartments, b) peroxynitrite, c) dinitrogen trioxide (N2O3), d) nitrite reduction catalyzed by peroxidases, and e) nitrite (NO2-) reduction at acidic pH. Protein tyrosine nitration (see also Chapter 6) represents a well characterized biomarker of cell injury and RNS reactions in vivo. This post-translational modification is a by-product of cell metabolism that could render proteins more susceptible to proteolytic degradation. Nitration of tyrosine involves an initial one-electron oxidation of a tyrosine residue to form tyrosyl radical -an aromatic radical cation- followed by a diffusion-controlled radical-radical termination reaction with •NO2 to yield 3-nitrotyrosine [10] (Figure 1). Different nitration pathways can contribute to in vivo nitration of tyrosine. Radicals arising from the homolysis of peroxynitrite were originally postulated as the major oxidizing species leading to the formation of the tyrosyl radical in hydrophilic compartments. More recently, alternative routes such as the nitrite/H2O2/heme-peroxidase and transition metal-dependent mechanisms [11], or the reaction of tyrosine with lipid peroxyl radicals in hydrophobic compartments [12] have been proposed.

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Figure 1. Free radical-mediated tyrosine nitration (adapted from [10]).

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We should differentiate nitration from nitrosation, defined as the donation of a nitrosonium ion (NO+) to a nucleophile. Sources for NO+ include N2O3 and •NO2, and the main products for nitrosation are nitrosoamines and S-nitrosothiols. Finally, nitrosylation is the formation of a nitrosyl adduct between •NO and metal protein constituents (e.g., iron). In vivo, nitrosylation reactions involve mainly ferric metalloproteins (e.g., guanylate cyclase, hemoglobin, cytochrome P450, nitric oxide synthase). Alternatively, •NO might nitrosylate thiolic components in proteins to form S-nitrosothiols by its reaction with a thiyl radical (S•), formed during thiol oxidation (see Chapter 4). Formation of S-nitrosothiols (RSNO) represents an alternative route for •NO to elicit biological signaling [13]. It is often referred to in the literature as “S-nitrosylation”, even though the nitrosation reaction is thought to predominate in vivo. S-nitrosothiols have the generic structure R-S-N=O, where a reactive thiol moiety in protein residues or low molecular weight antioxidants (e.g., glutathione) is covalently attached to a •NO group. In vivo, formation of S-nitrosothiols is expected to occur when •NO reaction with O2 is favored in the proximity of thiolic targets, with formation of the nitrosylating agent N2O3 (equation 1). 2 •NO + ½ O2 → N2O3 + RSH → RSNO + 2 NO2- + 2H+

eq. 1

RSNO + R`SH ↔ R`SNO + RSH

eq. 2

RSNO + R`SH → RSSR` + NO- + H+

eq. 3

The bond between the sulfur and nitrogen atoms in RSNO is not particularly susceptible to homolysis and S-nitrosothiols are fairly stable. Their role as cellular signaling molecules resides in their ability to act themselves as nitroso group donors, modifying thiols and other nucleophiles either by transnitrosation -transfer of the nitroso group from an S-nitrosothiol to a thiol- (equation 2) or by S-thiolation -generating a thiol disulfide and nitroxyl anion (NO-) (equation 3).

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3. Biological Aspects and Distribution of Unsaturated Fatty Acids

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Lipids represent a large and heterogeneous group of molecules that share the common feature of being insoluble in water. Lipids include fats, waxes, phospholipids, glycolipids and lipid-derived molecules such as glycerol, steroids, ketone bodies, hydrophobic vitamins, hormones and others. Knowledge in lipid biochemistry is necessary for the understanding of various biomedical areas of great interest, such as the pathological conditions associated with atherosclerosis, obesity and diabetes mellitus, or the role of essential fatty acids in nutrition, cell signaling and health. Fatty acids are the simplest constituents among lipids. Due to their hydrophobicity, fatty acids occur mainly as esters of more complex lipids such as oils, for the storage of fats in adipose tissue, or phospholipids, critical for membrane structure and function. They are aliphatic carboxylic acids containing a straight hydrocarbon chain with an even number of carbon atoms (typically from 12 to 24). Unsaturated fatty acids (UFAs) contain at least one double bond (e.g., 9-octadecenoic acid or oleic acid, 18:1 Δ9 in IUPAC nomenclature), while polyunsaturated fatty acids (PUFAs) contain two or more double bonds (e.g., linoleic acid, 18:2 Δ9,12 or linolenic acid, 18:3 Δ9,12,15 ), which are non-conjugated in mammals (i.e., they are separated by a methylene group). PUFAs serve a variety of biological roles including energy provision and storage, membrane structure, cell signaling and regulation of gene expression.

Figure 2. Four possible diastereomers of nitro-oleic acid.

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Fatty acid isomers differ in their structural formula. In UFAs, the geometric isomers are termed cis or trans, depending on the orientation of atoms or groups around the axes of double bonds, which do not allow rotation. Naturally-occurring UFAs in mammals have cis double bonds: acyl chains are on the same side of the bond such that the hydrocarbon chain is “bent” 120 degrees around the double bond. The various “bends” introduced into PUFAs that are constituents of biological membranes have important physicochemical consequences regarding the packaging and fluidity of the membrane. As discussed later in this Chapter, reaction of RNS with UFAs leads in some cases to a cis-trans isomerization of the acyl chain. Position isomers are structural isomers that differ only in the carbon atom to which a substituent group is bound in the acyl chain. As an example, all four theoretically possible isomers of the UFA nitro-oleic acid are illustrated in Figure 2. In the last 20 years, bioactive lipids have been shown to regulate a multitude of cellular responses including cell growth and death, and also inflammatory reactions. Representative examples are cyclooxygenase- and lipooxygenase-derived eicosanoids, peroxisome proliferation activating receptor (PPAR) activators, cannabinoids, sphingolipids and 4hydroxynonenal (4-HNE). As a general rule, bioactive lipid signaling pathways are basically composed of a regulated enzyme, the bioactive lipid itself or an active mediator derived from it, and specific downstream targets to reach the final effector. In particular, 4-HNE, a product of lipid peroxidation (see also Chapter 7), is a stable aldehyde and exhibits a great reactivity with thiol and amino groups, easily undergoing Michael additions with nucleophilic protein residues (i.e, cysteines and histidines). Covalent modification of cellular proteins is largely responsible for 4-HNE´s role as a signaling molecule. As we shall see later, electrophilicity is also critical for nitro-fatty acids to function as cell signaling molecules.

4. Nitration Chemistry in Hydrophobic Compartments Given the hydrophobic nature of lipids, fatty acid nitration is expected to occur in vivo mainly in hydrophobic compartments such as the lipid bilayer of cellular membranes or the lipophilic core of lipoproteins. Hence, hydrophobic compartments represent a reservoir for nitro-fatty acids which are likely to exert their biological effects upon cleavage from phospholipids by phospholipase-A2. Chemistry associated with the formation and decomposition of •NO-derived reactive species in hydrophobic compartments exhibits several significant differences when compared to the aqueous medium in the cytosol and other cellular domains. •NO freely diffuses into membranes and lipoproteins, due to its small molecular radius, neutral charge and hydrophobic character. •NO auto-oxidation (equation 4) is significantly accelerated in hydrophobic compartments relative to the surrounding aqueous environment [14-16]. In fact, while cell membranes represent narrowly 3% of the total cell volume, approximately 90% of • NO auto-oxidation takes place in this compartment [14]. This event is referred to as the “membrane lens effect”: the concentration of both reagents •NO and O2 inside the lipid bilayer is greatly enhanced due to a favoring partition, increasing the rate of reactions leading to formation of •NO2 despite the rather low reaction constant for •NO autooxidation (k = 1.5 - 3.0 x106 M-2 s-1). In addition, •NO2 hydrolysis within the lipid phase is minimal,

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favoring its reaction with targets available in the membrane. Reaction of •NO2 with a second molecule of •NO (eq. 4) leads to the formation of N2O3, a potent nitrating/nitrosating agent. In the aqueous phase, N2O3 would rapidly hydrolyze to NO2-. The membrane lens effect contributes to the role of •NO as an effective lipid antioxidant. During lipid peroxidation •NO reacts with unsaturated lipid reactive species such as alkyl (L•), epoxyallylic (L(O•)), alkoxyl (LO•) or peroxyl (LOO•) radicals generating radical-radical nitrogenated termination products [3]. 2 •NO + O2 → 2 •NO2 NO2 + •NO ↔ N2O3 N2O3 + H2O → 2 NO2- + 2 H+ ---4 •NO + O2 + 2 H2O → 4 NO2- + 4 H+

eq. 4

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On the other hand, peroxynitrite is a highly pH-dependent (pKa=6.8) oxidant and nitrating molecule that participates in direct one-and two-electron oxidation reactions. It easily crosses lipid membranes either as an anion through anion channels or in its protonated form by passive diffusion. Two different pathways have been suggested for peroxynitrite reactions with lipophilic targets: a) peroxynitrite-derived radicals are formed in aqueous media and then react with a substrate bound to the membrane surface, or b) peroxynitrous acid diffuses into the membrane and undergoes homolysis to generate •NO2 [17]. The latter is the most relevant peroxynitrite-mediated pathway for lipid oxidation and nitration in vivo [18].

Figure 3. Main radical routes for •NO2 formation in hydrophobic compartments.

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Figure 4. Main biological decomposition pathways for •NO. Reaction with superoxide, oxygen and metals leads to the formation of peroxynitrite, •NO2 and metal nitrosyl complexes, respectively.

Figure 5. Probable mechanisms for •NO-dependent inhibition of lipoperoxidation and formation of nitrogenated end products (adapted from [23]).

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A

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B

Figure 6. A) Hydrogen abstraction and B) addition mechanisms for •NO2-dependent fatty acid nitration under low and high oxygen tension (adapted from [20]).

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Although the detailed mechanisms for nitro-fatty acid synthesis in vivo still remain unknown, available knowledge suggests that •NO2 formation represents the limiting step. Figure 3 sketches the main pathways for •NO2 formation inside hydrophobic compartments. Under nitro-oxidative conditions, •NO2 derived from •NO auto-oxidation or proton-catalyzed homolysis of peroxynitrite might react directly with membrane UFAs leading to the formation of nitro-fatty acids, as discussed below. Alternatively, lipid peroxidation within the membrane can be terminated by •NO2 or •NO, leading to formation of nitrated derivatives, as shown in Figure 3. Overall, the predominance of nitration over oxidation will depend mainly on the local O2 concentration. A shift from lipid oxidation to lipid nitration should be expected during hypoxic conditions (e.g., ischemia-reoxygenation, inflammation), when tissue O2 levels are often suppressed while •NO levels are elevated, favoring lipoperoxidation termination reactions with •NO and/or direct homolytic reaction of UFAs with •NO2.

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5. Mechanisms of Fatty Acid nitration In vitro studies have described an ensemble of reactions leading to the introduction of nitrogenated groups into the hydrocarbon chain of fatty acids, where the combination of different mechanisms and nitrogenating agents promote the formation of a complex mixture of products [19, 20]. Extrapolating, however, in vitro observations to what actually occurs in biological systems poses a great challenge, because one should also consider the cellular sites of synthesis and biological distribution of reactive species, the half life of the species formed, and the presence of other biological targets which might also be susceptible to biological nitration. Despite all these uncertainties, nitration pathways involving free radical species have been fairly well characterized and actual knowledge on lipid nitration suggests that formation of •NO2 represents the limiting step. In this section, we will review what has been deciphered regarding the nitration mediated by •NO, peroxynitrite and •NO2. It should be stressed that besides radical-mediated nitration, the ionic addition of a nitronium ion (NO2+) via an electrophilic substitution at the double bond may also be relevant in nitration of fatty acids in hydrophobic compartments [21].

5.1. Nitric Oxide-dependent Mechanisms •

NO is synthesized during the oxidation of the amino acid L-arginine to citrulline, catalyzed by the enzyme nitric oxide synthase (NOS). •NO modulates vasodilation, inflammatory cell function and neurotransmission, through direct interaction with soluble guanylate cyclase (sGC). Additionally, •NO represents a key mediator of diverse physiological functions via sGC-independent pathways. •NO per se as well as •NO-derived RNS directly interact with critical biomolecules [22]. Figure 4 shows the main routes affecting •NO bioavailability. •NO can up- or downregulate biological oxidations thereby modulating free radical-mediated injury. This dual role of •NO in free radical-mediated oxidations is clearly demonstrated in the lipid peroxidation process [3, 23]. A typical doseresponse curve for •NO shows that for low •NO/O2•- ratios, increasing •NO levels leads to the generation of peroxynitrite and the initiation of lipid peroxidation (pro-oxidant effect) while

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NO terminates lipid peroxidation (anti-oxidant effect) when the •NO/O2•- ratio is equal or greater than 1 [3]. Lipid peroxidation is a complex process that generates hydroperoxides as primary oxidation products and cyclic peroxides as secondary oxidation products of UFAs. It is a free radical chain process having chain initiation, propagation and termination steps (Chapter 7). • NO is not reactive enough to initiate lipid oxidation, however it is highly reactive with lipidderived radicals generated during the propagation stage, such as alkoxyl (LO•) or peroxyl (LOO•) radicals (k=1-3x109 M-1s-1) [24], yielding a variety of nitrogenated products (Figure 5). In this case, •NO acts as an effective anti-oxidant, terminating lipid radical-mediated chain propagation reactions and thus protecting membrane lipids and lipoproteins from oxidative modification and redirecting the cytotoxic reactions mediated by superoxide anion (O2•-) and peroxynitrite towards other oxidative pathways. As mentioned earlier, the membrane “lens effect” which arises from the high diffusivity of •NO into biological membranes makes this fatty acid nitration pathway highly probable under nitro-oxidative stress. Additionally, based on relative rate constants, •NO is a more potent inhibitor of lipid peroxidation propagation reactions than α-tocopherol (vitamin E) (k ~ 5x105 M-1s-1), thus sparing this membrane antioxidant from oxidation [25].

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•

Figure 7. Main routes for peroxynitrite-mediated UFA nitration: peroxynitrite decomposition through H+- and CO2- catalyzed homolysis (I and II) leads to the formation of •NO2, a potent oxidizing and nitrating molecule and •OH a very potent oxidizing molecule.

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In vitro studies have shown that addition of •NO to peroxidizing lipid mixtures leads to inhibition of oxygen consumption, which resumes at original rates once all •NO has been consumed. Kinetic analysis indicates that at least 2 molecules of NO• are consumed per LOO• generated. An organic peroxynitrite (LOONO) is generated as the initial product of the reaction of LOO• and NO•. LOONO is a non-stable intermediary and may follow different decomposition routes via the caged radical pair [LO••NO2] which account for the stoichiometry of the inhibition process (Figure 5). Homolysis of the O-O bond in LOONO yields LO• and •NO2, and a second •NO molecule is consumed via reaction with LO• generating an alkylnitrite (LONO) as the final product [21, 23]. Alternative end products can be generated, as shown in Figure 5.

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5.2. Nitrogen Dioxide-dependent Mechanisms Unlike •NO, •NO2 is a fairly potent oxidizing and nitrating agent and its synthesis is likely to represent the limiting step in biological fatty acid nitration. In hydrophobic compartments the most probable source for •NO2 is the autooxidation of •NO, which concentrates up to 20fold in the membrane bilayer, where it is quickly consumed by O2 in a third order kinetics reaction (k=2x106 M-1s-1) [14] (Figure 3). •NO2 can also be generated from the decomposition of peroxynitrite, peroxidase-catalyzed oxidation of NO2- to •NO2 or reduction of NO2- in acidic tissue environments (e.g., gastric compartment). Altogether, these pathways account for the basal concentration of •NO2 found in tissues. The currently accepted mechanisms for •NO2-mediated oxidation and nitration of UFAs involve three routes: hydrogen atom abstraction and addition reactions, described in the 1980s by Pryor et al [1], and electrophilic substitution by a nitronium ion (NO2+) [26]. Reaction of • NO2 with UFAs leads to the generation of isomerized, oxidized and/or nitro-allylic, nitroalkene, dinitro, or nitro-hydroxy lipid derivatives (Figure 6). The profile of synthesized species depends mainly on •NO2 concentration. Also, since •NO2 can initiate lipid oxidation reactions, O2 levels impact the nitration versus oxidation yield, after reaction of •NO2 with UFAs [1, 27, 28]. There is a competition between both pathways for •NO2-dependent nitration (Figure 6). When •NO2 concentration is low (under 100 ppm) the hydrogen abstraction pathway is likely to predominate (Figure 6A). Generally speaking, H-atom abstraction by •NO2 is extremely slow, but it becomes very relevant in biological systems with activated hydrogen atoms (i.e., weak atom-hydrogen bond). In PUFAs the bis-allylic methylene centers represent the most probable reactive sites with •NO2 given the lability of their hydrogen atoms. The allylic or bis-allylic hydrogen abstraction in UFAs and PUFAs, respectively, generates a carboncentered lipid radical and nitrous acid (HONO), which rapidly decomposes turning the hydrogen abstraction reaction irreversible. In anaerobic conditions or when O2 tension is low, a second molecule of •NO2 reacts with the carbon-centered radical generating a nitro-alkane (i.e., a nitro-allylic derivative, where the -NO2 moiety is bound to a saturated carbon center). However, when the O2 tension is higher, the carbon-centered radical reacts with O2 to generate a nitro-peroxyl radical, an unstable and very reactive species that can react with a neighboring UFA, thus initiating the propagation stage of lipoperoxidation, or with a second • NO2 radical generating a nitro-nitrate alkane.

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Unlike the hydrogen abstraction pathway, the addition of •NO2 to an unsaturated carbon center is a reversible reaction. Nonetheless, when the concentration of •NO2 is fairly high, the addition mechanism is more likely to account for •NO2 -dependent fatty acid nitration/oxidation (Figure 6B). Moreover, in the presence of species that exhibit high reactivity towards the β-nitroalkyl radical, the first intermediary of the reaction, the equilibrium is shifted toward its production, disfavoring the hydrogen abstraction mechanism. In this conditions, a homolytic attack of •NO2 on the double bond in UFAs yields the βnitroalkyl radical. In an aqueous environment, where the solubility of •NO and its derived reactive species is rather low, the reaction will most probably revert, causing a cis-trans isomerization of the UFA and the loss of HONO. The generation of trans-fatty acids along with nitro-fatty acids is unique to •NO2 -dependent mechanisms. Otherwise, when the O2 levels are low, the β-nitroalkyl radical reacts with a second •NO2 radical generating a dinitroalkane acid and/or a nitro-nitrite alkane. Both these species are unstable and decompose to a nitroalkene, with the concurrent loss of HONO. Alternatively, the loss of HONO leads to the generation of trans-nitroalkenes and/or trans-nitroalkanes. However, the nitro group preferentially binds the double bond in nitro-alkene derivatives in the cis configuration. Trans-nitro isomers are expected to be a minority among the products generated. Yet another possible product is a nitro-hydroxy alkane (a nitro-alcohol), generated from the hydrolysis of a dinitroalkane/nitro-nitrite alkane. When the β-nitroalkyl radical is generated under aerobic conditions, its reaction with O2 yields a nitro-peroxyl radical. By analogy to the hydrogen abstraction mechanism under aerobic conditions, the nitro-peroxyl radical will initiate the propagation stage of lipid peroxidation, or react with •NO2 to form a nitro-nitrate alkane.

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5.3. Peroxynitrite-dependent Mechanisms Peroxynitrite is formed in vivo as a result of the diffusion controlled reaction between NO and O2•- radicals (k=109-10 M-1s-1). Given the high rate constant for the reaction, •NO sometimes out competes SOD for O2•- resulting in steady-state concentrations of peroxynitrite in the nanomolar concentration range [29]. Peroxynitrite anion (ONOO-) and its conjugate acid (ONOOH, pKa = 6.8 at 37ºC) are strong oxidizing species that react with a wide variety of biological targets, including protein tyrosine residues, thiols and UFAs. Peroxynitrite per se does not react with the C-H bonds in UFAs; rather, the main route for peroxynitritedependent fatty acid nitration is the generation of •NO2 following ONOOH homolysis. Reaction of peroxynitrite with UFAs yields a variety of oxidation and nitration products. For linoleic acid (18:3), major products include nitroso-peroxo-linolenate, hydroxyl-nitrosoperoxo-linolenate and hydro-peroxo-nitroso-peroxo-linolenate [3]. The biological chemistry of peroxynitrite is fairly complicated. In particular, pH and CO2 redirect peroxynitrite reactivity, enhancing lipid nitration and limiting oxidation reactions [30]. Figure 7 provides an overview of peroxynitrite reactivity with UFAs. Since both peroxynitrite and •NO2 readily diffuse through the membrane bilayers, reactions leading to •NO2 generation may take place in the aqueous environment in proximity to the membrane or inside the lipid bilayer (Figure 3). In the absence of CO2, the proton-catalyzed homolysis of peroxynitrite at the O-O bond yields •NO2 and •OH. The latter is an extremely potent oxidant capable of initiating lipid peroxidation, whilst •NO2 is capable of both oxidation and nitration of UFAs, as reviewed above. In the presence of CO2, peroxynitrite rapidly decomposes to •NO2 and CO3•-. CO3••

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radical is a strong oxidant but its oxidative damage is constrained to surface-exposed lipids, because it is unable to penetrate the biological membranes due to its anionic nature.

6. Biochemical Characterization of Nitro-fatty Acids The unique physicochemical properties of nitro-fatty acids compared to unmodified fatty acids are due to the presence of the nitrogen-containing group bonded to the hydrocarbon chain (i.e., a -NO2 group in nitro-alkenes or nitro-alkanes). Nitro-fatty acids are slightly more polar than their unmodified analogs, nonetheless maintaining a strong hydrophobic behavior due to the hydrocarbon chain. The most relevant physicochemical characteristic of nitro-fatty acids is their strong electrophilic nature. The high electron-withdrawing capacity of the nitro group renders the adjacent β-carbon in the hydrocarbon chain very electrophilic in addition to increasing the acidity of the adjacent hydrogens. The bis-allylic hydrogens of the methylene group found in nitro-PUFAs (i.e., located between two unsaturated carbon centers) are particularly reactive/labile, providing explanation for the lower stability of nitro-PUFAs compared to nitro-UFAs. Altogether, the biochemical characteristics conferred secondary to the addition of a nitrogen-containing group into an UFA account for the growing number of anti-inflammatory properties attributed to nitrated lipids.

7. Biological Stability of Nitro-fatty Acids: NO-donor Properties and Post-translational Modification of Proteins

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•

Nitro-fatty acids stored by esterification to the complex lipid constituents of membranes and lipoproteins remain stable and non-reactive. Whether they are generated in situ or esterified following nitration, A2-type phospholipases are capable of releasing nitro-fatty acids during inflammatory conditions or in response to other stimuli. Mobilization of nitrolipids, which allows them to reach specific cellular targets and exert signaling actions, involves the formation of reversible covalent adducts with cytoplasmatic and/or plasma proteins and low molecular weight thiols, as discussed below. The biological stability of nitroalkenes is greatly altered in aqueous environments, where they spontaneously decay releasing •NO. Direct evidence for nitroalkene •NO-donor capacity has been demonstrated by electron paramagnetic resonance (EPR) spectroscopy using a selective spin trap for •NO, as well as by UV-visible spetroscopic analysis following formation of the decomposition product at 320 nm. Additionally, indirect evidence shows that nitroalkenes activate sGC and exert vasorelaxation, presumably through •NO release [31-33]. A hydrophobically-controlled modified Nef reaction has been proposed as the chemical basis leading to the release of •NO from nitroalkenes; it occurs via formation of a nitroso intermediate that rapidly degrades to release •NO. Alternatively, the nitroalkene might rearrange to a nitrite ester, followed by N-C bond homolysis to generate •NO [4, 32].

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A

B

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Figure 8. Nitro-fatty acid reaction with nucleophiles. A) Mechanism of Michael addition reaction; the nucleophile reacts the electrophilic β-position to form a covalent adduct. B)ofSpectral Fig. 8.atNitro-fatty acid reaction with nucleophiles. A) Mechanism Michaelanalysis addition of nitrolinoleic acid reaction; (100 μM)the reaction with glutathione (GSH) (1 mM); extracted [35]. a covalent nucleophile reacts at the electrophilic β-positionfrom to form

B) Spectral analysis of nitro-linoleic acid (100 μM) reaction with glutathione

• 35 of nitrolipids as NO reservoirs in vivo, the most Despite(GSH) the (1well-ascertained mM); extracted fromrole . relevant cell signaling activities of nitroalkenes are linked to their receptor-mediated reactions and strong electrophilic nature. The highly electronegative nitro functional group facilitates reaction of the carbon β adjacent to the nitro group with nucleophilic cellular targets (i.e., cysteine, histidine or lysine residues in proteins, glutathione, other thiol groups) via a Michael addition reaction, often referred to in the bibliography as a nitroalkylation reaction when the electrophile is a nitroalkene (Figure 8A). Importantly, nitroalkylation-induced adducts are easily reversed in the physiological range of concentration of glutathione, thus representing a distinct mode of signal transduction from more traditional non-covalent ligand-receptor interactions that elicit responses by inducing conformational changes. The electrophilic property of nitroalkenes was first suggested by the biological detection of nitro-hydroxy fatty acid derivatives [4]. Nitroalkenes outside the hydrophobic environment of membranes or lipoproteins interconvert to a vicinal nitrohydroxy fatty acid following reaction with hydroxide anions present in aqueous solution at physiological pH. Naturally, this is a pH-regulated reaction greatly favored by basic conditions. Since nitrohydroxy fatty acids remain in equilibrium with their precursors while being much more polar and stable, the formation of these products might be relevant for nitroalkene cellular trafficking. Even so, most of the nitroalkenes released into the cytosol are expected to be

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initially bound as glutathione adducts, given the high intracellular concentration (up to 5 mM) of this major cellular antioxidant (Vol. II, Chapter 1). In addition, adducts between glutathione and nitro -oleic and -linoleic acids, the most abundant UFAs, are present in healthy human red blood cells [34]. Besides its role in the intracellular trafficking of nitro-fatty acids, nitroalkylation is involved in the regulation of target proteins. Evidence of protein nitroalkylation in vivo has been obtained from matrix-assisted laser desorption and ionization time of flight mass spectrometry (MALDI-TOF MS) [34]. Remarkably, nitroalkenes display the largest secondorder rate constants for the bimolecular reaction with cysteine and glutathione (k=183 and 355 M-1s-1 for nitro-oleic acid and nitro-linoleic acid, respectively, at pH 7.4 and 37 ºC), when compared to other biologically relevant non-nitrated lipid-derived electrophiles (e.g., 4hydroxynonenal, isoprostanes, etc). Reactions of nitroalkenes with nucleophilic targets can be easily followed by UV spectrophotometry (Figure 8B). Nitroalkenes have distinctive spectral properties compared to their unmodified analogs. A maximum of absorbance is registered at around 270 nm, characteristic of the conjugated system that is generated in nitroalkenes upon addition of a nitro group to a vinyl group, while unmodified fatty acids do not display absorbance outside the far UV region. During the course of a nitroalkylation reaction, the NO2-vinyl group in nitroalkenes is reduced to a NO2-allylic group (i.e., -NO2 is bound to a saturated C-C bond); as a consequence, nitroalkylation of nucleophilic targets can be easily characterized by UV spectrophotometric analysis following the decrease in absorbance at 270 nm during the course of product formation. As a representative example, Figure 8B shows glutathione nitroalkylation mediated by nitro-linoleic acid [35].

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8. Metabolic Fate of Nitro-fatty Acids Since nitro-fatty acids are synthesized in vivo and are very similar to unmodified fatty acids in their overall structure, it is reasonable to suppose that they will be metabolized, transformed and degraded in a similar fashion. At present, this is one of the least-studied aspects regarding nitrated lipid biochemistry. Recent evidence shows that nitro-fatty acids are metabolized via saturation of the double bond to generate derived nitroalkanes and hepatic βoxidation reactions that terminate at the site of acyl-chain nitration [36]. This reinforces the hypothesis that nitro-fatty acids share, to some extent, the metabolic fate characteristic of unmodified fatty acids; research in the next few years will probably shed light on these aspects.

9. Cell Signaling and Anti-inflammatory Properties of Nitrated Lipids Lipid nitration represents a novel mechanism for nitrated biomolecules to influence adaptive and anti-inflammatory cell responses, and may prove to be relevant in the progression of inflammation-related diseases. Inflammation (Vol. II, Chapter 21) is a

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complex biological response of the body to penetration of an infectious agent, entrance of antigen, or cell damage; it represents an attempt by the organism to remove the injurious stimuli as well as initiate the healing process for the tissue. This critical biological process represents the most frequent sign of disease, and there is a key involvement of inflammatory reactions in the pathogenesis of various diseases (e.g., atherosclerosis, myocardial infarction, chronic heart failure, Alzheimer´s disease, diabetes mellitus, cancer) [37]. Inflammatory cells, including blood neutrophils, monocytes/macrophages, platelets or endothelial cells, respond to pro-inflammatory signals by switching to an inflammatory phenotype, characterized by the expression of molecules for leukocyte and endothelium adhesion (i.e., selectins) that cause adhesion of blood cells to endothelium cells, penetration of blood cells across vascular walls and migration into the inflammation focus. Secondary to the phenotype switch, inflammatory mediators are released. Among these, cytokines play key roles in the initiation and regulation of the inflammatory response, controlling most of the regulatory and effector substances and inflammatory reactions mediated by them. Some of the proinflammatory enzymes expressed following activation of inflammatory genes are phospholipase-A2 (releases fatty acids from membrane phospholipids), cyclooxygenase (catalyzes the limiting step in the synthesis of prostanoids), inducible nitric oxide synthase, and NADPH oxidases. Acute inflammation is the primary response to infection; it is characterized by an increased vascular permeability, entry of activated leukocytes into the tissues, and activation of platelets and endothelial cells. An eventual resolution (suppression) of the inflammatory state is critical to shelter healthy cells from „collateral‟ damage. A persistent and chronic inflammatory state leads to a progressive shift in the types of cells present at the site of inflammation and to the development of a host of inflammatory-related diseases [37]. Nitrolipids have emerged over the last few years as very promising vascular protective compounds that aid in the resolution of inflammation. Despite the demonstrated role of nitrofatty acids as stimulators of smooth muscle relaxation [4, 5, 33], their anti-inflammatory effects described so far are mostly independent of •NO release. In general terms, they exert anti-inflammatory activity through rapid electophilic and receptor-mediated reactions leading to nitroalkylation of proteins playing critical roles in the regulation of the inflammatory response. Next, we will review the nitro-fatty acid-mediated inhibition of inflammatory cell function and the modulatory effects on transcription factors, which span most of the biological signaling roles ascribed to nitro-fatty acids. However, it should be noted that research in this area is rapidly growing and other nitroalkylation-dependent anti-inflammatory roles for nitro-fatty acids have been described, such as cardioprotection during ischemiareperfusion injury or ischemic-preconditioning, as well as nociceptive nerve activation during nitroxidative stress [34, 38-40]. Importantly, nitro-fatty acids have been detected in plasma from healthy individuals as well from patients with inflammatory conditions [8, 41]. Currently, there is no accord concerning actual nitro-fatty acid concentrations found in vivo.

9.1. Inhibition of Inflammatory Cell Function Nitro-fatty acids are potent inhibitors of inflammatory cell function. Nitro-linoleic acid alone has been reported to inhibit the pro-inflammatory activity of monocytes/ macrophages, neutrophils, platelets and endothelial cells [6, 7, 42, 43]. Nitro fatty acid-dependent

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attenuation of the inflammatory phenotype observed in these cell types follows •NOindependent mechanisms. Although the anti-inflammatory effects of nitro-fatty acids have mostly been characterized in vitro, the identification of an enhanced generation of endogenous nitro-fatty acid derivatives during macrophage activation [43], together with the identification of endogenously-generated nitro-fatty acids in human plasma and urine [8], reinforce the conception that nitrated lipids are generated in vivo as an adaptive response to nitro-oxidative inflammatory conditions. The initial observation that nitro-linoleic acid inhibits neutrophil activity sparked a further characterization of nitrated lipids as anti-inflammatory signaling molecules. Activation of neutrophils, the most abundant phagocytic cells found in the bloodstream, is a central feature of inflammatory disease. In response to inflammatory stimuli, neutrophils are recruited from the bloodstream and adhere to the endothelial lining of capillaries to reach the tissues. Nitro-linoleic acid inhibits neutrophil activation via down-regulation of superoxide generation, calcium mobilization, degranulation and integrin CD11b expression (involved in adhesion and transendothelial migration to inflammatory sites), all of which are requisites for activated neutrophils to mediate cytotoxicity during the inflammatory response [7]. The inhibition occurs upstream of NADPH oxidase (responsible for the generation of superoxide) and independently from •NO generation. Rather, it is associated to an increase in cAMP levels. Likewise, nitro-linoleic acid exibits an anti-thrombotic effect through inhibition of platelet activation in a cAMP-dependent mechanism [6]. Platelets are key players during the first stage of hemostasis, undergoing a series of reactions such as adhesion to the endothelium, aggregation, release of granule content and morphological changes that lead to the formation of the platelet plug. Platelet-derived mediators during the adhesion process have strong pro-inflammatory effects altering chemotactic, adhesive, and proteolytic properties of endothelial cells, which in turn support the interactions among platelets, leukocytes and endothelial cells typically observed in a site of inflammation. The level of platelet activation is tightly regulated through the action of both pro- (e.g., thromboxanes) and anti- (e.g., prostacyclin) aggregatory eicosanoids as well as through •NO-dependent mechanisms. However, under certain pathological conditions, platelet-induced chronic inflammatory processes at the vascular wall result in development of atherosclerotic lesions and atherothrombosis, which is often the primary cause of death in atherosclerotic vascular disease. Nitro-linoleic acid inhibits the thrombin-stimulated aggregation of platelets and the expression of the adhesion molecule P-selectin, which plays a key role for platelet adhesion to the endothelial lining. This antithrombotic effect is governed by a cAMP-dependent modulation of VASP (vasodilator stimulated phosphoprotein) phosphorylation on Ser-159 and by attenuation of thrombin-induced calcium mobilization [6]. Nitro-fatty acids also exert strong anti-inflammatory effects in macrophages, decreasing the expression of the inducible enzyme nitric oxide synthase-2 (NOS2) and down-regulating the lipopolysaccharide-induced secretion of proinflammatory cytokines (IL-6, TNFα) independently of •NO formation, PPARγ activation or induction of heme oxygenase 1 (HO-1) [42]. Macrophages are phagocytic cells derived from monocytes that promote the generation of an inflammatory phenotype by regulation of the differentiation and function of vascular and nonvascular cells through the expression and secretion of cytokines, reactive species, and other chemical mediators. Of note, the activation of macrophages following inflammatory stimulus leads to the endogenous generation of nitro-fatty acid-derivatives, capable of

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shutting down the inflammatory response mediated by these cells [43] and aiding the resolution of the inflammatory state. Also, in vitro studies show that nitro-fatty acids inhibit the adhesion of monocytes to the endothelial lining [42], a key initial step for both acute and chronic inflammation in the vascular system. As a consequence of endothelial cell activation, monocytes adhere to the endothelium first by rolling and then by firm adhesion to migrate ultimately into the intima, where they differentiate into macrophages. Nitro-fatty acids modulate the transendothelial migration of monocytes by down-regulation of adhesion molecules present in the activated endothelial cells (e.g., intracellular adhesion molecule-1, VCAM-1, E-selectin and P-selectin). In this regard, both nitro- oleic and -linoleic acids inhibit VCAM-1 expression when present in the nanomolar range [42].

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9.2. Modulation of Transcription Factors When the strong electrophilic reactivity of nitro-fatty acids was described, posttranslational modification of proteins emerged as a plausible mechanism for the observed modulation of inflammatory cells. Indeed, nitro-fatty acids covalently bind several critical protein residues altering their biological function [34, 44]. Particularly, nitroalkylation of nuclear transcription factors seems to be the main mechanism for nitro-fatty acids to modulate inflammatory cells responses, as reviewed above. In this regard, the transcription factor PPARγ contains a critical cysteine residue in the ligand-binding domain subject to nitroalkylation by nitro- oleic and -linoleic acids [41]. In the vasculature, PPARγ is expressed in monocytes/macrophages, smooth muscle cells and endothelium and plays key roles in the regulation of energy balance and adipogenesis. Agonists for PPARγ, mainly fatty acids, upregulate target gene transcription, thereby modulating adipocyte differentiation, lipid trafficking, glucose metabolism and inflammation. Nitro -oleic and -linoleic acids are potent agonists for PPARγ, resulting in macrophage CD36 expression, adipocyte differentiation, and glucose uptake at a potency comparable to thiazolidineadiones (TZDs) [32, 41]. Synthetic PPARγ ligands are widely used in the treatment of various metabolic disorders (e.g., type II diabetes). The dose-dependent activation of PPARγ mediated by nitro-fatty acids, particularly by nitro-oleic acid, showed a much greater binding affinity and overall potency compared to previously reported natural agonists like free fatty acids, eicosanoid derivatives and platelet-activating factor, whose reported concentration requirements for PPARγ activation actually exceed their physiological concentration ranges. PPARγ is activated by nitro-linoleic acid at concentrations as low as 100 nM. The transcription factor NF-κB (Vol. II, Chapter 12) is also subject to negative regulation by several naturally-occurring electrophiles through alkylation of highly conserved cysteine residues in the DNA-binding domains p50 and p65. NF-κB plays a crucial role in the induction of inflammatory cytokines and enzymes, chemokines, cell adhesion molecules, acute phase proteins, and growth factors. Nitro -oleic and -linoleic acids nitroalkylate subunit p65 of NF-κB in vitro as well as in intact macrophages, inhibiting its interaction with DNA and thus the NF-κB-mediated proinflammatory responses [42]. Also, fatty acid nitration products inhibit endotoxin-mediated STAT proinflammatory signaling through the induction of mitogen-activated protein kinase-1 (MAPK-1), a MAPK phosphatase known to contribute to anti-inflammatory signaling through alteration in the

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translation of mRNA to proteins [45]. Induction of MAPK-1 is independent of PPARγ activation, •NO release and also nitroalkylation, revealing that there are novel molecular mechanisms that contribute to the anti-inflammatory signaling actions of nitroalkenes. Furthermore, nitro-linoleic acid-dependent HO-1 induction [42, 46, 47] has been recently linked to the MAPK/ERK signaling pathway [48]. HO-1, which is also up-regulated by other electrophilic derivatives such as arachidonate-derived 15d-PGJ2 [49], mediates central events in the cytoprotective responses triggered during inflammatory conditions (Vol. II, Chapter 11). The degradation of pro-oxidant heme catalyzed by HO-1 releases carbon monoxide, iron and biliverdin and has a protective effect against vascular disease as well as acute inflammatory disorders such as sepsis or organ transplant rejection [49]. Regulation of the expression of HO-1 is subject to complex gene regulation via antioxidant response elements (Vol. II, Chapter 13), and binding sites for PPAR α or γ and NF-κB, while alternative molecular targets for nitro-linoleic acid modulation have been proposed [47].

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10. In Vivo Detection of Nitrosyl-fatty Acids: Current Knowledge, Challenges and Potential Pitfalls Demonstration that nitro lipids are present in biological systems is important to support and sustain the growing amount of evidence for the beneficial role of nitro-fatty acids as potent anti-inflammatory signaling mediators. In 2002, stable nitrated derivatives of nitrolinoleic acid were identified in human blood plasma from normolipidemic and hyperlipidemic donors [8]. Shortly afterwards, cholesteryl nitrolinoleate was detected in human blood plasma and lipoproteins of normolipidemic patients [50]. Nitro linoleic acid has also been detected in healthy human red blood cells both free and esterified to other molecules [34, 51], while nitro-oleic acid was identified in plasma, red cells, and urine from healthy humans [41]. More recently, endogenous formation of nitro-linoleic acid was found in heart-isolated mitochondria exposed to simulated ischemic preconditioning, and exhibited cardioprotection under this condition [39, 52]. Also, endogenous cholesteryl nitro-linoleate levels increase up to 20-fold following macrophage activation leading to the suppression of macrophage inflammatory responses [43]. Despite the mounting evidence for endogenous formation of nitro-fatty acids, several key aspects remain unclear [20]. Accordingly, reported values for nitro-fatty acid concentration in biological samples has switched from the micromolar [41] to the picomolar range [53] in the last few years. A lack of appropriate standards is partly responsible for the disagreement as to the net tissue concentration of nitrated lipid derivatives. Mostly, the identification and quantification of nitro-fatty acids in biological samples has been performed using nitroalkenes as standards; however there are other structural possibilities as well as nitro-allylic lipid derivatives probably present in biological samples. Additionally, the internal standards for nitro-fatty acids used so far constitute a mixture of positional isomers. On one hand, this leads to underestimation of the total concentration of the nitro-fatty acid in the sample, and on the other hand, it does not allow detection of the isomer preferentially formed in vivo. In this regard, in 2006 Woodcock et al. reported the regio- and stereospecific synthesis of all four

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possible diastereomers of nitro oleic acid [54] (shown in Figure 2). However appropriate standards for nitro-arachidonic acid and other biologically relevant nitro-fatty acids are still unavailable. The hydrophobic nature of nitro lipids also poses an additional methodological complexity that should be considered during the experimental design. Given the biochemical characteristics described previously, various cellular pools of nitrolipids are expected to be found in vivo: free, esterified to complex lipids in hydrophobic compartments and proteinadducted nitrolipids. At present, the release and stability of nitro-fatty acids in membranes or lipoproteins remain unknown, probably affecting quantification studies. The handling of the biological samples to be analyzed should include protocols for de-esterification from proteins and complex lipids. Finally, acidic nitration is often favored during sample processing and can mediate artifactual generation of nitro-fatty acids, leading to overestimated concentrations of nitrofatty acids; acidic conditions should be avoided at all times during handling and analysis of the sample, or at least this aspect should be considered in order to have adequate controls for artifactual nitration.

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Conclusions Lipid nitration represents a novel mechanism for •NO and •NO-derived free radical species to transduce metabolic and inflammatory information. Nitro-fatty acids are present in the vasculature [51] and their formation is induced under inflammatory conditions [43], supporting a role of lipid nitration in adaptive redox-sensitive signaling. They are present in the nanomolar to micromolar range of concentrations, sufficient to exert biological actions that aid in the resolution of inflammation, including inhibition of inflammatory cell activation, pro-inflammatory cytokine and prostaglandin [55] secretion and vascular smooth muscle proliferation [20]. Overall, these data reveal that inflammatory-derived fatty acid nitration products mediate homeostatic signaling reactions in vivo. Further work is necessary to determine whether nitro-fatty acid supplementation exerts anti-inflammatory and cytoprotective actions in inflammatory diseases.

Acknowledgments This work was supported by Welcome Trust (UK) and ICGEB (Italy) to Homero Rubbo and ANII (Agencia Nacional de Investigación e Innovación, Uruguay) to Lucía Bonilla Cal.

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[17] Rudakov ES, Lobachev VL, Geletii YV. Does peroxynitrite partition between aqueous and gas phases? Implication for lipid peroxidation. Chem Res Toxicol. 2001 Sep;14(9):1232-8. [18] Rubbo H, Trostchansky A, O'Donnell VB. Peroxynitrite-mediated lipid oxidation and nitration: mechanisms and consequences. Arch Biochem Biophys. 2009 Apr 15;484(2):167-72. [19] Freeman BA, Baker PR, Schopfer FJ, Woodcock SR, Napolitano A, d'Ischia M. Nitro-fatty acid formation and signaling. J Biol Chem. 2008 Jun 6;283(23):15515-9. [20] Trostchansky A, Rubbo H. Nitrated fatty acids: mechanisms of formation, chemical characterization, and biological properties. Free Radic Biol Med. 2008 Jun 1;44(11):1887-96. [21] O'Donnell VB, Eiserich JP, Bloodsworth A, Chumley PH, Kirk M, Barnes S, et al. Nitration of unsaturated fatty acids by nitric oxide-derived reactive species. Methods Enzymol. 1999;301:454-70. [22] Fukuto JM. Nitric Oxide: Biology and Pathobiology, Chapter 2: The Chemical Properties of Nitric Oxide and Related Nitrogens Oxides; Ignarro, L.J. 2000. [23] O'Donnell VB, Chumley PH, Hogg N, Bloodsworth A, Darley-Usmar VM, Freeman BA. Nitric oxide inhibition of lipid peroxidation: kinetics of reaction with lipid peroxyl radicals and comparison with alpha-tocopherol. Biochemistry. 1997 Dec 9;36(49):15216-23. [24] Padmaja S, Huie RE. The reaction of nitric oxide with organic peroxyl radicals. Biochem Biophys Res Commun. 1993 Sep 15;195(2):539-44. [25] Rubbo H, Radi R, Anselmi D, Kirk M, Barnes S, Butler J, et al. Nitric oxide reaction with lipid peroxyl radicals spares alpha-tocopherol during lipid peroxidation. Greater oxidant protection from the pair nitric oxide/alpha-tocopherol than alphatocopherol/ascorbate. J Biol Chem. 2000 Apr 14;275(15):10812-8. [26] O'Donnell VB, Eiserich JP, Chumley PH, Jablonsky MJ, Krishna NR, Kirk M, et al. Nitration of unsaturated fatty acids by nitric oxide-derived reactive nitrogen species peroxynitrite, nitrous acid, nitrogen dioxide, and nitronium ion. Chem Res Toxicol. 1999 Jan;12(1):83-92. [27] Gallon AA, Pryor WA. The reaction of low levels of nitrogen dioxide with methyl linoleate in the presence and absence of oxygen. Lipids. 1994 Mar;29(3):171-6. [28] Napolitano A, Panzella L, Savarese M, Sacchi R, Giudicianni I, Paolillo L, et al. Acid-induced structural modifications of unsaturated Fatty acids and phenolic olive oil constituents by nitrite ions: a chemical assessment. Chem Res Toxicol. 2004 Oct;17(10):1329-37. [29] Szabo C, Ischiropoulos H, Radi R. Peroxynitrite: biochemistry, pathophysiology and development of therapeutics. Nat Rev Drug Discov. 2007 Aug;6(8):662-80. [30] Radi R. Nitric Oxide: Biology and Pathobiology, Chapter 4: The Biological Chemistry of Peroxynitrite. Ignarro, L.J. 2000. [31] Lima ES, Bonini MG, Augusto O, Barbeiro HV, Souza HP, Abdalla DS. Nitrated lipids decompose to nitric oxide and lipid radicals and cause vasorelaxation. Free Radic Biol Med. 2005 Aug 15;39(4):532-9. [32] Schopfer FJ, Baker PR, Giles G, Chumley P, Batthyany C, Crawford J, et al. Fatty acid transduction of nitric oxide signaling. Nitrolinoleic acid is a hydrophobically stabilized nitric oxide donor. J Biol Chem. 2005 May 13;280(19):19289-97.

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[33] Trostchansky A, Souza JM, Ferreira A, Ferrari M, Blanco F, Trujillo M, et al. Synthesis, isomer characterization, and anti-inflammatory properties of nitroarachidonate. Biochemistry. 2007 Apr 17;46(15):4645-53. [34] Batthyany C, Schopfer FJ, Baker PR, Duran R, Baker LM, Huang Y, et al. Reversible post-translational modification of proteins by nitrated fatty acids in vivo. J Biol Chem. 2006 Jul 21;281(29):20450-63. [35] Baker LM, Baker PR, Golin-Bisello F, Schopfer FJ, Fink M, Woodcock SR, et al. Nitro-fatty acid reaction with glutathione and cysteine. Kinetic analysis of thiol alkylation by a Michael addition reaction. J Biol Chem. 2007 Oct 19;282(42):3108593. [36] Rudolph V, Schopfer FJ, Khoo NK, Rudolph TK, Cole MP, Woodcock SR, et al. Nitro-fatty acid metabolome: saturation, desaturation, beta-oxidation, and protein adduction. J Biol Chem. 2009 Jan 16;284(3):1461-73. [37] Kulinsky VI. Biochemical aspects of inflammation. Biochemistry (Mosc). 2007 Jun;72(6):595-607. [38] Liu H, Jia Z, Soodvilai S, Guan G, Wang MH, Dong Z, et al. Nitro-oleic acid protects the mouse kidney from ischemia and reperfusion injury. Am J Physiol Renal Physiol. 2008 Oct;295(4):F942-9. [39] Nadtochiy SM, Baker PR, Freeman BA, Brookes PS. Mitochondrial nitroalkene formation and mild uncoupling in ischaemic preconditioning: implications for cardioprotection. Cardiovasc Res. 2009 May 1;82(2):333-40. [40] Taylor-Clark TE, Ghatta S, Bettner W, Undem BJ. Nitrooleic acid, an endogenous product of nitrative stress, activates nociceptive sensory nerves via the direct activation of TRPA1. Mol Pharmacol. 2009 Apr;75(4):820-9. [41] Baker PR, Lin Y, Schopfer FJ, Woodcock SR, Groeger AL, Batthyany C, et al. Fatty acid transduction of nitric oxide signaling: multiple nitrated unsaturated fatty acid derivatives exist in human blood and urine and serve as endogenous peroxisome proliferator-activated receptor ligands. J Biol Chem. 2005 Dec 23;280(51):42464-75. [42] Cui T, Schopfer FJ, Zhang J, Chen K, Ichikawa T, Baker PR, et al. Nitrated fatty acids: Endogenous anti-inflammatory signaling mediators. J Biol Chem. 2006 Nov 24;281(47):35686-98. [43] Ferreira AM, Ferrari MI, Trostchansky A, Batthyany C, Souza JM, Alvarez MN, et al. Macrophage activation induces formation of the anti-inflammatory lipid cholesteryl-nitrolinoleate. Biochem J. 2009 Jan 1;417(1):223-34. [44] Kelley EE, Batthyany CI, Hundley NJ, Woodcock SR, Bonacci G, Del Rio JM, et al. Nitro-oleic acid, a novel and irreversible inhibitor of xanthine oxidoreductase. J Biol Chem. 2008 Dec 26;283(52):36176-84. [45] Ichikawa T, Zhang J, Chen K, Liu Y, Schopfer FJ, Baker PR, et al. Nitroalkenes suppress lipopolysaccharide-induced signal transducer and activator of transcription signaling in macrophages: a critical role of mitogen-activated protein kinase phosphatase 1. Endocrinology. 2008 Aug;149(8):4086-94. [46] Khoo NK, Rudolph V, Cole MP, Golin-Bisello F, Schopfer FJ, Woodcock SR, et al. Activation of vascular endothelial nitric oxide synthase and heme oxygenase-1 expression by electrophilic nitro-fatty acids. Free Radic Biol Med. 2009 Oct 24.

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[47] Wright MM, Kim J, Hock TD, Leitinger N, Freeman BA, Agarwal A. Human haem oxygenase-1 induction by nitro-linoleic acid is mediated by cAMP, AP-1 and E-box response element interactions. Biochem J. 2009 Sep 1;422(2):353-61. [48] Iles KE, Wright MM, Cole MP, Welty NE, Ware LB, Matthay MA, et al. Fatty acid transduction of nitric oxide signaling: nitrolinoleic acid mediates protective effects through regulation of the ERK pathway. Free Radic Biol Med. 2009 Apr 1;46(7):866-75. [49] Maines MD. The heme oxygenase system: a regulator of second messenger gases. Annu Rev Pharmacol Toxicol. 1997;37:517-54. [50] Lima ES, Di Mascio P, Abdalla DS. Cholesteryl nitrolinoleate, a nitrated lipid present in human blood plasma and lipoproteins. J Lipid Res. 2003 Sep;44(9):16606. [51] Baker PR, Schopfer FJ, Sweeney S, Freeman BA. Red cell membrane and plasma linoleic acid nitration products: synthesis, clinical identification, and quantitation. Proc Natl Acad Sci U S A. 2004 Aug 10;101(32):11577-82. [52] Schopfer FJ, Batthyany C, Baker PR, Bonacci G, Cole MP, Rudolph V, et al. Detection and Quantification of Protein Adduction by Electrophilic Fatty Acids: Mitochondrial Generation of Fatty Acid Nitroalkene Derivatives. Free Radic Biol Med. 2009 Jan 17. [53] Tsikas D, Zoerner A, Mitschke A, Homsi Y, Gutzki FM, Jordan J. Specific GCMS/MS stable-isotope dilution methodology for free 9- and 10-nitro-oleic acid in human plasma challenges previous LC-MS/MS reports. J Chromatogr B Analyt Technol Biomed Life Sci. 2009 Sep 15;877(26):2895-908. [54] Woodcock SR, Marwitz AJ, Bruno P, Branchaud BP. Synthesis of nitrolipids. All four possible diastereomers of nitrooleic acids: (E)- and (Z)-, 9- and 10-nitrooctadec-9-enoic acids. Org Lett. 2006 Aug 31;8(18):3931-4. [55] Trostchansky A., Bonilla L., et al. Nitroarachidonic acid: A novel peroxidase inhibitor of Prostaglandin Endoperoxide H Synthase 1 and 2. JBC. 2011. In Press

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Chapter 9

DNA Oxidation Jean-Luc Ravanat* Laboratoire "Lésions des Acides Nucléiques" CEA-Grenoble, 38054 Grenoble Cedex 9, France

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1. Introduction This chapter provides an overview of free radical-mediated DNA decomposition reactions. An almost complete degradation pathway of DNA bases initiated by one electron oxidation, hydroxyl radical (HO•) and singlet oxygen is now available. More than 70 DNA lesions have been characterized to date, mostly using model compounds such as isolated nucleosides or short oligonucleotides. Some of them could be detected in vivo following exposure of cells to oxidative stress. Recent data on the formation of complex DNA lesions, generated by only one radical event, will also be highlighted. Various analytical methods developed for measuring oxidatively generated DNA lesions will be presented and discussed, particularly regarding their sensitivity and specificity. Due to space restrictions it is impossible to present all data available in literature; nevertheless, efforts will be made to sensitize the reader to the advantages and limitations of the developed assays, allowing a critical assessment.

*

Tel : +33 (0)4 38 78 47 97; Fax : +33 (0)4 38 78 50 90; Email : [email protected] CEA, DSM/INAC/SCIB UMR-E 3 CEA/UJF

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2. Oxidation of DNA Components by Free Radicals: Mechanistic Aspects of Formation of Single Lesions 2.1. One Electron and HO•-mediated Formation of DNA Lesions Mechanistic insights into one-electron oxidation reactions and HO• radical-mediated formation of modified nucleobases in aerated aqueous solutions were gained from extensive studies involving pyrimidine and purine 2‟-deoxyribonucleosides as relevant DNA model compounds. Information was inferred mostly from the characterisation of the final decomposition products and also from identification of the transiently generated radicals. Importantly, most of the degradation pathways that were uncovered by the model compounds have been validated with oligonucleotides and/or isolated DNA. The degradation pathways of the four DNA bases upon one-electron oxidation reaction and HO•-mediated DNA decomposition are presented in parallel, as some of the produced DNA radicals and thus diamagnetic compounds are generated by both mechanisms 1.

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2.2. Thymine Comprehensive mechanistic information on the one-electron oxidation reactions of the thymine moiety mostly relies on the characterization of final degradation products of the 2methyl-1,4-naphthoquinone (MQ)-mediated sensitization of thymidine (1) with UVA radiation in aerated aqueous solutions [2, 3]. Two main decomposition pathways of thymidine radical cations (2) generated upon one-electron transfer from the base moiety of 1 to triplet excited MQ have been identified, i.e. deprotonation for approximately 30% and a competitive hydration reaction for 70% (Figure 1). Deprotonation of 2 occurs exclusively on the methyl group generating the 5-methylyl-2‟-deoxyuridine radical (3) that upon reaction with molecular oxygen produces unstable 5-hydroperoxymethyl-2‟-deoxyuridine (4); this further decomposes into 5-hydroxymethyl-2‟-deoxyuridine (5) and 5-formyl-2‟-deoxyuridine (6). The second decomposition pathway involves hydration of 2 generating 6-hydroxy-5,6dihydrothymidyl-5-yl radical (7). Reaction of 7 with molecular oxygen produces unstable peroxyl radicals that upon decomposition gives rise mostly to the four cis and trans diastereoisomers of 5,6-dihydroxy-5,6-dihydrothymidine (8), also named thymidine glycols, and to a minor extent to N-(2-deoxy-ß-D-erythro-pentofuranosyl) formamide (9) and 1-(2deoxy-ß-D-erythro-pentofuranosyl)-5-hydroxy-5-methyl-hydantoin (10). Information concerning hydroxyl radical-mediated oxidation reactions of (1) was mostly obtained from the characterization of the bulk of radiation-induced decomposition products of thymidine in aqueous aerated solutions. Under these conditions, the main decomposition products of (1) were identified as thymidine glycols (8) and their mechanism of formation involved addition of HO• produced by water radiolysis onto mostly C6 of 1 to produce 6hydroxy-5,6-dihydrothymidyl-5-yl radical (7), and to a minor extent by addition at C5. Following reaction with molecular oxygen, these two radicals generate unstable hydroperoxides that further decompose to produce mostly 8 and also 9, 10. In addition, the

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two oxidation products of the methyl group (5 and 6) are also produced by reaction of HO• with thymidine. Their formation could be explained by a hydrogen atom abstraction reaction occurring on the methyl group of 1 to produce 3, which further decomposes as described above generating 4, a precursor of 5 and 6.

Figure 1. Main one electron and HO•-mediated decomposition reactions of the base moiety of thymidine (1) in aqueous aerated solution (dR = 2-deoxyribose).

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2.3. Cytosine

Figure 2. Main one electron and HO•-mediated decomposition reactions of the base moiety of 2‟deoxycytidine (1) in aqueous aerated solution.

As reported for thymine, following one electron oxidation of 2‟-deoxycytidine (11), the resulting cytosine radical cation (12) decomposes by two competitive reactions involving either deprotonation or hydration (Figure 2). Deprotonation occurs on the exocyclic amino group yielding aminyl radical 13, which may undergo deamination leading to the formation of 2‟-deoxyuridine (14). Hydration of the cytosine radical cation 12 that is the predominant pathway (80%) occurs mostly at C6 producing 6-hydroxy-5,6-dihydro-2'-deoxycytidyl-5-yl radical (15) as inferred from labelling experiments. Decomposition of 15 is similar to what is observed for the corresponding thymine derivative (7) and generates mainly the four cis and trans diastereoisomers of 5,6-dihydroxy-5,6-dihydro-2'-deocytidine (17) also named cystosine glycols, and also N-(2-deoxy-ß-D-erythro-pentofuranosyl)formamide (9) and 1-(2-

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deoxy-ß-D-erythro-pentofuranosyl)-5-hydroxyhydantoin (16). Cytosine glycols (17) are unstable and further decompose either by a dehydration reaction producing 5-hydroxy-2‟deoxycytidine (18) or though hydrolytic deamination to produce the corresponding 5,6dihydroxy-5,6-dihydro-2'-deoxuridine that upon dehydration gives rise to 5-hydroxy-2‟deoxyuridine (19).

2.4. Guanine

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By analogy to pyrimidine bases, decomposition of the guanine radical cation (21) produced upon one electron oxidation of 2‟-deoxyguanosine (20) may proceed by two mechanisms, either through a hydration reaction or following deprotonation [4] (Figure 3). The former mechanism generates the unstable radical 22 that further decomposes to either 2,6-diamino-4-hydroxy-5-formamidopyrimidine (23) upon reduction, or following oxidation (in the presence of oxygen) to the familiar oxidative DNA lesion, 8-oxo-7,8-dihydro-2‟deoxyguanosine (8-oxodGuo, 24). Deprotonation of guanine radical cation 21 generates radical 25 that upon addition of both a molecule of oxygen and a molecule of water followed by a rearrangement of the purine ring generates 2-amino-5-[(2-deoxy-ß-D-erythropentofuranosyl)amino]-4H-imidazol-4-one (26). The latter nucleoside was found to be unstable and upon hydrolysis is converted quantitatively to 2,2-diamino-4-[(2-deoxy-ß-Derythro-pentofuranosyl)amino]-5-(2H)-oxazolone (27) [5]. Similar products were observed upon HO•-mediated decomposition of 20. Under these conditions, formation of nucleosides 26 and 27 could be explained by the transient formation of radical 25 that is produced upon addition of HO• on C4 followed by a dehydration reaction. Competitive addition of HO• onto the C8 of guanine explains the formation of 23 and 24 though the transient formation of unstable radical 22.

Figure 3. Main one electron and HO•-mediated decomposition reactions of the base moiety of 2‟deoxyguanosine (20) in aqueous aerated solution.

It should be noted that at the nucleoside level, 8-oxodGuo (24), the main 2‟deoxyguaonsine (dGuo) oxidation product in double stranded DNA, is not generated at significant levels. The absence of 8-oxodGuo could be explained by the fact that this nucleoside is consumed as soon as it is produced, by a one electron reaction with guanine

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radical cation (21) or its deprotonated derivative (25) [6]. However, such a reaction has a low probability to occur in DNA and thus, secondary decomposition reaction of 8-oxodGuo in double-stranded DNA is at best a minor process. In addition, it should also be noted that guanine is a critical target for one electron oxidation of DNA because that base has the lowest ionization potential among the DNA constituents. Therefore, upon one-electron mediated DNA oxidation, guanine radical cation 21 is the expected predominant species that could be produced either directly upon oxidation of the guanine base or through efficient charge transfer reactions in double-stranded DNA [7].

2.5. Adenine

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Likewise, following one-electron oxidation of 2‟-deoxyadenosine (28), decomposition of the corresponding radical cation 29 could proceed either by hydration or deprotonation (Figure 4). As reported for dGuo (20) hydration of adenine radical cation is at the origin of two decomposition products, generating through oxidation 8-oxo-7,8-dihydro-2‟deoxyadenosine (32) and under reducing conditions the corresponding formamido-pyrimidine derivative (31). Deprotonation of 29 occurs on the exocyclic amino group producing an aminyl radical (33) that upon decomposition gives rise through deamination to hypoxanthine (34). HO•-mediated formation of 31 and 32 is explained, as reported for guanine, by decomposition of transiently produced radical 30 arising from addition of HO• onto the C8 of the purine base. However, in contrast to what is observed with guanine, no decomposition product of adenine arising from addition of HO• at C4 has been identified so far [2].

Figure 4. Main one electron and HO•-mediated decomposition reactions of the base moiety of 2‟deoxyadenosine (28) in aqueous aerated solution.

2.6. Singlet Oxygen-mediated DNA Oxidation Singlet oxygen in its lowest excited state reacts exclusively with the guanine base (Figure 5). Diels-Alder addition of singlet oxygen (1O2) across C4 and C8 carbons of guanine produces an unstable endoperoxide (35). Decomposition of the latter nucleoside (35)

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produces the corresponding hydroperoxide 37 that decomposes mostly into the two isomers of spironucleosides [8] 38 and to a minor extent into 8-oxodGuo (24). However, in doublestranded DNA, reaction of singlet oxygen with guanine base gives rise exclusively to 8oxoGuo [9]. Notably, 8-oxodGuo can also react efficiently with singlet oxygen and such secondary decomposition reaction also produces spironucleosides 38 as the main decomposition products. Recently it was shown that decomposition of endoperoxide 35 also produces nucleoside 36; this was found specific for singlet oxygen reaction with guanine as 36 is not produced, in contrast to what is observed for 38, by reaction of 1O2 with 8-oxodGuo. Interestingly, 1O2 produced by decomposition of an unstable endoperoxide is able to react with guanine base of cellular DNA to produce 8-oxodGuo [10].

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Figure 5. Main singlet molecular oxygen-mediated degradation pathways of the guanine moiety of dGuo (20).

3. Complex DNA Lesions Generated by a Single Oxidation Event As already mentioned, the single DNA lesions described above were mostly identified by extensive studies performed using nucleosides as DNA model compounds. However, recent data obtained with short oligonucleotides or directly with double-stranded DNA have highlighted the possible formation of more complex DNA lesions produced upon either oneelectron or HO•-mediated DNA oxidation. 3.1. DNA-protein Cross-links Under conditions of oxidative stress, the formation of covalent cross-links between DNA and proteins is well documented [11]. However, only few experiments have been performed to determine the nature of the covalent cross-links, as well as the mechanism of their formation. One study focused on the addition of nucleophilic amino acids onto guanine radical cation produced upon one-electron oxidation. Data available from the literature have

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.

highlighted the electrophilicity of guanine radical cation. Indeed, addition of water at C8 is known to produce 8-oxodGuo (vide supra), while nucleophilic addition of the C5‟ hydroxyl group, a reaction that could not occur in DNA, gives rise to a cyclonucleoside. A similar nucleophilic addition at C8 of 21 was observed with an amino group [12]. More recently, upon one-electron oxidation of a short oligonucleotide containing one guanine residue oxidized in the presence of a tri-lysine peptide, formation of lysine-guanine adduct 39 has been reported [13]. Further results demonstrate that the ε amino group of lysine is able to add to the C8 of the guanine radical cation 21 or its corresponding deprotonated neutral radical 25 (Figure 6). As observed for 8-oxodGuo, the C8-lysine adduct was found capable of undergoing secondary decomposition reactions producing spiro-related nucleosides. Such a nucleophilic addition of amino acids onto the C8 of guanine was also found to occur in double-stranded DNA and with other amino acids such as serine and arginine (Ravanat et al. unpublished).

Figure 6. Mechanism of formation of lysine-guanine cross-link (39) involving nucleophilic addition of lysine (K) onto the C8 of guanine radical cation (21).

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3.2. Complex Lesions Arising from Sugar Oxidation It is generally admitted that HO•-mediated hydrogen abstraction reactions occurring on the 2-deoxyribose moiety of double-stranded DNA give rise to single strand breaks, lesions known to be rapidly repaired in cells. However, recent evidence suggests that complex DNA lesions may also be produced by this mechanism. Experiments were recently performed to search for unknown DNA lesions in double-stranded DNA directly exposed to ionizing radiation [14]. Four new radiation-induced DNA lesions were detected and particular attention was given to one of these lesions that was present in cellular DNA. Characterisation of that lesion and of the mechanism for its formation revealed a complex DNA modification generated following a single oxidation event [15]. Indeed, the initial step involves a hydrogen abstraction at the C4-position of the 2-deoxyribose moiety (Figure 7). Decomposition of the produced radical 40 generates a C4 oxidized abasic site 41 that, following ß-elimination, gives rise to keto-aldehyde 42. Nucleophilic attack of the exocyclic amino group of a cytosine base 11 onto 42 produces adduct 43 identified upon DNA hydrolysis as the four isomers of 6(2-deoxy-ß-D-erythro-pentofuranosyl)-2-hydroxy-3(3-hydroxy-2-oxopropyl)-2,6dihydroimidazo[1,2-c]-pyrimidin-5(3H)-one. Formation of complex DNA lesion 43 highlights the fact that subsequently to a single oxidation event, in this case C4‟-hydrogen abstraction, complex DNA lesions could be produced. Therefore, oxidation of the sugar moiety of double-stranded DNA does not give rise to only single strand breaks. Reaction of the reactive aldehyde 42 was found to occur

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predominantly with a cytosine base located on the complementary strand [16]; thus, the lesion involves a strand break, a modified cytosine base and an inter-strand cross-link. The half-life of that lesion in cells was estimated at about 10 hours, indicating that the repair of such complex DNA damage is relatively slow, as compared to the repair of single oxidized DNA bases.

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Figure 7. Mechanism of formation of a complex DNA lesions 43 produced subsequently to C4‟ hydrogen abstraction.

A similar mechanism was reported for the formation of cross-links arising from a depurination reaction [17]. The detected DNA adduct was similar to that produced by reaction of 4-oxo-2-pentenal with 2‟-deoxycytidine. In addition, ribonolactone, the oxidation product generated following hydrogen abstraction at C1‟ of 2-deoxyribose moiety, was shown able to produce DNA-protein cross-links through nucleophilic addition of amino acids.

3.3. Tandem DNA Lesions The pioneering work of Box and co-workers illustrated that in short oligonucleotides, tandem DNA lesions, containing adjacent base damages on the same strand, could be produced at a high yield [18]. In addition, it has been reported that such tandem lesions could be generated, at least in short oligonucleotides, by a single oxidation event [17]. Covalent cross-links between the methyl group of thymidine and the C8 of either guanine or adenine can be formed between two adjacent DNA bases. Such tandem lesions result from addition of the 5-methylyl-2‟-deoxyuridine radical (3) onto the C8 of a purine base located preferentially at its 3‟-relative position [19]. Formation of the cross-link thymidine-purine tandem lesions is more efficient with guanine compared to adenine and can be inhibited by the presence of oxygen, providing a tentative explanation for the fact that the yield of radiation-induced formation of such damage in cellular DNA is very low. Adducts between C5 of cytosine and C8 of guanine are also produced meanwhile at very low yield in cellular DNA [20]. Pyrimidine-purine cross-link formation is inhibited by the presence of oxygen, because initially produced radicals rapidly react with molecular oxygen to produce peroxyl radicals.

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However, such peroxyl radicals are able to react with surrounding DNA bases, thus producing tandem lesions comprised of adjacent oxidized DNA bases [21]. Indeed, formation of tandem lesions containing formylamine (dF) and 8-oxodGuo account for about 10% of totally produced 8-oxodGuo upon ionizing radiation of aqueous aerated DNA solutions. The formation of dF/8-oxodGuo tandem lesions (Figure 8) is due to addition of a pyrimidine peroxyl radical 44 onto the C8 of a vicinal guanine base [22] creating endoperoxide 45 between a pyrimidine and the C8 of guanine. Decomposition of the unstable endoperoxide 45 produces 8-oxodGuo and a vicinal formylamine residue (46). In fact, as reported recently [23], the latter mechanism accounts for about 50% of totally produced 8-oxodGuo and 8oxodAdo in double-stranded DNA exposed to hydroxyl radicals. Moreover, in doublestranded DNA, peroxidation reactions were found to increase the yield of formation of 8oxodGuo by a factor of 20. Such an increase is explained only partly by the formation of tandem lesions generated upon addition of pyrimidine peroxyl radicals onto the C8 of guanine. In addition, such an increase is also due to a one-electron transfer mechanism, from guanine to peroxyl radicals [23]. This proposed mechanism provides, for the first time, an elegant explanation for the high yield of formation of 8-oxodGuo compared to 8-oxodAdo observed in double-stranded DNA.

Figure 8. Mechanism of formation of tandem DNA lesions 8-oxodGuo/dF 46 involving initial formation of a peroxyl radical.

4. Method for Measuring Oxidative DNA Lesions in Cells Measuring cellular levels of oxidized DNA bases poses a challenging analytical problem [24]. The major difficulty arises from the fact that the method should allow the detection of DNA lesions at levels of around one modification per million DNA bases using a few µg of DNA. Another problem is that DNA oxidation may occur during the sample preparation, generating spurious DNA lesions, and resulting in overestimated results. During the last three decades, considerable efforts have been made to use oxidized DNA lesions as potential biomarkers of in vivo oxidative stress. Several analytical approaches have been developed for such a purpose (Figure 9), which may be classified into two categories: a) Those using physico-chemical analytical approaches subsequently to DNA extraction and digestion (direct approaches), and b) those requiring biochemical tools (indirect approaches). The objective of this Chapter is not to cover comprehensively the abundant literature in this field, but to

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provide readers with sufficient background to evaluate critically the reported results. Many of them should be considered with caution because methods employed in the past to measure oxidative DNA lesions were not always appropriate.

Indirect approaches

Direct approaches Cells

Alkaline cell lysis DNA N-glycosylase

DNA extraction

Underestimation ?

Strand break detection

Comet/Fpg AE/Fpg

Overestimation ?

Antibodies

Fluorescence detection (ELISA) Radioactive detection (RIA)

DNA digestion or hydrolysis

Overestimation ? Underestimation ?

Nucleotides or nucleosides or bases 32

P-ATP

Overestimation 32P-Label

derivatization Overestimation

HPLC-EC GC-MS HPLC-MS/MS

Figure 9. Schematic representation of the available methods aimed at measuring cellular levels of oxidative DNA lesions. Possible causes for under- or overestimations are highlighted.

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4.1. Direct Approaches Following DNA extraction from cells or tissues, DNA is digested to the nucleoside level and then modified nucleosides are detected using appropriate analytical tools [25]. With this technique, most attention has been focused on 8-oxodGuo, a popular marker of oxidized DNA lesions since the mid-1980s, when an HPLC assay coupled to electrochemical detection (HPLC-EC) was developed for its detection. To obtain reliable and non-underestimated data, a quantitative digestion of DNA into nucleosides is required. Enzymatic digestion with nuclease P1 is appropriate, at least for detection of 8-oxodGuo. A cocktail of endo- and exonucleases may be used for lesions that cannot be released by nuclease P1 alone. At least for 8-oxodGuo, hydrolysis with formic acid can also be employed; however, this method is unsuitable for unstable bases like formamidopyrimidine derivatives 23 and 31. Recent results confirmed that enzymatic digestion is free of significant artifactual DNA oxidation [26, 27]. Gas-chromatography coupled to mass spectrometry (GC-MS) was also developed for measuring simultaneously several oxidized DNA bases [28]. This technique requires acidic DNA hydrolysis for the release of lesions as isolated bases that are further derivatized for detection by GC-MS. Interestingly, it appears that results obtained by the GC-MS assay are on average significantly higher than respective results obtained by HPLC-EC [29, 30]. This discrepancy was later attributed to spurious DNA oxidation occurring during the derivatization step of the GC-MS assay that is performed at high temperatures [31].

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Therefore, published data obtained by the controversial GC-MS assay should be considered with caution, unless the lesion was purified prior to derivatization [32]. At the end of the last century, HPLC coupled to tandem mass spectrometry through electrospray ionization became available and was applied for the measurement of 8-oxodGuo [33] and other oxidized DNA lesions [34]. This method has been extensively used during the last decade and appears to offer the most suitable means for measuring low levels of lesions because it is highly specific, sensitive and versatile. It should be emphasized that extraction of DNA from cells or tissues, which is necessary for the detection of lesions, constitutes another (and probably the major) potential source of spurious oxidation. Therefore, considerable work has been done, mostly through the European project ESCODD, to minimize this problem. Nowadays, optimized protocols are available [26, 35]. A 32P-post-labelling approach, initially developed for measuring bulky DNA adducts, was tentatively applied to oxidized DNA bases; however, spontaneous selfradiolysis occurring during sample preparation does not allow accurate determination of oxidized DNA lesions in cellular DNA [25].

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4.2. Indirect Approaches Several approaches have been developed for measuring DNA strand breaks in cellular DNA. These include the so-called single cell gel electrophoresis, also known as the “comet assay” [36], and the alkaline elution (AE) [37] and alkaline unwinding [38] techniques. These methods are very sensitive to measure DNA single-strand breaks and were also adapted for the measurement of oxidized DNA bases. To this end, DNA repair enzymes, such as Fpg, are used to convert oxidized bases into strand breaks that are then quantified. It is generally admitted that the amount of detected Fpg sensitive sites correlates to the amount of 8oxodGuo, a lesion that is efficiently excised by Fpg protein. The measurement of DNA damage by such indirect approaches does not require isolation of cellular DNA thus reducing the risk of potential spurious DNA oxidation. In addition, compared to direct approaches, the amount of cells required for the analysis is much lower and the sensitivity of indirect approaches is higher. The major limitation is that these approaches are not quantitative. For instance, results obtained by the comet assay are expressed as the percentage of DNA in the tail of the comet that does not directly correlate to the amount of damaged DNA per cell. To overcome this limitation, calibration of the assay can be performed using ionizing radiation, which is considered to induce 1000 single-strand breaks per cell and per Gy [39]. In general, results obtained by the indirect approaches are significantly lower, sometimes by several orders of magnitude, than those reported by direct methods (Figure 10). By evaluating only recent data obtained with optimized protocols, the gap between the results is reduced [40] to a factor of 3 to 5 and background levels of 8-oxodGuo in untreated cells are considered to vary between 0.1 to 1 8-oxodGuo per million nucleosides. The discrepancy between the reported data is generally attributed to an overestimation of the level of 8oxodGuo when measured using direct approaches due to spurious DNA oxidation during sample preparation (mostly during DNA extraction). However, potential underestimation of data obtained by indirect approaches should be also considered. Indeed, recent results showing that Fpg protein is unable to excise quantitatively 8-oxodGuo from tandem lesions

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produced in high yields by a single oxidation event [23] strongly suggest that methods using DNA glycosylases may underestimate the level of 8-oxodGuo. In addition, it is well known that Fpg protein has a broad specificity and Fpg sensitive sites are not merely constituted of 8oxodGuo. Moreover, it is also possible that not all lesions are accessible to the enzyme when DNA is embedded in the agarose gel. The employment of the hOGG protein may, at least partially, overcome this limitation [41], at least when 8-oxodGuo is not involved in tandem damage. Antibody-based methods have also been developed for measuring oxidized DNA lesions [42]. However, due to lack of sufficient specificity of antibodies, these methods are not appropriate for measuring very low levels of damage encountered in cellular DNA [43], unless they are coupled to chromatographic techniques. Nevertheless, immunohistochemical localization of 8-oxodGuo has been applied successfully to localize oxidative lesions in tissues [42]. The difficulties encountered during measurement of oxidative DNA lesions explain the significant variability and inconsistency of reported levels for 8-oxodGuo background levels in untreated cells (Figure 10). High values that are not biologically relevant could be attributed to inappropriate analytical approaches (vide supra). It should also be emphasized that significant variations were observed even between studies using the powerful HPLCMS/MS analytical tools, but this could be explained by the lack of optimized DNA extraction protocols. Most of the origins of the drawbacks have now been identified, and recent results obtained following optimization of the analytical assays confirm that the background levels of 8-oxodGuo in untreated cells are around one oxidative DNA lesion per million nucleosides. Only few reports on other oxidatively generated DNA lesions are currently available, and it appears that their levels are lower than those of 8-oxodGuo [44].

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Cellular background levels of 8-oxodGuo 32P

Label

GC-MS HPLC-EC HPLC-MS/MS RIA ELISA Comet/Fpg AE/Fpg 8-oxodGuo/ 106 DNA bases

0.1

1

10

100

1000

Figure 10. Overview of the background levels of 8-oxodGuo reported in the literature. The illustration indicates that some methods are clearly not appropriate for measuring cellular levels of 8-oxodGuo. Even by using a defined method such as HPLC-EC, significant variations are observed and these could be explained by variability in DNA extraction protocols.

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Conclusions The measurement of oxidative lesions in cellular DNA is crucial for understanding biological responses to oxidative stress. By surveying relevant literature several inconsistencies are apparent, very likely due to technical limitations, suggesting that many previous results should be interpreted with caution. Recent results illustrated that decomposition reactions of initially produced DNA radicals in double-stranded DNA may be influenced by their environment; thus the importance of complex DNA lesions is probably underestimated. The repair of such complex lesions appears to be, at least in vitro, less efficient than that of single lesions. Therefore, the biological importance of such damage should be a focus of future research. The development of accurate analytical assays, such as HPLC coupled to tandem mass spectrometry, aimed at detecting specific DNA lesions, could be useful to reassess the decomposition pathways of transiently produced DNA radicals in vivo.

References

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[1]

Cadet J, Douki T, Gasparutto D, Ravanat J-L. Oxidative damage to DNA: formation, measurement and biochemical features. Mutat Res 2003;531:5-23. [2] Cadet J, Carell T, Cellai L, et al. DNA damage and radical reactions: mechanistic aspects, formation in cells and repair Studies. Chimia 2008;62:742-9. [3] Ravanat J-L, Douki T, Cadet J. Direct and indirect effects of UV radiation on DNA and its components. J Photochem Photobiol B 2001;63:88-102. [4] Cadet J, Douki T, Ravanat JL. Oxidatively generated damage to the guanine moiety of DNA: mechanistic aspects and formation in cells. Acc Chem Res 2008;41:10757083. [5] Cadet J, Berger M, Buchko GW, Joshi PC, Raoul S, Ravanat J-L. 2,2-Diamino-4[(3,5-di-O-acetyl-2-deoxy-ß-D-erythro-pentofuranosyl)amino]-5-(2H)- oxazolone : a novel and predominant radical oxidation product of 3',5'-di-O-acetyl-2'deoxyguanosine. J Am Chem Soc 1994;116:7403-4. [6] Ravanat J-L, Saint-Pierre C, Cadet J. One-electron oxidation of the guanine moiety of 2'-deoxyguanosine: influence of 8-oxo-7,8-dihydro-2'-deoxyguanosine. J Am Chem Soc 2003;125:2030-1. [7] Douki T, Ravanat J-L, Angelov D, Wagner RJ, Cadet J. Effects of duplex stability on charge-transfer efficiency within DNA. Top Current Chem 2004;236:1-25. [8] Ravanat J-L, Cadet J. Reaction of singlet oxygen with 2'-deoxyguanosine and DNA. Isolation and characterization of the main oxidation products. Chem Res Toxicol 1995;8:379-88. [9] Ravanat J-L, Martinez GR, Medeiros MGH, Di Mascio P, Cadet J. Singlet oxygen oxidation of 2'-deoxyguanosine. Formation and mechanistic insights. Tetrahedron 2006;62:10709-15. [10] Ravanat J-L, Di Mascio P, Martinez GR, Medeiros MH, Cadet J. Singlet oxygen induces oxidation of cellular DNA. J Biol Chem 2000;275:40601-4.

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[11] Barker S, Weinfeld M, Murray D. DNA-protein crosslinks: their induction, repair, and biological consequences. Mutat Res 2005;589:111-35. [12] Cadet J, Douki T, Ravanat J-L. One-electron oxidation of DNA and inflammation processes. Nat Chem Biol 2006;2:348-9. [13] Perrier S, Hau J, Gasparutto D, Cadet J, Favier A, Ravanat J-L. Characterization of lysine-guanine cross-links upon one-electron oxidation of a guanine-containing oligonucleotide in the presence of a trilysine peptide. J Am Chem Soc 2006;128:5703-10. [14] Regulus P, Spessotto S, Gateau M, Cadet J, Favier A, Ravanat J-L. Detection of new radiation-induced DNA lesions by liquid chromatography coupled to tandem mass spectrometry. Rapid Commun Mass Spectrom 2004;18:2223-8. [15] Regulus P, Duroux B, Bayle P-A, Favier A, Cadet J, Ravanat J-L. Oxidation of the sugar moiety of DNA by ionizing radiation or bleomycin could induce the formation of a cluster DNA lesion. Proc Nat Acad Sci, USA 2007;104:14032-7. [16] Sczepanski JT, Jacobs AC, Greenberg MM. Self-promoted DNA interstrand crosslink formation by an abasic site. J Am Chem Soc 2008;130:9646-7. [17] Wang Y. Bulky DNA lesions induced by reactive oxygen species. Chem Res Toxicol 2008;21:276-81. [18] Box HC, Dawidzik JB, Budzinski EE. Free radical-induced double lesions in DNA. Free Radic Biol Med 2001;31:856-68. [19] Bellon S, Ravanat J-L, Gasparutto D, Cadet J. Cross-linked thymine-purine base tandem lesions: Synthesis, characterization, and measurement in gamma-Irradiated isolated DNA. Chem Res Toxicol 2002;15:598-606. [20] Hong H, Cao H, Wang Y. Formation and genotoxicity of a guanine-cytosine intrastrand cross-link lesion in vivo. Nucleic Acids Res 2007;35:7118-27. [21] Hong IS, Carter KN, Sato K, Greenberg MM. Characterization and mechanism of formation of tandem lesions in DNA by a nucleobase peroxyl radical. J Am Chem Soc 2007;129:4089-98. [22] Douki T, Riviere J, Cadet J. DNA tandem lesions containing 8-oxo-7,8dihydroguanine and formamido residues arise from intramolecular addition of thymine peroxyl radical to guanine. Chem Res Toxicol 2002;15:445-54. [23] Bergeron F, Auvré F, Radicella JP, Ravanat J-L. HO° radicals induce an unexpected high proportion of tandem base lesions refractory to repair by DNA glycosylases. Proc Nat Acad Sci USA 2010;107:5528-33. [24] Ravanat J-L. Measuring oxidized DNA lesions as biomarkers of oxidative stress: An analytical challenge. FABAD, J Pharm Sci 2005;30:100-13. [25] Cadet J, Douki T, Ravanat J-L. Artifacts associated with the measurement of oxidized DNA bases. Environ Health Perspect 1997;105:1033-9. [26] Ravanat J-L, Douki T, Duez P, et al. Cellular background level of 8-oxo-7,8dihydro-2'-deoxyguanosine: an isotope based method to evaluate artefactual oxidation of DNA during its extraction and subsequent work-up. Carcinogenesis 2002;23:1911-8. [27] Huang X, Powell J, Mooney LA, Li C, Frenkel K. Importance of complete DNA digestion in minimizing variability of 8-oxo-dG analyses. Free Radic Biol Med 2001;31:1341-51.

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[28] Dizdaroglu M. The use of capillary gas chromatography-mass spectrometry for identification of radiation induced DNA bases damage and DNA base-amino acid cross-links. J Chromatogr 1984;295:103-21. [29] Halliwell B, Dizdaroglu M. Commentary. The measurement of oxidative damage to DNA by HPLC and GC/MS techniques. Free Radic Res Commun 1992;16:75-87. [30] Hamberg M, Zhang L-Y. Quantitative determination of 8-hydroxyguanine and guanine by isotope dilution mass spectrometry. Anal Biochem 1995;229:336-44. [31] Ravanat J-L, Turesky RJ, Gremaud E, Trudel LJ, Stadler RH. Determination of 8oxoguanine in DNA by gas chromatography-mass spectrometry and HPLCelectrochemical detection. Overestimation of the background level of the oxidized base by the gas chromatography-mass spectrometry assay. Chem Res Toxicol 1995;8:1039-45. [32] Douki T, Ravanat J-L, Cadet J. Measurement of oxidized bases in DNA and biological fluids by gas chromatography coupled to mass spectrometry. In: Lunec J, Griffiths HR, eds. Measuring in vivo oxiative damage: A practical course. Chichester: John Wiley & Sons, LTD; 1998:15-26. [33] Singh R, Sweetman GM, Farmer PB, Shuker DE, Rich KJ. Detection and characterization of two major ethylated deoxyguanosine adducts by high performance liquid chromatography, electrospray mass spectrometry, and 32Ppostlabeling. Development of an approach for detection of phosphotriesters. Chem Res Toxicol 1997;10:70-7. [34] Frelon S, Douki T, Ravanat J-L, Pouget J-P, Tornabene C, Cadet J. High performance liquid chromatography - tandem mass spectrometry for the measurement of radiation-induced base damage to isolated and cellular DNA. Chem Res Toxicol 2000;13:1002-10. [35] Hofer T, Moller L. Optimization of the workup procedure for the analysis of 8-oxo7,8- dihydro-2'-deoxyguanosine with electrochemical detection. Chem Res Toxicol 2002;15:426-32. [36] Collins AR, Dobson VL, Dusinska M, Kennedy G, Stetina R. The comet assay: what can it really tell us? Mutat Res 1997;375:183-93. [37] Epe B, Pflaum M, Boiteux S. DNA damage induced by photosensitization in cellular and cell-free systems. Mutat Res 1993;299:135-45. [38] Hartwig A, Dally H, Schlepegrell R. Sensitive analysis of oxidative DNA damage in mammalian cells: use of the bacterial Fpg protein in combination with alkaline unwinding. Toxicol Lett 1996;88:85-90. [39] Pouget J-P, Ravanat J-L, Douki T, Richard M-J, Cadet J. Measurement of DNA base damage in cells exposed to low doses of gamma radiation: comparison between the HPLC-EC and the comet assays. Int J Radiat Biol 1999;75:51-8. [40] ESCODD. Measurement of DNA oxidation in human cells by chromatographic and enzymic methods. Free Radic Biol Med 2003;34:1089-99. [41] Smith CC, O'Donovan MR, Martin EA. hOGG1 recognizes oxidative damage using the comet assay with greater specificity than FPG or ENDOIII. Mutagenesis 2006;21:185-90. [42] Toyokuni S, Tanaka T, Hatton Y, et al. Quantitative immunohistochemical determination of 8-hydroxy-2'-deoxyguanosine by a monoclonal antibody N45.1: Its

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application to ferric nitrilotriacetate-induced renal carcinogenesis model. Laboratory Investigation 1997;76:365-74. [43] Breton J, Sichel F, Bianchini F, Prevost V. Measurement of 8-hydroxy-2'deoxyguanosine by a commercial available ELISA test: Comparison with HPLC/Electrochemical detection in calf thymus DNA and determination in human serum. Anal Lett 2003;36:123-34. [44] Cadet J, Douki T, Frelon S, Pouget J-P, Sauvaigo S, Ravanat J-L. Assessment of oxidative base damage to isolated and cellular DNA by HPLC-MS/MS measurement. Free Radic Biol Med 2002;33:441-9.

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 10

Methods of Investigation of Selected Radical Oxygen/Nitrogen Species in Cell-free and Cellular Systems Jacek Zielonka* and Balaraman Kalyanaraman Department of Biophysics and Free Radical Research Center, Medical College of Wisconsin, 8701 Watertown Plank Road, Milwaukee, WI 53226 U.S.

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1. Introduction The methodology of investigation of biologically-relevant free radicals both in cell-free and cellular systems is of great importance for the understanding of physiological and pathological functions of those species in living organisms [1]. In fact, in many cases the progress in understanding the role of radical species in biological systems is hampered by the lack of reliable methods for their specific detection and quantification [2, 3]. One of the reasons for that difficulty lies in the intermediary character of radicals, due to their short lifetime, especially in biological systems. Therefore, an array of probes has been developed, which react directly with the radical species of interest and form a relatively stable and easily detectable product [1-15]. However, in many cases due to the lack of the understanding of the chemistry behind the assay, the interpretation of the experimental data remains questionable [2, 3, 16-18]. Not only specificity of the probe towards specific radical species needs to be tested, but also the reaction mechanism needs to be established and the potential confounding effects of different compounds present in cellular systems should be examined. Therefore in this Chapter the methods of detection as well as of generation of selected free radicals in both cellular and cell-free systems will be discussed.

*

Address correspondence to: Jacek Zielonka, Department of Biophysics, Medical College of Wisconsin, 8701 Watertown Plank Road, Milwaukee, WI 53226, USA. Phone: 414-955-4789; Fax: 414-456-6512; E-mail: [email protected]

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2. General Principles of the Detection of Free Radicals in Chemical and Biological Systems 2.1. Methods of Investigation of Free Radicals Due to the short life-time of most radicals, their direct detection is limited to fast, kinetic methods, combined with pulse techniques of their generation. The two most commonly used techniques for that purpose are pulse radiolysis and laser flash photolysis, both of them generating relatively high concentration of radicals (~ µM) at the sub-microsecond timescale. For detection of the radicals formed by these techniques, the fast spectrophotometric method is most widely used. Other fast kinetic methods include electron paramagnetic resonance (EPR) spectroscopy, which can detect paramagnetic compounds and electrochemical methods, including conductometry, which responds to the changes in the conductivity of the solutions, typically due to acid-base equilibration occurring during radical reactions. In some cases also steady-state methods of detection, again including spectrophotometry, EPR spectrometry and electrochemical techniques, can be used, for example during continuous mixing of the reagent with the formation of radical intermediate, which reaches the steady-state concentration that is detectable by the method used. In most cases, however, the concentrations of the free-radical intermediates are below the detection limit, and the addition of the radical-reactive probes is necessary.

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2.2. Probes for Free Radical Detection The application of the probes for the detection of radical species is based on the difference in the specific properties between the probe and the product of the reaction of the probe with the radical investigated. Due to the availability of the instrumentation in most laboratories and the possibility of non-invasive, real time monitoring of the conversion of the probe into the products, the difference in the spectroscopic properties are most widely employed in the probe design and use. Generally, the spectroscopic probes can be divided into several categories, depending on the methods of the detection, including: Spectrophotometry Chemiluminescence Fluorescence Electron paramagnetic resonance (EPR) In many cases, however, the probe can be a member of more than one category. For example, this applies to probes, which upon reaction with the radical, yield the fluorescent product of different electronic absorption spectra than the parent compound. The need for the knowledge of the extent of probe uptake and for the separation of the possibly multiple reaction products with the specific quantification of each of them implicates the requirement for use of chromatographic techniques (e.g. HPLC, GC). Typical HPLC systems are equipped with UV-Vis absorption, fluorescence, electrochemical and/or mass

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detectors, which again may not fit to the above-mentioned categories. For example the fluorescent products can also be detected in many cases with high sensitivity using electrochemical and/or MS detectors. Spectrophotometric Probes Spectrophotometric detection is based on the changes occurring in the UV-Vis absorption spectrum during the process of trapping of the radical R• by the probe (reaction 1). probe + R• →→ product(s)

(1)

The sensitivity of the assay is proportional to the difference of the extinction coefficients between the product and the probe at the wavelength used. Due to the low transmittance of biological materials for UV and visible light, the use of spectrophotometric probes for realtime monitoring of radical formation is limited almost exclusively to cell-free systems, cell lysates or detection of radicals released from cells to extracellular medium. In case of endpoint measurements, the spectrophotometric probes can be also used intracellularly and before the measurement the samples are prepared in a way to minimize light absorption/scattering by cell components. Chemiluminescent Probes Chemiluminescent probes react with the radicals with the emission of light during the reaction (see also Chapter 12). Actual mechanism involves the formation of the product in the excited state (product)*, which relaxes to the ground state with the emission of light of energy h (reaction 2).

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probe + R• →→ (product)* → product + h

(2)

As the chemiluminescent detection is typically highly sensitive, very low levels of radicals can be measured using this method. Fluorogenic Probes Fluorogenic probes (also called fluorescent probes) react with the radicals to form highly fluorescent products (reaction 3). probe (non-fluorescent) + R• →→ product (fluorescent)

(3)

The probes are typically non- or minimally-fluorescent, at least in the spectral region where the product exhibits fluorescence. The sensitivity of the assay is proportional not only to the product‟s absolute quantum yield of the fluorescence, but also to the difference in the fluorescence intensity between the product and the probe itself. Thus, even if the product is highly fluorescent, but the probe is fluorescent per se to a significant degree, the fluorescence of the product due to low amount of radicals will be masked by the probe‟s autofluorescence.

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EPR Spin Traps and Probes EPR traps and probes yield persistent free radicals during the reaction with short-lived radical species. Two major classes of EPR probes involve spin probes, which react with the short-lived radical R• to from a stable radical (e.g. conversion of hydroxylamines into nitroxides, reaction 4) and spin traps which form adducts with radicals, resulting in secondary, long-lived radicals called spin adducts (e.g. reaction of nitrones and nitroso compounds to form nitroxides, reactions 5 and 6, respectively). In contrast to spin probes, which are typically non-specific, the spin traps yield the product, which in most cases is specific for the radical trapped and thus also provides structural information about the trapped species. +

R•

+

N OH

+ N O

H

NO R3

R• N O

R1 R2

RH

N O

+

R•

(4) R H

(5)

R1 R2

O N R3 R

(6)

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While the EPR technique could be used to detect directly the radical of interest (R•), its steady-state concentration is typically below the detection limit in biological systems. Although the amount of the secondary radical (typically nitroxide) cannot be higher than the total amount of the primary radical R•, due to its longer life-time it can accumulate to a higher extent and thus exceed the threshold of the detection limit of the EPR instrument. The Properties of the Ideal Probe There are several requirements for the reliable use of probe for detection and quantification of the radical of interest: A. The probe should be present at concentration high enough to effectively compete with other pathways of radical decay, and thus scavenge the whole pool of the radical formed. Small changes in the concentration of the probe should not affect the scavenging efficacy. B. At the concentration used, the probe should not be toxic, nor modify the rate of radical production. C. The probe should react selectively with the radical of interest and/or produce easily distinguishable, radical-specific product. D. The product should be stable under experimental conditions and its amount should be proportional to the amount of radical formed. E. The probe, its end-product of the reaction with the radical, or any reaction intermediate should not be a source of the radical or in any other way affect the rate of the radical production during the assay.

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F. It is preferable that the product formation rather than the probe consumption be monitored, as at low concentration of radicals the relative increase in the signal from the product will be higher than relative decreases in the signal intensity of the probe. In reality, there is no perfect probe and one has to understand the limitation of the assay in order to properly design the experiment and to correctly interpret the experimental data. While in cell-free systems controlling of the factors, like the probe‟s concentration and product quantification are typically straightforward, in cellular systems the situation is in many cases more problematic, including achievement of the proper intracellular concentration of the probe, quantification of the product(s) and minimizing the interference with the assay by intracellular components. One way to overcome the cell loading problem, especially in the case of hydrophilic probes, is to design the probe carrying carboxylic group(s) (probe(COOH)n) and prepare their esters (probe-(COOR)n). The ester (probe-(COOR)n) is not charged and can passively diffuse into cells, where it undergoes esterase-catalyzed hydrolysis to (probe-(COO-)n, reaction 7), which being negatively charged is trapped inside the cells. probe-(COOR)n + n -OH → probe-(COO-)n + n ROH

(7)

A similar approach can be used for phenolic compounds, which can form esters with carboxylic acids, and after hydrolysis yield a phenolate anionic probe. An example are the diacetate derivatives of fluorescein-based probes (reaction 8).

O

COOH

O O

O

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O

O

O

fluorescein diacetate

esterase

+ HO

O

2 CH3COOH

O

fluorescein

(8) Reactivity of the Intermediates of the Reaction between the Probe and Radical R• The typical problem in the quantification of radical species (R•) using probes comes from the radical character of the primary product of the reaction of the radical R• with the probe (the formation of probe-derived radical P•, reaction 9). probe + R• → P• + R

(9)

Due to very fast and in many cases barrierless radical-radical recombination reactions, the probe-derived radical P• may compete with the probe itself for the radical of interest (reaction 10). P• + R• → product(s)

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(10)

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This reaction will not only deplete the steady-state concentration of the radical R•, but in many cases may also prevent the formation of the product being monitored, leading to significant underestimation of the amount of radical R• being produced. However, in some cases (e.g. hydroethidine- and lucigenin-based assays for superoxide radical anion) the reaction between P• and R• is responsible for the formation of the product being measured, and thus is beneficial. Depending on the nature of the probe-derived radical P•, it may also lead to formation of additional radicals, by reduction or oxidation of the system components. Some of the important reactants to consider in biological systems include oxygen, reduced glutathione (GSH) and reduced nicotinamide adenine dinucleotide NAD(P)H. If the radical P• is reactive towards oxygen, it may form secondary peroxyl radical (POO•, reaction 11) or directly reduce oxygen to superoxide radical anion, O2•- (reaction 12). P• + O2 → POO•

(11)

P• + O2 → P(-H) + H+ + O2•-

(12)

The probe-derived peroxyl radical POO• is an oxidizing radical (see below) and in many cases may be able to oxidize the parent probe molecule (reaction 13), thus leading to chain reaction of probe oxidation. POO• + probe → POOH + P•

(13)

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If the radical P• (or POO•) reacts with GSH, it forms thiyl radical GS• (reaction 14 and 15), which may react with thiolate anion (GS-) or thiol (GSH) to form radical anion of disulfide GSSG•- (reaction 16), which in the presence of oxygen will produce O2•- (reaction 17) [19]. P• + GSH → P + GS•

(14)

POO• + GSH → POOH + GS•

(15)

GS• + GS-(or GSH) → GSSG•- (+ H+)

(16)

RSSR•- + O2 → RSSR + O2•-

(17)

Similarly, the reaction of P• with reduced nicotinamide coenzymes NAD(P)H may lead to the radical NAD(P)• (reaction 18), which in the presence of oxygen will yield O2•- (reaction 19) [20]. P• + NAD(P)H → P + NAD(P)•

(18)

NAD(P)• + O2 → NAD(P)+ + O2•-

(19)

These examples show how the probe-derived radicals may generate an additional pool of radicals (O2•- and possibly other secondary radicals) regardless of their reductive or oxidative Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

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properties. Those reactions will typically lead to overestimation of the rate of production of the radical under investigation. In some cases they may even mislead in terms of the identity of the radical produced in the system. Other possible reactions of the radical P• in the investigated system should be also considered. Therefore, an understanding of the mechanism of the reaction between the probe molecule and the radical species R• as well as of the chemical reactivity of any reaction intermediate formed (exemplified above as the probe radical P•) is essential.

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2.3. Radical Footprints The alternative and complementary techniques to the use of external spectroscopic probes for detection of free radicals in biological systems is to monitor the radical-induced modifications of cellular components, including DNA, proteins, lipids, glutathione, etc [1]. Therefore, in those techniques, termed “footprinting” or “fingerprinting”, the cellular constituents can be regarded as natural “probes” for the radicals. The advantage of those assays lies in the fact that there is no need for the supply of the xenobiotics (probes), so there is no interference with the biological processes being studied. Examples of the products of radical-induced modifications to DNA include 8-hydroxydeoxyguanosine and degradation products of deoxyribose (Chapter 9). In case of radical-induced post-translational protein modifications, the most-widely used markers include protein carbonyls, nitrotyrosine (indicating the presence of nitrogen-containing radical(s)) and dityrosine (Chapter 6). The products of the reaction of lipids with radical species include hydroperoxides, conjugated dienes, aldehydes (e.g. malondialdehyde, 4-hydroxynonenal) and isoprostanes (Chapter 7). With a few exceptions (e.g. monitoring of fluorescence due to dityrosine) the footprinting techniques rely on methods of analysis of destructive character, and thus are limited to endpoint measurements. The lack of specificity of the product detected to the radical species of interest is one of the major problems with those techniques, and with the progress in understanding of the oxidative chemistry of biomolecules, often comes the realization of a lack of specificity towards specific radicals of interest. In addition to the possibility of an artificial rise of the level of oxidatively modified biomolecules during processing of the biological samples for analysis, the cellular repair and/or secretion of the products of interest may be confounding factors even in semi-quantitative analysis of the production of radical species. As in most cases the radical “footprints” are not the direct products of the reaction with the radicals, these methods will not be discussed further in this Chapter.

2.4. Scavengers and Inhibitors Whenever possible, the detection of the radical should be accompanied by the use of specific scavenger(s) of the radical, to confirm the specificity of the assay and/or to establish the portion of the detected signal attributable to this radical. The requirements for the radical scavengers are similar to the ones for the probes themselves:

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Jacek Zielonka and Balaraman Kalyanaraman A. Should be selective towards the radical of interest. In case of low-molecular weight compounds, the specificity towards specific radical is questionable, with the exception of the hydroxyl radical due to its high reactivity, and thus the possibility of the reaction with compounds which are not reactive towards other radicals/oxidants. In case of scavengers of less reactive radicals, they can also react with more reactive radicals so they are less selective. An example of a scavenger of high selectivity includes superoxide dismutase (SOD), which selectively scavenges superoxide radical anion and is widely used in cell-free assays for O2•-. B. Should be present at concentration high enough to efficiently compete for the radical. The amount of the scavenger used in cell-free assay is typically limited by its solubility and/or the possible interference with the assay used. Therefore scavenging efficiency is not a major problem. In biological systems, low-molecular weight scavengers may or may not accumulate intracellularly, depending on the physicochemical characteristics of the compound. In many cases the rational modification of the scavenger‟s chemical structure may result in better cellular uptake, while its radical scavenging activity is retained. The situation is different for enzymatic scavengers. For instance, the applicability of SOD for the cellular assays for O2•- is limited to the extracellularly detected O2•- because of the lack of cell membrane permeability of the enzyme. The strategies to increase the intracellular concentration of the enzymatic scavengers include the attachment of polyethylene glycol (PEG) chains to the enzyme to increase its membrane permeability or the stimulation of intracellular expression of the enzyme using molecular biology techniques. C. The scavenger should not affect the rate of radical production. In many cases, based on the known chemical reactivity of the compound towards a specific radical, it has been successfully applied to protect the cells from the oxidative insult. At the same time the compound inhibits the oxidation of the probe used. This may lead to the false (positive) conclusion that in the biological system the compound scavenges the specific radical of interest. Polyphenolic compounds are an example of that situation. Although in chemical systems they have been shown to be potent radical scavengers, at the typical concentrations used in biological systems their effect cannot be attributed to radical scavenging. In fact, many of polyphenols are able to prevent radical formation by interfering with the pathways leading to radical production (e.g. inhibition of protein kinases, NADPH oxidase). The opposite situation occurs when the scavenger causes radical production. An example may be the formation of scavenger-derived radical, which is able to cause one-electron reduction of oxygen to O2•- via one of the pathways discussed above in the case of probe-derived radicals P•. This will lead to a false (negative) conclusion that the scavenged radicals are not involved in the probe oxidation. D. In case of biological systems, the scavenger should not be toxic at the concentration used. An obvious limitation of the scavengers is that they may alter cellular function, in the extreme case causing the cell death. For example, as discussed below, to scavenge effectively hydroxyl radical, the compound added should be present at concentration equal to total concentration of the cellular organic matter. This cannot

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be achieved without significant alteration of cellular structure and function, most probably leading to cell death. E. The radical scavenger should not interfere (chemically or physically) with the assay. The radical scavengers may lead to the decrease of the signal detected not only by scavenging the radical but also by interfering with the assay. This includes both chemical and physical interaction with the assay components. The chemical interference is typically due to the reaction of the scavenger with the probe, the product from the probe and/or with any reaction intermediates on the pathway between the probe and the product detected. For instance, the radical scavenger may be able to reduce the probe derived radical P• back to the probe, and in this way inhibit product formation. An example of physical interference is the absorption of the excitation and/or emission light by the scavenger when fluorescent product is being monitored. While radical scavengers are used to confirm the radical identity, the inhibitors of the radical production systems are also used for the establishment of the source of the radical. For example, in the case of superoxide radical anion, inhibitors of mitochondrial function, NADPH oxidase and xanthine oxidase are used both in cell culture studies and in vivo to establish sources of O2•- production. The requirements for the inhibitors are similar to the ones listed above for the radical scavengers, except point C, as inhibitors should affect (decrease) the rate of radical production. On the other hand, they should not directly scavenge the radical under study.

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3. Methods of Generation and Detection of Selected Free Radicals 3.1. General Considerations In case of most radicals, multiple ways of generation and detection of specific species are available. Some of the methods can be used in both cell-free and cellular systems, while others can be reliably used only in cell-free or cellular systems. For example, radiolytic methods of generation of specific radical can be used in well-defined chemical settings, but will yield poorly-defined mixtures of radicals in biological systems. Some photolytic methods can be used both in cell-free and cellular systems, with retention of the identity of the radical produced. On the other hand, there are methods of radical generation which are useful in biological systems but are practically useless in cell-free conditions. For instance, redox cycling compounds (e.g. paraquat) stimulate intracellular O2•- production, but alone in a pure chemical environment are not capable of producing O2•-. Similarly, some probes which are useful in chemical settings cannot be used in biological systems and vice versa. The applicability and limitations of the methods of generation and detection will be discussed separately for each radical described in this Chapter. 3.1.1. Methods of Free Radical Generation For studies on the chemical reactivity of specific radicals, development of the appropriate methods of detection, and on the effects of the radical‟s presence in biological systems,

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method(s) of controlled generation of the radical both in cell-free and cellular conditions should be available. Generally, the methods of radical generation can be divided into two groups: a) Pulse methods, in which the radical is “introduced” into the investigated system within a time period significantly shorter than the lifetime of the radical. These methods are typically used in connection with fast kinetic monitoring of radicalinduced processes. b) Steady-state methods, in which the radical is continuously generated within the system over a time range significantly exceeding the radical‟s lifetime. Those methods may be used in connection with real-time monitoring of the product(s) formed using non-destructive analytical methods, or with end-point measurement after stopping the generation of the radicals.

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3.1.1.1. Pulse Methods The most-widely used pulse methods of radical introduction into investigated system involve pulse radiolysis, laser flash photolysis and fast-mixing (e.g. stopped flow) techniques. Pulse Radiolysis Pulse radiolysis is based on the exposure of the sample to short (typically in the range of nano or microseconds) pulse of ionizing radiation, which leads to ionization of the molecules present in the investigated system [21-23]. Due to the lack of selectivity of ionizing radiation, the ionization is almost exclusively limited to the solvent molecules, at least in diluted (< 0.1 M) solutions. For example the concentration of water molecules in aqueous solution is ca. 55 M, and thus in most cases water will be the primary target for ionization in aqueous systems. The radiolysis of neutral water leads to the formation of three highly reactive species: hydrated electron eaq- (2.6), hydroxyl radical •OH (2.7) and hydrogen atom H• (0.6) in addition to the formation of less reactive products, hydrogen peroxide H2O2 (0.7), molecular hydrogen H2 (0.45) and hydronium cation H3O+ (2.6) (numbers in parentheses are the radiation yield values G defined as the number of the species formed per 100 eV energy absorbed, equation 1).

G(X) 100

number of species X formed energy absorbed (in eV)

(Equation 1)

Of the three highly reactive species formed, two (H• and eaq-) exist in acid-base equilibrium (reaction 20). H• ⇄ eaq- + H+

(20)

While the hydroxyl radical (•OH) is a highly oxidizing species, hydrated electron (eaq-) is a very strong reductant and hydrogen atom (H•) may behave as reductant or oxidant depending on the nature of co-reactant. To investigate the reactivity of the specific radical, however, only one type of radical should be generated. The oxidizing radicals are typically produced with the use of hydroxyl radical; thus the reducing eaq- should be eliminated and/or

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converted into an oxidizing radical. For the conversion of eaq- into •OH, the solution may be saturated with nitrous oxide (N2O, also known as laughing gas), which reacts with eaq- to yield • OH (reaction 21, k21 = 9.1 × 109 M-1s-1). eaq- + N2O + H2O  •OH + -OH + N2

(21)

For investigation of reducing radicals, the hydroxyl radical needs to be eliminated and/or converted into reducing radical. An example of the conversion of •OH into the reducing radical involves its reaction with 2-propanol (reaction 22, k22 = 1.9 × 109 M-1s-1), which produces the corresponding ketyl radical bearing reducing properties. (CH3)2CHOH + •OH  (CH3)2•COH + H2O

(22)

Another example involves the conversion of •OH into carbon dioxide radical anion (CO2•-) that is formed in the reaction of •OH with formate anion HCOO- (reaction 23, k23 = 3.2 × 109 M-1s-1): HCOO- + •OH  CO2•- + H2O

(23)

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More detailed strategies for the radiolytic generation of specific radicals are provided below. Laser Flash Photolysis Laser flash photolysis is based on the exposure of the sample to short (typically ranging from pico to microseconds) pulses of light (within UV or visible range) [22]. In contrast to radiation chemical methods, in the photolytic techniques the primary target of the radiation is the solute (S) rather than solvent. The basic requirement of the photolytic techniques is that the solute S is able to absorb the light used, i.e. it is characterized by the absorption band in the spectral region overlapping with the spectral characteristics of the light source. Absorption of the electromagnetic radiation (h ) in UV or visible range will lead to the change in electronic configuration of the molecule and the formation of its excited state (S)* (reaction 24), which may undergo photodecomposition into radical S• (reaction 25). The radical S• may be a radical of interest (for example, in studying •OH, the photo-dissociation of H2O2 into •OH). S + h → (S)*

(24)

(S)* → S•

(25)

In many cases, however, the primary product formed upon photolysis of S (excited state or radical photoproduct) is used in a secondary reaction to form the radical of interest R• (for example via reactions 26-28). S• + R → S + R •

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(26)

212

Jacek Zielonka and Balaraman Kalyanaraman (S)* + R → S + (R)*

(27)

(R)* → R•

(28)

Stopped Flow The stopped flow method is an example of fast-mixing techniques and is based on mixing of two solutions, one containing the radical of interest and the other the molecular target of interest. Commercial stopped flow systems are characterized by the mixing time down to 1 millisecond, and thus processes occurring over time ranges exceeding 10 milliseconds can be reliably studied. Typically, after the mixing, the reaction is monitored using spectrophotometric or fluorescence techniques. An example of the use of stopped-flow system are studies of superoxide reactivity. Of the two solutions one contains the superoxide solution in DMSO, in which solvent superoxide is stable, and the other solution contains the buffered aqueous solution of the substrate, the reactivity of which towards superoxide is being investigated. To maintain the properties of the aqueous solution, asymmetric mixing is used, i.e. during mixing the volume ratio of aqueous solution to DMSO solution should be 10 or higher. During mixing, O2•- is introduced into aqueous solution and its decay in the absence and presence of the co-reactant is kinetically examined.

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3.1.1.2. Steady-state Methods The steady-state methods are based on continuous supply of the radical to the investigated system. These methods can be divided into several sub-classes including but not limited to: a) Chemical b) Enzymatic c) Radiation-chemical and photolytic generation a) Chemical (thermal) Generation of the Radicals The chemical source may involve the compound (S), which spontaneously decomposes with the formation of the radical species S• (reaction 29, for example diazenium diolates as the sources of •NO). S → S•

(29)

The radical S• may also be the precursor of the radical of interest R• (reaction 30, for example azo-compounds as the precursors of peroxyl radicals). S• →→ R•

(30)

The radical may be continuously formed during the slow chemical reaction between reactants A and B (reaction 31, for example formation of •OH in Fenton-like reaction) A + B → R• + other products

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(31)

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While the chemical system is typically well-defined, in some cases problems may arise with the ability to stop the radical production, and the use of radical scavenger seems to be the most universal approach to terminate the reaction studied. In other cases the change of pH or scavenging of the reaction substrate (for instance the addition of catalase to Fenton‟s system) may be an alternative. b) Enzymatic Generation of the Radicals Enzymatic generation of radical R• is based on enzyme catalyzed formation of the radical of interest. For example xanthine oxidase is a well-defined enzymatic source of O2•-. Another example may be the formation of nitrogen dioxide (•NO2) via myeloperoxidase (MPO)catalyzed oxidation of NO2- by H2O2. In case of enzymatic reaction, in addition to radical and/or substrate scavenging, the use of enzyme inhibitors provides the possibility to control the amount of the radical formed and thus to terminate the studied reaction.

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c) Radiation-chemical and Photolytic Generation The radiation-chemical and photolytic methods of radical generation have been described above in the case of pulse methods. The major difference is that the sample is exposed to continuous radiation and thus there is no need to use sophisticated pulse generating instruments. In fact, the steady-state radiolytic techniques described in this Chapter can be conveniently carried out in many medical centers, by application of the accelerators used for cancer radiotherapy or X-ray generating equipment. Both radiolytic and photolytic methods of radical generation have the advantage that the amount of radical generated can be easily controlled by manipulating the time of exposure and the intensity (dose rate) of the radiation used. 3.1.2. Principles of the Analytical Methods of Radical Detection Due the presence of an unpaired electron, and the open-shell electronic character, radicals can be directly detected by EPR as well as in most cases by UV-Vis absorption spectroscopy. These methods are, however, often not very sensitive, and thus significant accumulation of the radical species is required for direct detection. On the other hand, the probes for radicals are designed to provide easily detectable product with low detection limit, down to nanomolar radical concentrations. As fluorescence spectroscopy offers both high specificity and sensitivity, most probes have been designed to yield fluorescent product. As discussed above, the quantitative analysis of the probe concentration as well as of the product(s) of its reaction with the radical require separation of the reaction mixture and separate quantification of each analyte. This requirement is typically fulfilled by the use of chromatographic techniques and HPLC systems equipped with various detector types. The basic principles for each of those detection methods (EPR, UV-Vis absorption and fluorescence spectroscopy, HPLC) are described below. Electron Paramagnetic Resonance (EPR) EPR is a technique which detects molecules carrying unpaired electrons and is widely used in the free radical field [24]. The unpaired electron may be in two spin states, represented by spin quantum numbers: +½ or -½. The difference in the energy ( E = E+½ - E-½) between both spin states is described by equation 2.

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Figure 1. Splitting of the spin energy states and resonance conditions as a function of the magnetic field strength B. Lower lines represent the EPR spectra (absorption and first derivative) corresponding to the electron spin transition at the microwave radiation energy equal to h .

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E = ge

BB

(Equation 2)

where ge is the g-factor of the electron which is equal to 2.0023 in the case of free electron, -24 J T-1, and B is the strength of the magnetic field. B is Bohr magneton equal to 9.274 × 10 This equation implies that in the absence of the magnetic field, both spin states have the same energy, and that with increasing magnetic field strength, the E value increases (Figure 1). The transition between both spin states is possible only if the system absorbs or emits electromagnetic radiation of energy h equal to the value of E (equation 3) h = ge

BB

(Equation 3)

Equation 3, in which h is the Planck constant equal to 6.626 × 10-34 J s and is the frequency of the electromagnetic radiation, is called the resonance condition and defines the relationship between the magnetic field strength and electromagnetic radiation frequency for resonance to occur. When the resonance occurs, the sample absorbs the electromagnetic radiation resulting in the appearance of EPR signal. To fulfill the resonance condition, one can vary the radiation frequency , magnetic field strength B, or both. For technical reasons, the frequency is typically kept constant and the magnetic field strength is varied to obtain EPR spectra (Figure 1). Moreover, in practice the magnetic field is constantly modulated (sinusoidally), yielding the EPR signal as the first derivative of the absorption spectrum (Figure 1). As the resonance frequency depends on the field strength B, it cannot be used as the specific parameter of the radical studied. Indeed, it is the parameter ge (equal to BB/h ) that is used for identification of the radical species. However, in most organic radicals, the value of ge of the unpaired electron is close to that of free electron (ca. 2.0) and the knowledge of its

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value may not be sufficient for radical identification. Fortunately, a number of atomic nuclei (including hydrogen and nitrogen) possess magnetic moment and therefore will modify the effective magnetic field strength B at the unpaired electron. This interaction between the unpaired electron and the atomic nuclei of non-zero spin present in proximity of that electron is called the hyperfine interaction and leads to the splitting of the single EPR band into multiple lines, giving the so-called hyperfine structure. In case of the presence of several nuclei of non-zero spin close to the unpaired electron, the spectral pattern may be complex. The structural information provided by the hyperfine structure of the EPR spectrum is used for the identification of the detected radical. Examples of the influence of the substituent and the geometry of the adduct on the spectral pattern of selected nitroxides are shown in Figure 2.

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Figure 2. Hyperfine interactions in selected nitroxides and their effects on the observed EPR spectra; aN and aH represent the hyperfine splitting constants of nitrogen and hydrogen atoms, respectively.

UV-Vis Absorption Spectroscopy UV-Vis absorption spectroscopy (spectrophotometry) is based on the absorption of the electromagnetic radiation in the UV and visible range by the molecule of interest [25]. The absorption of the radiation (light) is accompanied by transition of the molecule from the ground (S0) to the excited electronic state (Sn, n > 0, Figure 3). When light passes through the sample, its intensity decreases from I0 to I, and the ratio of I/I0 is called the transmittance T. The most-often used parameter describing the extent of light absorption by the sample is called absorbance (A) and is defined by equation 4. I0 (Equation 4) I The absorption spectrum of the molecule typically shows the dependence of the absorbance vs. the radiation wavelength, and in many cases is specific for the given species. However, at the current stage, the structural information “encoded” in the electronic spectrum is in most cases too difficult to be extracted, and therefore the UV-Vis absorption spectrum is rarely used alone for structural identification, unless the spectrum was well characterized in prior experiments and is very characteristic of the species investigated. Thanks to the linear dependence of the absorbance on the concentration of the absorbing species (equation 4, A

log

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known as Lambert-Beer law), spectrophotometry is a frequently-used method for the determination of the concentration of known species in solution. A=

cl

(Equation 5)

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The equation 5, in which is the molar extinction coefficient, c is the concentration of the absorbing species and l is the optical pathlength of the sample, is typically valid in modern instruments at least up to the absorbance value of 2, if no chemical or physical interactions between the solute molecules occur at high concentration. The value of of the product observed defines the sensitivity of the detection; the higher the molar extinction coefficient, the lower the detection limit.

Figure 3. Schematic representation of the transition from the ground state S0 to electronically excited states (transitions a-c) during absorption of light (panel A) and the corresponding absorption spectrum with absorption bands reflecting the transition of molecules between different electronic states (panel B).

Fluorescence Spectroscopy Fluorescence spectroscopy relies on two processes mediating the transitions of the molecule between the electronic ground and excited states [26]. The primary event is the same as in UV-Vis absorption spectroscopy and is based on the transition of the molecule from the ground to the excited electronic state upon absorption of light (Figure 4). The second event is the return of the molecule to the ground state, which may occur by radiative and/or non-radiative pathway(s). Fluorescence occurs when the molecule in the singlet excited state returns to the singlet ground state via radiative decay, i.e. when the transition is accompanied by the emission of light.

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Figure 4. (A) Schematic representation of the transitions between electronically and vibrationally excited states during excitation of the molecule and its relaxation via non-radiative (internal conversion) and radiative (fluorescence) transitions. Transitions a, b and c represent excitation of the molecule (absorption of light) and transition d represents radiative (fluorescent) decay of the excited state. (B) Fluorescence excitation and emission spectra corresponding to the transitions a, b, c and d. Note that the excitation spectrum typically resembles the absorption spectrum of the molecule (Figure 3).

The efficiency of the radiative (fluorescent) decay of the excited state is called fluorescence quantum yield ( ) and is defined by equation 6.

number of photons emitted number of photons absorbed

(Equation 6)

There are two types of fluorescence spectra: excitation spectrum and emission spectrum. Excitation spectrum is obtained when the fluorescence intensity in monitored at set wavelength under the conditions where the wavelength of the excitation light is varied. It therefore reflects the events of light absorption and typically the excitation spectrum closely resembles the UV-Vis absorption spectrum of the molecule. Emission spectra are collected by varying the wavelengths at which the emission light is detected, at set wavelengths of the excitation light. For quantitative analysis, the excitation and emission wavelengths are typically set at the maxima of the corresponding fluorescence bands to achieve good

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sensitivity and selectivity. The sensitivity of the fluorescence detection is proportional to the amount of light emitted, and thus to both the efficiency of light absorption and emission. The parameter used for the comparison of different fluorescent compounds is the brightness, defined as the product of molar extinction coefficient and fluorescence quantum yield ( × , M-1cm-1). High Performance Liquid Chromatography (HPLC) High performance liquid chromatography (HPLC, also called high pressure liquid chromatography) is the most widely used technique for separation and selective detection of individual components in mixtures of compounds [27]. The typical HPLC system consists of pumps which deliver the mobile phase (the solution used to elute the compounds from the column), the autosampler with the injector (used to load the sample onto the column), the column on which the individual compounds are separated, and the detector(s) used for quantification of the compounds of interest. The type of mobile phase and column can be tailored for individual projects besting order to achieve optimal separation and low organic solvent consumption. The separation is based on differences in the strength of interaction of the separated compounds with the column filling, which results in varying retention times during constant flow of the mobile phase through the column. The most commonly used separation method is reversed phase separation utilizing columns containing hydrophobic filling. In that case the strength of interaction of the analytes with the column is proportional to their hydrophobicity; the more hydrophilic compounds generally elute earlier than the hydrophobic ones. In the other method based on normal phase separation, the hydrophobic compounds elute earlier followed by the more hydrophilic analytes. Two different types of elution may be employed: isocratic and gradient. In isocratic elution, the mobile phase during sample elution is kept the same, while in gradient elution the mobile phase composition is modified during the elution process. The isocratic method is used for separation of compounds exhibiting only minor differences in the strength of interaction with the column, while the gradient method facilitates the elution of compounds characterized by major differences in interaction strength with the column filling. Different methods of detection can be used in connection with HPLC systems, with the UV-Vis absorption detector being most widely used. Other popular, HPLC-compatible detection methods utilize fluorescence, mass spectrometry and electrochemical detectors.

3.2. Superoxide Radical Anion (O2•-) 3.2.1. Physicochemical Properties Superoxide radical anion (O2•-) is a product of one-electron reduction of molecular oxygen (see also Chapter 2). In water it exists in acid-base equilibrium with hydroperoxyl radical (HO2•, pKa = 4.8, reaction 32) [28]. HO2• ⇄ O2•- + H+

(32)

Superoxide radical anion can act as an oxidant (E(O2•-,H+/HO2-) = 1.03 V) or as reductant (E(O2/O2•-) = -0.33 V) while hydroperoxyl radical is a strong oxidizing species

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(E(HO2•,H+/H2O2) = 1.45 V). In aqueous solution superoxide radical anion is unstable and undergoes dismutation via reactions 33 and 34: O2•- + HO2• → O2 + HO2-

(33)

HO2• + HO2• → O2 + H2O2

(34)

with rate constant k33 = 9.7 107 M-1s-1 and k34 = 8.3 105 M-1s-1. Therefore even in the absence of superoxide scavengers, in aqueous solution at neutral pH its lifetime is within the millisecond time scale and its decay rate is accelerated at lower pH values and higher steadystate concentrations. Aprotic solvents (DMSO, MeCN) do not promote dismutation of superoxide radical anion and therefore have been used as the convenient medium for superoxide storage. 3.2.2. Generation

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3.2.2.1. Cell-free Systems For cell-free systems, there are various methods of O2•- supply, including delivery of O2•solution in aprotic solvents or in situ O2•- generation by thermal decomposition of “superoxide donors”, as well as by electrochemical, photochemical, radiolytic and enzymatic methods. Solutions of O2•- in Aprotic Solvents As mentioned above, the superoxide radical anion is relatively stable in aprotic solvents and thus can be used for superoxide delivery. For that purpose the salt potassium superoxide (KO2) is dissolved in DMSO [29]. However, due to limited solubility of KO2 in DMSO, crown ether is dissolved in DMSO prior to addition of KO2 powder in order to increase KO2 solubility by complexing K+ cations. Thermal Sources of Superoxide The thermal sources of superoxide radical anion involve compounds which upon decomposition react with oxygen to generate O2•-. Thus, these methods require the presence of oxygen and cause its consumption in the investigated system. While many compounds may undergo “autooxidation” with the formation of O2•- (e.g. pyrogallol), they typically also react with O2•-; thus the flux of superoxide is poorly defined in those systems. The other approach is to use a precursor of the radical with reducing properties, which upon reaction with oxygen will yield O2•-. The diazo compound, which spontaneously decomposes to give hydroxybenzyl ketyl radical via intermediate benzoxy radical (reactions 35 and 36), is a well-defined superoxide thermal source (SOTS) [30]. The radical formed reacts with oxygen, generating superoxide and benzaldehyde (reaction 37).

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N N

O

O

+

2 COOH

N2

COOH

COOH

(35)

SOTS-1

O

H

COOH

H

OH

(36)

COOH

OH

H +

O +

O2

COOH

O2•- + H+

COOH

(37)

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Radiolytic Generation of Superoxide The production of superoxide by radiolysis of oxygenated aqueous solutions is regarded as one of the best-defined and controlled methods used for quantitative analysis of superoxide reactions [31]. Superoxide and its protonated form are formed by direct reaction of oxygen with hydrated electron (reaction 38) and hydrogen atom (reaction 39), respectively. eaq- + O2 → O2•-

(38)

H• + O2 → HO2•

(39)

At a specific pH one can obtain the desired ratio O2•-/HO2• due to fast acid-base equilibration (reaction 32). This method, however, requires the addition of an hydroxyl radical scavenger, typically tert-butanol, which produces relatively low-reactive carboncentered radicals in reaction with •OH radical. The other possibility is to add sodium formate to covert •OH into CO2•-, which upon reaction with O2 produces additional amounts of O2•(reaction 40). CO2•- + O2 → CO2 + O2•-

(40)

Additional pathways for radiolytic generation of O2•-/HO2• involve hydrogen abstraction from H2O2 (reaction 41) and production of -hydroxyalkyl radicals, which upon reaction with O2 form O2•- and/or HO2• (reactions 42-43). •

OH + H2O2 → H2O + HO2•

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species … •

221

OH + RHOH → H2O + R•(OH)

(42)

R•(OH) + O2 → R(OH)OO• → R=O + HO2• (O2•- + H+)

(43)

Enzymatic Generation of Superoxide A commonly used source of superoxide is xanthine oxidase (XO)-catalyzed oxidation of hypoxanthine by molecular oxygen to uric acid (U) via intermediate formation of xanthine (X) (reactions 44 and 45) [32]. HX + O2 → X, O2•-, H2O2

(44)

X + O2 → U, O2•-, H2O2

(45)

Both reactions yield superoxide radical anion and hydrogen peroxide and both HX and X are substrates for XO-catalyzed production of O2•-.

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3.2.2.2. Cellular Systems Due to low cell membrane permeability of O2•- and its rapid dismutation, the increase in intracellular O2•- level is typically achieved by stimulation of intracellular production, rather than incubation of the cells with extracellular sources of O2•-. Different strategies have been applied to stimulate O2•- production, with redox-cycling agents (menadione, paraquat) most frequently employed for that purpose. Other strategies involve stimulation of endogenous O2•-generating systems, including the electron transfer chain in mitochondria, and activation of NADPH oxidase (e.g. by phorbol myristate acetate, PMA). 3.2.3. Detection Due to the short lifetime of O2•- at and below neutral pH, direct monitoring of its reactions is possible only under alkaline conditions (pH > 12) using steady-state techniques; at pH < 12 pulse radiolysis, a time-resolved method, is the most widely used. Superoxide radical anion has an absorption band in UV range with maximum at 245 nm (with the extinction coefficient of 2350 M-1cm-1), which can be used to follow its formation during the reaction. For the detection of low concentrations of O2•- and/or its formation on longer time-scales, O2•--reactive probes have to be used. While there are many probes available for superoxide detection, the two most-widely used (reduction of ferricytochrome c and EPR spin-trapping) are described below. In case of all probes the use of SOD is recommended to test whether, and to what extent, the signal observed is due to the reaction with O2•-. Ferricytochrome C Assay In cell-free systems the detection of superoxide is accomplished by the use of ferricytochrome c (cyt c(Fe3+)) which is reduced by O2•- with the formation of ferrocytochrome c (cyt c(Fe2+), reaction 46) [33]. cyt c(Fe3+) + O2•- → cyt c(Fe2+) + O2

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Jacek Zielonka and Balaraman Kalyanaraman

The reaction is relatively fast (k46 ~ 5 × 105 M-1s-1) and the progress of reduction can be monitored by UV-Vis spectrophotometry by following the build-up of absorbance at 550 nm (the difference in the extinction coefficients between reduced and oxidized forms of cytochrome c is reported within the range (1.8 – 2.1) × 104 M-1cm-1). The cytochrome c-based assay cannot be used in the presence of cyt c – reactive compounds (e.g. reduced glutathione, ascorbate) and enzymes capable of direct reduction of cyt c. To inhibit the enzymatic reduction of cyt c, one can use the acetylated analog of the protein which has lower affinity towards the active sites of the enzymes. The re-oxidation of reduced cyt c may be another confounding factor and use of catalase to scavenge H2O2 is recommended. EPR Spin-trapping Another widely used method for detection of O2•- is based on its reaction with cyclic nitrones, with the formation of secondary radicals (nitroxides) of higher stability and with very characteristic EPR spectra (reactions 47 and 48) [34].

H3C H3C

H

N O

+ O2 + H

H

DEPMPO

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N O

OOH H

H3C H3C

DMPO-•OOH

DMPO

C2H5O O C2H5O P H3C N O

H3C H3C

+ O2 + H

C2H5O O C2H5O P H3C N O

N O

OH H

DMPO-•OH

(47)

OOH H

DEPMPO-•OOH

(48)

Although these methods require access to an EPR instrument, the advantage of detection of specific product makes it relatively popular. Because the rate constant of the reaction of O2•- with nitrones is rather low (k < 100 s-1 for most spin traps), high concentration (> 20 mM) are typically used. The rate of the spontaneous decay of the O2•- adduct depends on the spin trap structure and in some cases the secondary product is identical with the hydroxyl radical adduct to the spin trap (e.g. DMPO spin trap, reaction 47). The adduct of O 2•- to DEPMPO spin trap is more stable and does not undergo spontaneous conversion to hydroxyl radical adduct, and thus use of DEPMPO spin trap for O2•- detection (reaction 48) is recommended. Measurement of Intracellular O2•- Production The above mentioned methods are applicable to cell-free or extracellular O2•measurements due to low cell membrane permeability of those probes. There is an urgent need for new superoxide-specific probes that would allow for reliable quantification of intracellular O2•- flux in order to ascertain its contribution to physiological and pathophysiological processes.

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Hydroethidine Assay Currently, the fluorogenic probe hydroethidine is widely used for O2•- detection [35]. The probe reacts with O2•- to form 2-hydroxyethidium (reaction 49), which is considered highly specific for O2•- as it is not formed with other biologically relevant oxidants. Hydroethidine is not, however, selective for O2•-, as other oxidants can convert it into ethidium cation and/or dimeric products. As both 2-hydroxyethidium and ethidium emit red fluorescence, the fluorescence-based techniques of 2-hydroxyethidium in biological samples have been called into question. In fact, the HPLC-based assay has been praised as most accurate, in so far as it allows separation of HE and its oxidation products and selective monitoring of their intracellular levels. Thus, one can monitor probe uptake and the profile of its oxidation products, including superoxide-specific 2-hydroxyethidium. The conversion of hydroethidine into 2-hydroxyethidium is a multistep process involving radical of HE (reaction 49). H2N

+

H N C 2 H5 H2 N

NH2 H 2N

HE•+ H N C2H5

NH2

O2 (49a)

+H

+

O2 (49b)

-H+

H2 N

OH N C2 H5

NH2

2-OH-E+ HE

H N C2 H 5

NH

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HE(•NH)

(49) As the first step, one-electron oxidation of HE (step a) can be accomplished by many oxidants, the specificity of the product (2-OH-E+) towards O2•- is attributed to the consecutive step, the reaction of HE-derived radical with O2•- (step b). This implies that the yield of 2OH-E+ in cellular systems will be proportional to the steady state concentrations of both O2•and HE radical. As steady-state concentration of HE radical will be a function of HE concentration and the rate of its one-electron oxidation, knowledge of intracellular levels of HE and the non-specific products of HE oxidation (ethidium cation and dimeric products) is crucial for proper interpretation of the changes in the levels of 2-OH-E+. Aconitase Inactivation Assay Complementary to the use of the externally added probes for O2•-, the superoxide-specific effects on the endogenous enzymes may be studied as well. The most commonly used is the assay of the activity of aconitase, as this enzyme undergoes inactivation by O2•- [36]. Aconitase catalyses the interconversion of citrate and isocitrate via a cis-aconitate intermediate. The rate constant of the reaction of aconitase with O2•- is relatively high (k ~ 106 - 107 M-1s-1). During the reaction between aconitase and O2•-, iron is released from its iron-

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Jacek Zielonka and Balaraman Kalyanaraman

sulfur cluster, leading to enzyme inactivation. The advantage of the assay is the possibility to distinguish between intramitochondrial and extramitochondrial O2•- production by measurement of aconitase activity in subcellular mitochondrial and cytosolic fractions, respectively. Because the enzyme inactivation is reversible, the conclusions about the increase in O2•- formation should be made with caution. Aconitase may be also inactivated by peroxynitrite (ONOO-) and therefore the assay should not be regarded as totally specific towards O2•-. Other Assays for Intracellular Superoxide Other methods for detection of intracellular O2•- include the chemiluminescent assay using lucigenin and luminol, and EPR-based detection of hydroxylamine oxidation. As discussed later in this Chapter, lucigenin and luminol may increase the rate of O2•- formation, and thus their use should be avoided. The oxidation of EPR-silent hydroxylamines into EPRactive nitroxide (reaction 50) has been proposed for O2•- detection but is not very specific. COOH

COOH

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+

O2

+ H

+

N OH

N O

CPH

CP

H2O2

(50)

In addition, the nitroxide formed may act as an SOD-mimetic (see Vol. II, Chapter 6) and thus may decrease the steady-state concentration of O2•- leading to underestimation of its actual intracellular concentration. Nitroxide is also prone to undergoing conversion back to hydroxylamine by cellular reductants, again causing underestimation of O2•- levels. As O2•- is converted into H2O2 in non-enzymatic and SOD-catalyzed processes, the detection of H2O2 released to the cell culture medium has also been used to monitor intracellular O2•- production. Those assays cannot, however, discriminate between intracellular O2•- and H2O2 formation. Moreover, the O2•- - independent changes in intracellular H2O2 concentration (for example due to changes in H2O2-scavenging systems) may lead to false conclusions concerning changes in O2•- levels. 3.2.4. Inhibitors of O2•- Production and O2•- Scavengers In case of cell-free or extracellular experiments, the specificity of the signal detected for •O2 can be tested by addition of SOD, which is highly specific and a fast catalytic scavenger of O2•-. SOD is not cell membrane-permeable, and thus it cannot be used to validate the assays of intracellular O2•-. Several strategies have been applied to inhibit intracellular O2•-, including increasing membrane-permeability of SOD by linking the enzyme with polyethylene glycol. The other approaches include the application of cell-permeable SODmimetic compounds, e.g. cyclic nitroxides and metal complexes with porphyrins (see Vol. II, Chapter 6). However, the specificity of those compounds towards O2•- is not always clear and most superoxide-scavenging metal-porphyrin complexes can also scavenge ONOO-. Of note, MnTBAP, a manganese-porphyrin complex widely used as SOD mimetic has been reported to lack superoxide-scavenging activity [37] .

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3.3. Nitric Oxide (•NO) 3.3.1. Physicochemical Properties Nitric oxide (•NO) is a radical molecule of high stability in aqueous solution in the absence of co-reactants (e.g. molecular oxygen) [38, 39] (see also Chapter 3). •NO can undergo one-electron reduction to nitroxyl anion, NO- (E(•NO/NO-) = -0.8 V), or one-electron oxidation with the formation of nitrosonium cation, NO+ (E(NO+/•NO) = 1.2 V). In the presence of oxygen •NO undergoes oxidation with the formation of NO2- (reactions 51-53) via intermediate formation of nitrogen dioxide (•NO2) and dinitrogen trioxide (N2O3). 2 •NO + O2 → 2 •NO2 •

(51)

NO2 + •NO → N2O3

(52)

N2O3 + H2O → 2 HNO2

(53)

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3.3.2. Generation 3.3.2.1. Cell-free Systems When handling •NO gas and its aqueous solution, one has to bear in mind its spontaneous oxidation by molecular oxygen [40]. Thus, all solutions used to purify and store • NO should be deoxygenated, for example by purging with pure argon. Although nitric oxide is a relatively stable gaseous compound, its storage in gas tanks may lead to its decomposition, as under high pressure it undergoes disproportionation to give nitrous oxide (N2O) and nitrogen dioxide (•NO2). Thus, •NO needs to be purified by passing it through a solution of sodium hydroxide (NaOH, 1 M) followed by the solution of the buffer used. The alternative to the use of •NO gas tank is to generate it in the laboratory. •NO can be prepared by decomposition of nitrous acid (reaction 54) 3 HNO2 → 2 •NO + HNO3 + H2O

(54)

Again, the gas evolved should be purified, as described above. The alternative to use of gaseous •NO or deoxygenated •NO solutions is to employ compounds which generate •NO in situ, under well controlled conditions [41]. Such compounds, termed •NO-donors, are simple to use and may mimic physiological conditions of continuous flux of •NO. Moreover, the wide range of •NO-donors allows ascertainment whether the effects observed are due to •NO formed or to •NO-donor per se or its decomposition by-product(s). Additionally, different • NO donors display various half-lives under physiological conditions, and thus one can control both the flux of •NO and the duration of •NO exposure. The most often used •NOdonors include S-nitrosothiols (thionitrites) and diazeniumdiolates (NONOates). The use of diazeniumdiolates is recommended, as the mechanism of •NO release is well established and the reaction lifetime ranges from minutes to hours [42]. While theoretically one molecule of the donor should generate two molecules of •NO (reaction 55), the reported stoichiometry depends on the actual structure of the donor and should be determined under the experimental conditions used [42, 43].

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Jacek Zielonka and Balaraman Kalyanaraman R1 R2

O N N N O

+

H

R1 NH

+

2 •NO

R2

NONOate

(55)

An obvious way to mimic intracellular •NO production is use of nitric oxide synthase (NOS). While possible, this is often impractical due to instability of the enzyme and the need for several cofactors for enzymatic •NO generation. In order to introduce •NO into solution rapidly (on the sub-millisecond time scale), pulse methods including flash photolysis and pulse radiolysis can be used. In the photolytic method, the excitation of nitrosothiols or metal-•NO complex releases •NO (reaction 56). RSNO + h → RS• + •NO

(56)

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In the radiolytic method •NO is generated by one-electron reduction of nitrite anion (reaction 57) followed by hydrolysis of the radical dianion formed (reaction 58). NO2- + eaq → NO2•2-

(57)

NO2•2- + H2O → •NO + 2HO-

(58)

3.3.2.2. Biological Systems In order to increase intracellular •NO levels, two general approaches are used: (1) the incubation of cells with •NO donors and (2) stimulation of endogenous production of •NO. The first approach is based on the reactions described above for generation of •NO in cell-free systems or by administration of compounds which are metabolized intracellularly to release • NO (e.g. organic nitrites). The second approach, stimulation of endogenous •NO production, is based on the induction of NOS expression and/or activation of the enzyme by increasing intracellular concentration of the NOS cofactors or induction of enzyme phosphorylation. In cells capable of expressing inducible isoform of NOS (iNOS), the addition of the corresponding stimuli (e.g. cytokines) leads to increases in the rate of •NO production. In the case of constitutively expressed NOS isoforms, augmentation of intracellular calcium concentration by calcium ionophores (e.g. ionomycin, A23187) has been reported to increase intracellular steady-state •NO levels. 3.3.3. Detection 3.3.3.1. Cell-free Systems Electrochemical Detection Direct monitoring of •NO is usually achieved using electrochemical methods [44]. •NOselective electrodes have been developed to monitor •NO levels in cell free systems as well as • NO released from cells to the extracellular medium. As the signal is proportional to the actual concentration of •NO at any given time, calculation of total •NO produced should be performed by integration of the electrode response over time. The calibration of the probe

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with pure deoxygenated solutions of •NO in the medium used in the actual experiment is crucial for quantitative analysis. Chemiluminescence Detection Another approach to monitoring •NO is chemiluminescent detection involving reaction of • NO gas with ozone to produce molecular oxygen and nitrogen dioxide in the excited state (•NO2)* (reaction 59) [40]. The spontaneous transition from excited to ground state of NO2 is accompanied by emission of light (photons of energy h , reaction 60), detected by the photomultiplier. •

NO + O3 → (•NO2)* + O2

(59)

(•NO2)* → •NO2 + h

(60)

In order to transfer •NO from the liquid phase of aqueous solution to the gas phase of the chemiluminescence reaction chamber, the chamber is connected to the reaction vessel with air-tight glassware and the reaction solution is continuously purged with argon gas. Therefore, this detection method is limited to •NO generating systems under anaerobic conditions and the reaction mixture is constantly modified by removing gaseous and volatile components. To avoid those limitations, the same system may be used to detect nitrite, the stable oxidation product of •NO. In that case, after specific incubation time, the sample is analyzed for conversion (reduction) of accumulated nitrite anions to •NO by potassium iodide (KI) under acidic conditions (reaction 61).

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I- + NO2- + 2 H+ → •NO + ½ I2 + H2O

(61)

The •NO formed is transferred into a chemiluminescence reaction chamber, as described above. Oxyhemoglobin Assay • NO detection may also be accomplished by spectrophotometric monitoring of the conversion of oxyhemoglobin (or oxymyoglobin) to methemoglobin (or metmyoglobin) by • NO (reaction 62) [40, 45]. Hb(Fe2+)O2 + •NO → Hb(Fe3+) + NO3-

(62)

The conversion can be conveniently monitored by following the decrease in absorbance at 420 nm and 577 nm with concomitant increase in absorbance at 401 nm. For low concentrations (fluxes) of •NO, the changes at 420 nm and 401 nm may be used ( 420 nm = -5.1 × 104 M-1cm-1, 401 nm = 4.9 × 104 M-1cm-1) after subtraction of the absorbance read at the isosbestic point at 410.5 nm, while for higher concentration of •NO, less sensitive absorption changes at 577 nm ( 577 nm = -1.03× 104 M-1cm-1) may be utilized. The assay is relatively simple and allows for real-time monitoring of •NO fluxes. In contrast to the electrochemical method, the response in the oxyhemoglobin assay is cumulative, i.e. the detector response reflects the total amount of •NO produced from the start of measurement.

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The drawback of the method is the possibility of auto-oxidation of oxyhemoglobin, and therefore the results should be corrected for that process. Assays Based on Fluorescent Aromatic Triazole Formation Another approach to monitor in situ •NO generation is to use aromatic diamino compounds, which, in the presence of both •NO and O2 undergo conversion to fluorescent aromatic triazoles [46]. The most commonly used probes include 2,3-diaminonaphthalene (DAN) and 4,5-diaminofluorescein (DAF-2) (reactions 63 and 64). NH2

H N N N

•NO/O 2

NH2 DAN

(63) NH2

N NH

H2N

N COO

O

O

COO

•NO/O 2

O

O

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DAF-2

O

O

DAF-2T

(64)

Although the mechanism of this reaction is not completely understood, it is assumed that N2O3 is the actual nitrosating species (reaction 65, step a), formed in the reaction of •NO with oxygen. Another possibility is that nitrogen dioxide (•NO2) oxidizes the diamino compounds with the formation of aminyl radical (reaction 65, step b) which reacts with •NO forming Nnitroso derivative (step c), yielding triazole derivative, DAF-2T with the elimination of water (step d). O NH2

N HN

H2N COO

NH2

N NH N COO

N2O3 (65a)

O

O

COO

- H2O (65d)

O

O

O

O

DAF-2

O

O

O

DAF-2T

one-electron oxidation (65b) e.g. NO2

NO (65c)

NH2 HN COO

O

O

O

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As in the case of the oxyhemoglobin assay, the observed signal reflects total amount of NO produced due to accumulation of the nitrosated product. Due to the indirect nature of the detection, the assay is limited to solutions containing oxygen. Also, as the reaction involves several steps, the rate of product formation will be determined by the rate of the slowest step which may not always represent •NO formation. The conversion of aromatic diamino compounds into triazoles may also be achieved by reaction with nitrite under acidic conditions (reaction 66) and this reaction is used to measure the total amount of nitrite produced in the system. •

O NH2

N HN

H2N COO

O

O

NH2

N NH N COO

NO2-, H+

COO

- H2O

O

O

O

O

O

O

DAF-2

O

DAF-2T

(66)

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The assay involves acidification of the medium after specific time of incubation and addition of probe. Before fluorescence measurement, the pH of the solution is brought back to ca. 8 with sodium hydroxide to increase the fluorescence intensity of the triazole formed. The major drawback of the assay is its end-point character, i.e. it cannot be used for real-time monitoring of NO2- formation. EPR Spin-trapping Another method of detecting •NO is use of spin traps to transform short-lived •NO radical into relatively stable spin adducts, the formation of which may be monitored by EPR spectrometry [13, 14]. The most widely used •NO spin traps are iron dithiocarbamates, e.g. Fe(DETC)2 and Fe(MGD)2 (reaction 67). R1

S N

R2

S

Fe2+

S

R1 N

S

DETC:

R1, R2 = -C2H5

MGD:

R1 = -CH3, R2 =

+

R1

•

R2

S N

NO R2

S

NO Fe2+

S

R1 N

S

R2

OH OH OH OH OH

(67)

The other class of compounds available for EPR monitoring of •NO are nitronyl nitroxides (e.g. PTIOs) which react with •NO by oxygen transfer yielding imino nitroxides (PTIs) and •NO2 (reaction 68).

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Jacek Zielonka and Balaraman Kalyanaraman O N R

+

N

•NO

R

N O

+

•NO 2

N O

PTIO: R =

carboxy-PTIO: R =

COO-

(68)

As PTIO exhibits a different EPR spectrum than PTI, both the decay of PTIO and formation of PTI can be monitored. Major drawbacks of this assay include the generation of another radical during the reaction (•NO2) and potential reduction of PTIO and PTI to EPRsilent hydroxylamines by reductants present in the investigated system.

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Griess Assay for Nitrite Anion The accumulation of nitrite anion as a measure of •NO formed may be followed using the Griess assay [47]. The assay is based on the diazotization of sulfanilic acid or sulfanilamide by nitrite under acidic conditions, followed by coupling of the diazonium cation with Nnaphthylethylene diamine to form an azo compound with strong absorption at 548 nm. The assay is relatively straightforward and can be used with plate readers capable of absorption measurements. 3.3.3.2. Biological Systems Essentially, all methods described above for cell-free detection of •NO may also be applied to biological systems. However, for the detection of intracellular •NO, cell-permeable probes are required. A cell-permeable derivative of DAF-2 is currently the most-widely used probe for monitoring intracellular •NO formation [48]. The diacetate derivative of DAF-2, DAF-2DA undergoes hydrolysis by intracellular esterases (reaction 69) and the actual probe, DAF-2, remains trapped inside cells due to its anionic character. DAF-2DA → DAF-2 + 2 CH3COOH

(69)

It has been argued that the indirect manner of detection (detection of •NO/O2 reaction product rather than •NO itself) may be an advantage, as the probe should not interfere with • NO signaling. Also, iron dithiocarbamates have been used for intracellular monitoring of • NO production by EPR spectrometry. The availability of both lipophilic (e.g. Fe(DETC)2) and hydrophilic (e.g. Fe(MGD)2) •NO-spin trapping agents allows for controlled accumulation of the probes in specific compartments. The advantage of iron dithiocarbamates is the possibility of monitoring •NO production in vivo by EPR L-band spectrometry or by magnetic resonance imaging (MRI) techniques. EPR spectroscopy can be also employed for monitoring nitrosohemoglobin, formed endogenously in vivo in the reaction of ferrous hemoglobin with •NO (reaction 70). Hb(Fe2+) + •NO → HbNO

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species …

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For that purpose, samples of plasma are taken from live animals and analyzed by lowtemperature EPR spectroscopy. Although electrochemical detection is currently limited to extracellular •NO detection, the high sensitivity and the possibility of miniaturization of the electrochemical sensors allow for monitoring of in vivo as well as single-cell •NO production. Akin to cell-free assays, another method to determine •NO production in biological systems is measurement of stable •NO-derived oxidation products. While in simple aqueous solution •NO is oxidized exclusively to nitrite anion, in biological systems both nitrite and nitrate anions are formed. Therefore the amount of both anions should be determined to account for total •NO production. In the case of aromatic diamino compounds and the Griess reagent, nitrate must be converted to nitrite; with chemiluminescence detection, nitrate must be converted into nitric oxide. A standard approach to convert NO3- into NO2- is based on enzymatic reduction utilizing nitrate reductase and NAD(P)H coenzyme as the reducing agent (reaction 71). NO3- + NAD(P)H → NO2- + NAD(P)+ + -OH

(71)

For the conversion of NO3- into •NO, the sample is mixed with vanadium chloride (VCl3) in hydrochloric acid and incubated at 90 oC (reaction 72). 2NO3- + 3V3+ + 2H2O → 2•NO + 3 VO2+ + 4H+

(72)

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Under those conditions nitrite anions and nitrosothiols will produce •NO, and therefore this method yields the sum of all three species: NO3-, NO2- and RSNO. 3.3.4. Inhibitors and Scavengers The most commonly used scavenger of •NO is PTIO or its water-soluble derivative, carboxy-PTIO. As the reaction with PTIO converts •NO to •NO2, PTIO will not suppress the nitrosation of aromatic diamino compounds or nitrite accumulation. In biological systems, where •NO is produced almost exclusively by NOS enzymes, specific inhibitors of the latter may be used to confirm the involvement of •NO in the measured signal. Standard inhibitors include analogs of L-arginine, such as N -nitro and N -methyl derivatives. As nitrocompounds slowly release •NO under conditions of conversion of NO3- into •NO (vanadium chloride method), nitro-derivatives of arginine (L-N-nitroarginine, L-NNA, and its methyl ester, L-NAME) should not be used in connection with the chemiluminescence assay for NO3. 3.4. Nitrogen Dioxide (•NO2) 3.4.1. Physicochemical Properties Nitrogen dioxide radical is the product of one-electron oxidation of nitrite anion (NO2-). It exists in equilibrium with N2O4 (reaction 73) [49-51]. 2 •NO2 ⇄ N2O4

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(73)

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Jacek Zielonka and Balaraman Kalyanaraman

Both •NO2 and N2O4 are oxidizing agents, but the monomeric form (•NO2) is much more reactive. Nitrogen dioxide radical is a relatively strong one-electron oxidant, with reduction potential E(NO2•/NO2-) = 1.03 V. 3.4.2. Generation 3.4.2.1. Cell-free Systems For the specific formation of nitrogen dioxide radical, the one-electron oxidation of nitrite anion can be carried out by employing radiolytic techniques. For that purpose the solution of nitrite is saturated with N2O to convert eaq- into •OH, which reacts with NO2- to generate •NO2 (reaction 74) [51]. •

OH + NO2- → •NO2 + -OH

(74)

Another pathway to generate •NO2 by a radiolytic method is to reduce NO3- by hydrated electron (reactions 75-76). eaq- + NO3- → NO3•2-

(75)

NO3•2- + H2O → •NO2 + 2 -OH

(76)

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In this case, the hydroxyl radical needs to be removed from the system, which may be achieved by addition of small amounts of nitrite to produce more •NO2 radicals via reaction 74. An alternative enzymatic method is based on MPO-catalyzed oxidation of NO2- by H2O2 (reactions 77-79) [52]. MPO + H2O2 → MPO-compound I + H2O

(77)

MPO-compound I + NO2- → MPO-compound II + •NO2

(78)

MPO-compound II + NO2- → MPO + •NO2

(79)

The decomposition of peroxynitrite (ONOO-) at neutral or acidic pH leads to the formation of •NO2; however, it is always accompanied by other reactive radicals such as •OH (reaction 80) or CO3•- (in the presence of CO2, reaction 81) [50, 53]. In addition, the reactivity of ONOO- or ONOOH per se with the investigated compounds should be taken into account. ONOO- + H+ ⇄ ONOOH → HNO3 (~70%); •OH + •NO2 (~30%)

(80)

ONOO- + CO2 ⇄ ONOOCO2- → NO3- (~65%); CO3•- + •NO2 (~35%)

(81)

3.4.2.2. Biological Systems In biological systems, the induction of NO2• formation is achieved by stimulation of •NO production. This may increase the concentration of •NO2 which is the intermediate involved

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in •NO autooxidation (reaction 51). Induction of •NO production will lead to increased concentration of NO2- which may be converted into •NO2 by H2O2/MPO system (reactions 77-79). Also stimulation of co-generation of •NO and O2•- will lead to formation of ONOO-, which upon decomposition will produce •NO2. Thus, when the production of both •NO and ROS are stimulated, the formation of •NO2 can be expected to proceed via peroxidasecatalyzed reactions and/or peroxynitrite formation/decomposition. 3.4.3. Detection 3.4.3.1. Cell-free Systems Because NO2• exhibits weak and broad absorption at 300 - 500 nm (with a maximum at 400 nm, = 200 M-1cm-1), its reaction can be directly monitored by fast kinetic spectrophotometry (e.g. pulse radiolysis) when high concentrations of the radicals are produced. In most cases, however, the detection of •NO2 is based on its reaction of with spectroscopic probes, including ABTS2-, dihydrorhodamine (RhH2), dichlorodihydrofluorescein (DCFH2) or phenolic compounds. As •NO2 is a relatively strong oxidant, the reaction with the probes proceeds via one-electron oxidation, with the formation of radical species derived from the probe molecule: ABTS•- (reaction 82, k82 = 2.2 × 107 M-1s-1), rhodamine radical (RhH•, reaction 83, k83 = 4-7 × 105 M-1s-1), fluorescein radical (DCFH•, reaction 84, k84 = 1.3 × 107 M-1s-1) or phenoxyl radical (PhO•, reaction 85, k85 = 2.9 × 107 M-1s-1 for tyrosine dianion (phenolate anion), k85 < 105 M-1s-1 for tyrosine at pH < 7). C2H5 -

O3S

C2H5

N S

+

N N

S

•NO 2

O3S

S

+

N N

SO3-

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N

-

S

NO2-

SO3-

N C2H5 ABTS•-

ABTS2-

(82)

O

O H O

O +

H2N

O

•NO 2

+ H2N

NH2

(83) OH

OH H O

O DCFH2

OH

O

Cl

Cl +

HO

+ H+

RhH•

RhH2

Cl

NO2-

NH2

O

Cl

•NO 2

+ HO

O DCFH•

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NO2-

+ H+

OH

(84)

234

Jacek Zielonka and Balaraman Kalyanaraman O

O

OH •NO 2

+ R

+

+ H+

R

R

PhO-

NO2-

PhO•

(PhOH)

(85)

While in the case of fast kinetic methods (pulse radiolysis, flash photolysis) all of these radicals may be detected in “real time”, only ABTS•- can be detected by steady-state methodologies due to its relative stability. [Note: ABTS•- is also referred to as ABTS radical cation (ABTS•+) as it is a product of one-electron oxidation of ABTS. However, because the parent molecule is a dianion, the product of one-electron oxidation is formally a radical anion and we will maintain this designation here]. ABTS•- can be detected and quantified by spectrophotometry using one of two absorption bands with maxima at 420 nm ( = 3.6 × 104 M-1cm-1) and 735 nm ( = 1.6 × 104 M-1cm-1). In the case of dihydrorhodamine and dihydrofluorescein, the radical species must undergo further oxidation to yield fluorescent product (in case of rhodamine 123 or fluorescein, the product should be excited ca. 490 nm and the emission detected ca. 515 nm). In deaerated cell-free systems, rhodamine (Rh) can be formed by disproportionation of the rhodamine radical (reaction 86).

O

O

O

H O

O

O +

2 H2N

O

NH2

H2N

RhH•

O

NH2

H2N

RhH2

O

NH2

Rh (fluorescent)

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(86) Similarly, disproportionation of the fluorescein radical will produce fluorescein (DCF, reaction 87). COOH Cl

Cl

H

Cl

COOH Cl

COOH Cl

Cl +

2 HO

O

OH

DCFH•

HO

O DCFH2

OH

HO

O

O

DCF (fluorescent)

(87) This would yield a stoichiometry of 2:1, i.e. one molecule of rhodamine or fluorescein formed per two •NO2 radicals. However, in the presence of suitable oxidants of RhH• or DCFH• (reactions 88 and 89), the stoichiometry will change to 1:1 and the one-electron reduction product of the oxidant will be formed. RhH• + Ox → Rh + OxH• (Ox•- + H+)

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(88)

Methods of Investigation of Selected Radical Oxygen/Nitrogen Species … DCFH• + Ox → DCF + OxH• (Ox•- + H+)

235 (89)

In the presence of oxygen, O2•- will be formed [54, 55]. The reactivity of that secondary radical should be also considered when interpreting the experimental data. In the case of simple phenols (e.g. tyrosine, 4-hydroxyphenylacetic acid), the phenoxyl radical undergoes dimerization to diphenol (di-PhOH, reaction 90) or recombination with • NO2 to form nitrophenol-type product (reaction 91). R O

OH

2

OH R

R

PhO•

(90)

di-PhOH

O

OH NO2 +

R

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PhO•

•NO 2

R 3-NO2-PhOH

(91)

The relative yield of both products depends on the concentration of the phenolic probe and steady-state concentration of •NO2 radical. While the quantification of di-phenol-type product can be accomplished based on fluorescence detection, the amount of nitrophenolic product formed is generally monitored by HPLC-based techniques (with UV-Vis absorption, electrochemical or mass spectrometric detectors). 3.4.3.2. Biological Systems The quantification of •NO2 is based on the detection of the products of nitration of protein-bound tyrosine and to a lesser extent of the oxidation products of fluorogenic probes (DCFH2 and RhH2). While the principles underpinning these methods have been described above, there are some additional factors/modifications germane to biological systems. The nitrotyrosine formed by nitration of protein-bound tyrosine residues may be detected by immunochemical methods as the nitrotyrosine antibodies are commercially available. Also, chromatographic methods may be used for the detection of nitrotyrosine and the digestion of cellular proteins coupled with LC-MS analysis may help in the determination of protein nitration sites. In case of fluorogenic probes (DCFH2, RhH2), the identity of the oxidizing species will remain unknown, as most strong oxidants, including intermediates in peroxidase/H2O2 systems, can mediate the oxidation of the probes. Moreover, as discussed above, the identity and reduction products of the oxidants of the probe-derived radical is an important factor in the detection process, and these constitute largely unknown parameters in biological systems.

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3.4.4. Inhibitors and Scavengers To inhibit the formation of •NO2 in biological systems one has to attenuate the formation of ONOO- or prevent its decomposition to •NO2. The formation of •NO2 via peroxidasemediated pathways can be inhibited by peroxidase inhibition and inhibition of production of NO2- and H2O2. Thus, both ONOO-- and peroxidase-mediated •NO2 can be inhibited by suppression of •NO production, as it is a substrate in ONOO- formation as well as a precursor of NO2-. Similarly, the inhibition of the production of O2•- will diminish the amount of •NO2 formed in both pathways. On the other hand, inhibition of peroxidase may selectively inhibit the second pathway, and therefore could be used to distinguish between both pathways of • NO2 production. Although the selective attenuation of ONOO--mediated pathway may be achieved with scavengers of ONOO-, metal complexes of porphyrins may be inadequate for that purpose, as it has been postulated that •NO2 is formed during the reaction of ONOO- with porphyrin-metal complexes. Although no specific scavengers of •NO2 are available, many of the biologically-relevant reductants can react with •NO2, including ascorbic acid (reaction 92, k92 = 3.5 × 107 M-1s-1) and reduced glutathione (reaction 93, k93 = 2 × 107 M-1s-1). •

NO2 + AscH2 → NO2- + AscH• + H+

•

NO2 + GSH → NO2- + GS• + H+

(92) (93)

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3.5. Carbonate Radical Anion (CO3•-) 3.5.1. Physicochemical Properties Carbonate radical anion is the product of one-electron oxidation of carbonate anion (CO32-) [49-51] (see also Chapter 2). In aqueous environment it can be regarded as the base formed by the deprotonation of the corresponding strong acid HCO3• (pKa < 0, reaction 94). HCO3• ⇄ CO3•- + H+

(94)

Carbonate radical anion is a strong oxidant with reduction potential E(CO3•-/CO32-) = 1.59 V and therefore in most reactions it oxidizes substrates by one-electron transfer.

3.5.2. Generation 3.5.2.1. Cell-free Systems CO3•- can be produced by radiolytic methods utilizing the hydroxyl radical-mediated oxidation of carbonate dianion (CO32-, reaction 95, k95 = 3.9 × 108 M-1s-1) and bicarbonate anion (HCO3-, reaction 96, k96 = 8.5 × 106 M-1s-1) [51]. CO32- + •OH → CO3•- + -OH

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(95)

Methods of Investigation of Selected Radical Oxygen/Nitrogen Species … HCO3- + •OH → CO3•- + H2O

237 (96)

Due to the acid-base equilibria of bicarbonate (pKa(HCO3-/CO2) = 6.4, pKa(HCO3-/CO32-) = 10.4), limited solubility of CO2 in water, and its scavenging property towards hydrated electron, the radiolytic methods of CO3•- generation are limited to alkaline solutions (pH > 10). An alternative method for CO3•- production, suitable at neutral pH, is the photolysis of carbonato-metal complexes, with [Co(NH3)4CO3]+ most widely used (reaction 97) [56, 57]. [Co(NH3)4CO3]+ + h → CO3•- + Co2+ + 4 NH3

(97)

Other sources of CO3•- include specific metalloenzymes, exhibiting peroxidase-like (e.g. SOD, xanthine oxidase) activity in the presence of H2O2 and bicarbonate. However, in this case another oxidant, peroxycarbonate anion (HCO4-) will be formed (reaction 98, K = 0.32) [58]. HCO3- + H2O2 ⇄ HCO4- + H2O

(98)

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Decomposition of ONOO- in the presence of bicarbonate will result in the formation of CO3•- (reaction 81), although •NO2 will be formed as a co-product [59]. 3.5.2.2. Biological Systems The formation of CO3•- has been postulated to occur in biologic systems via decomposition of peroxynitrite in CO2-rich environments (reaction 81). In fact, it has been estimated that in most biological systems the reaction with CO2 constitutes the major pathway of ONOO- decomposition [59]. As discussed above for •NO2, to induce intracellular ONOOformation stimulation of both •NO and O2•- production is required. This pathway will lead to co-production of CO3•- and •NO2 radicals. Another pathway of CO3•- generation in biological systems may be the oxidation of CO2 (or bicarbonate) by H2O2 catalyzed by SOD in a peroxidase-like manner. This pathway may be promoted by overexpression of SOD and stimulation of H2O2 production in the presence of CO2. However, the utility of this reaction for stimulation of intracellular production of CO3•- remains to be established. 3.5.3. Detection 3.5.3.1. Cell-free Systems Carbonate radical anion exhibits a broad visible absorption band with maximum at 600 nm ( = 1860 M-1cm-1). For direct detection of CO3•-, fast spectrophotometric techniques including both pulse radiolysis and laser flash photolysis may be applied. The stationary techniques involve the use of probes, which upon one-electron oxidation yield readily detectable product. Akin to the detection of •NO2 (reactions 82-84), this includes ABTS, RhH2 and DCFH2. Additionally, a spin trapping technique has been applied for CO3•detection. The rate constant of the reaction between DMPO spin trap and CO3•- has been determined as 2.5 × 106 M-1s-1 [60]. However, the spin adduct detected is identical to the

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adduct of •OH radical (reaction 99). Thus, appropriate control experiments must be performed to confirm the identity of the trapped radical. H3C H3C

N O

H

+ CO3

H2O

DMPO

H3C H3C

N O

OH H

DMPO-•OH

(99)

3.5.3.2. Biological Systems Because similarly to •NO2, no selective probe for CO3•- is currently available, nonspecific probes including RhH2 and DCFH2 are used as intracellular reporter molecules. However, as discussed below, the results obtained with these probes are questionable. EPR spin trapping remains an option for the detection of CO3•-, at least in cell culture experiments, if EPR instrumentation is available. 3.5.4. Inhibitors and Scavengers Depending on the mechanism of CO3•- formation in biological systems, inhibition of its generation may be achieved by preventing the formation of ONOO-, or by scavenging the latter before its reaction with CO2. In situations where CO3•- is generated in a peroxidase-like pathway (e.g. via SOD or XO), depletion of intracellular H2O2 may provide an effective method to inhibit CO3•- formation. Although no specific scavengers of CO3•- are available, ascorbic acid, reduced glutathione (GSH), NADH, histidine and tyrosine can efficiently reduce CO3•- to CO32-/HCO3-.

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3.6. Hydroxyl Radical (•OH) 3.6.1. Physicochemical Properties Hydroxyl radical is a product of one-electron oxidation of water (or hydroxyl anion, OH), and is the strongest one-electron oxidant that can be efficiently generated in aqueous solutions [23, 50] (see also Chapter 2). It exists in equilibrium with its deprotonated form, O•(pKa = 11.9, reaction 100). •

OH ⇄ O•- + H+

(100)

The reduction potential of •OH has been reported as 2.7 V in acidic solution and 1.8 V in neutral solution. While both acid-base forms of hydroxyl radical behave as strong oxidants and can abstract hydrogen from organic molecules, they behave differently in many reactions due to the electrophilic nature of •OH and nuclephilic properties of O•-. Therefore •OH readily adds to unsaturated bonds, including aromatic systems, while O•- does not.

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3.6.2. Generation 3.6.2.1. Cell-free Systems The radiolysis of N2O-saturated aqueous solution is the most selective method for generation of hydroxyl radical [23]. In that system •OH is generated in high yield (radiation yield of 6) due to conversion of hydrated electron into •OH by dissolved N2O (reaction 21). H2O2 can also be used to convert hydrated electron into •OH (reaction 101). eaq- + H2O2 → •OH + -OH

(101)

However, as H2O2 can also react with •OH to form O2•-/HO2• (reaction 41), one must adjust the concentrations of the reagents very carefully to assure the correct identity of the target oxidant. Additionally, irradiation of H2O2 solution with UV light can be used to generate •OH (reaction 102). H2O2 + h → 2 •OH

(102)

In the experimental design one must take into account the possibility of the occurrence of reaction 41. The other common approach to generation of hydroxyl radical, also described as a radiolysis “mimetic” system, is the use of Fenton‟s reagent. In this system, the reduced iron ion (ferrous cation, Fe2+) reacts with H2O2 to produce ferric cation (Fe3+) and hydroxyl radical (reaction 103) [61, 62] (see also Chapter 5).

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Fe2+ + H2O2 → Fe3+ + •OH + -OH

(103)

Although controversy exists whether •OH is actually produced in that system under different reaction conditions, with the alternative oxidant FeO2+ the chemical reactivity of the oxidant formed closely resembles that of •OH. Because at neutral pH Fe2+ undergoes rapid auto-oxidation, the ferrous ion needs to be chelated to inhibit its reaction with oxygen. EDTA or DTPA chelates of Fe2+ react with H2O2 relatively slowly, thus allowing for steady-state production of •OH. The major drawback of the Fenton‟s system is the possibility of interference by reaction components with the processes investigated. For example Fe2+ or iron chelator may react with •OH radical formed, and both iron ions and H2O2 may interfere with the reaction of the radical products of the reaction of •OH with the scavengers investigated. 3.6.2.2. Biological Systems It is generally assumed that the major source of hydroxyl radical in biological systems is Fenton-type reaction (reaction 103) [63]. Thus, the stimulation of the cellular production of H2O2 and of cellular iron uptake may lead to intracellular generation of •OH. Stimulation of the production of O2•- may foster the •OH generation by increasing H2O2 levels and by facilitating the Haber-Weiss reaction (also referred to as superoxide-driven Fenton reaction) (reaction 104) O2•- + H2O2 → •OH + -OH + O2

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(104)

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This reaction requires metal ion catalysis, and it is believed that in biological systems iron and copper ions are the most important catalysts. Another potential pathway of promoting •OH formation by O2•- is the release of iron ions from iron-sulfur clusters of metalloenzymes, e.g. aconitase. In this scenario, tightly-bound iron ion in protein is released and exists as “loosely bound iron” which may then catalyze reaction 104. H2O2 has been shown to induce cellular uptake of iron from extracellular fluid, further setting the stage for Fenton‟s reaction (see Vol. II, Chapter 19). Another pathway for intracellular •OH formation is the decomposition of ONOO- in a CO2-free environment (reaction 80). This process has been implicated in the pathogenic effects of ONOO-. However, as discussed before, in that system •OH is co-generated with • NO2 radical. 3.6.3. Detection

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3.6.3.1. Cell-free Systems Due to the presence of only weak ultraviolet absorption band (with a maximum at ~225 nm), •OH is typically detected using probe molecules, even with fast kinetic methods such as pulse radiolysis. The probes for spectrophotometry include ABTS2-, ferricyanide (Fe(CN)64-) and other compounds yielding relatively stable one-electron oxidation products. These methods are considered to lack selectivity, as they are based on the one-electron oxidizing properties of •OH which to some extent are shared with many other radical species (e.g. CO3•and peroxyl radicals). The common approach to determine the identity of the oxidant is the use of •OH scavengers which lack reactivity towards other radicals (see below). An approach that is more selective for •OH is based on •OH-induced hydroxylation of aromatic molecules. As hydroxyl radical adds to the aromatic rings, at least one of the products formed is the hydroxy derivative derived from the oxidation of the •OH adduct, hydroxycyclohexadienyl radical (reaction 105). Ox

H OH

OxH•

OH

H •

+

OH

(105) Examples of •OH probes include phenylalanine, which undergoes hydroxylation to the isomeric forms of tyrosine (reaction 106), and salicylic acid, which undergoes hydroxylation to dihydroxybenzoic acid (see below) [1]. In most cases the products require detection by chromatographic methods (e.g. HPLC) to separate and quantify probe and products. OH OH +

•OH

,

,

OH COOH H2N

COOH H2N ortho-

Phenylalanine

COOH H2N metaTyrosine

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COOH H2N para-

(106)

Methods of Investigation of Selected Radical Oxygen/Nitrogen Species …

241

Another widely used method for the detection of •OH is based on •OH-induced degradation of 2-deoxyribose to malondialdehyde (reaction 107), a so-called thiobarbituric acid (TBA) - reactive substance (TBARS); malondialdehyde reacts with thiobarbituric acid (TBA) to form a potent chromophore (reaction 108) absorbing at 532 nm ( = 1.54 × 105 M-1cm-1) under acidic conditions [64]. Because the product also emits fluorescence (with maximum at 553 nm), it can be quantified by UV-Vis absorption and/or fluorescence spectroscopic techniques. HPLC analysis of the product may determine the identity of the TBARS. O

OH

+

HO

•

O

O

OH H

H

HO 2-deoxyribose

malondialdehyde

O O

O +

H

H

2

malondialdehyde

OH

HN S

(107) OH

N N H

O

S

N N

OH HO

N

+

2 H2O

SH

chromogenic product

thiobarbituric acid

(108) A major class of probes which provides both qualitative and quantitative information on the production of •OH are spin traps. A prime example is DMPO, which upon reaction with • OH yields product (reaction 109) with a characteristic 4-line spectrum (Figure 2).

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H3C H3C

+ N O

•

H

OH

H3C H3C

N O

OH H

DMPO-•OH

DMPO

(109)

In contrast to the trapping of O2•-, the rate constant of the reaction of spin traps with •OH is high. Therefore high concentrations of the spin trap are needed only if there are other •OH scavengers present at high concentrations in the test system. Although the EPR spectrum of the spin adduct is characteristic for hydroxyl radical, the same radical adduct is observed in the case of carbonate radical anion (reaction 99). Also, with some spin traps, the superoxide spin adduct may undergo spontaneous (DMPO, reaction 47) or enzymatic (DEPMPO, reaction 110) conversion to hydroxyl radical spin adduct. C2H5O O C2H5O P H3C N O

OOH H

DEPMPO-•OOH

reduction

C2H5O O C2H5O P H3C N O DEPMPO-•OH

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OH H

(110)

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It has been also suggested, that the adduct may be formed by “reverse spin trapping” whereby metal-catalyzed addition of water (reaction 111) is followed by one-electron oxidation of the hydroxylamine to nitroxide radical (reaction 112). H3C H3C

N O

H3C H3C

+ H2O

H

DMPO

DMPO-hydroxylamine Ox

H3C H3C

N OH

N OH

OH H

(111)

OxH•

OH H

H3C H3C

N O

OH H

DMPO-•OH

DMPO-hydroxylamine

(112)

To confirm the identity of the species forming the •OH adduct, one may add dimethylsulfoxide (DMSO) or ethanol (EtOH) as scavengers of •OH (reactions 113 and 114).

H3C

O S

+

CH3

•

OH

O H3C S CH3 OH

O H3C S

DMSO

methanesulfinic acid

CH3CH2OH

•

+

OH

CH3•CHOH

+

•

CH3

(113)

H2O

EtOH

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+

OH

(114)

The occurrence of •OH radical is proven if the EPR signals of •OH adduct are replaced by methyl (from DMSO, reaction 115) or hydroxyethyl (from EtOH, reaction 116) radical adduct. H3C H3C

+ N O

H

•

CH3

H3C H3C

N O

CH3 H

DMPO-•CH3

DMPO

(115) OH

H3C H3C

+ N O DMPO

H

CH3•CHOH

H3C H3C

N O

H

CH3

DMPO-•CH(OH)CH3

(116)

3.6.3.2. Biological Systems The high value of rate constants of the reactions between •OH and the detection probes indicate that even at relatively low probe concentrations, the entire pool of •OH should be trapped (and detected). In reality, due the overall very high reactivity of •OH towards most

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species …

243

organic cellular constituents, only small portions of intracellular •OH are detected. Even if the intracellular concentration of the probe is raised to 10 mM, it can be assumed that less than 1% of •OH will be trapped. The most popular probe used is salicylic acid (SA), as it can be implemented in relatively high concentrations without apparent toxicity. The reaction between SA and •OH yields catechol (hydroxylation/decarboxylation product) and two isomeric hydroxylation products: 2,3-dihydroxybenzoic acid (2,3-DHBA) and 2,5dihydroxybenzoic acid (2,5-DHBA) (reaction 117). COOH OH

OH

COOH OH

OH +

•OH

,

salicylic acid

catechol (11%)

COOH OH ,

OH HO 2,3-dihydroxybenzoic 2,5-dihydroxybenzoic acid (49%) acid (49%)

(117)

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Of the two dihydroxy isomers, 2,5-DHBA can be also formed enzymatically (e.g. via cytochrome P-450-catalyzed oxidation), while 2,3-DHBA is believed to be a specific product of the reaction with •OH. HPLC-based separation coupled with electrochemical detection and quantification of 2,3-DHBA is useful for monitoring the formation of •OH in biological systems. However, factors such as the stability of the product formed and intracellular uptake of the probe (SA or its precursor, aspirin) must be considered. EPR spin-trapping of •OH has been used for its detection in cellular systems. However, the applicability of this technique for intracellular detection of •OH (and of any other radical) is limited by the possibility of conversion of EPR-active nitroxide (spin adduct) to EPR-silent oxidation and/or reduction products (oxo-ammonium cation and hydroxylamine, respectively, reactions 118 and 119). H3C H3C

N O

OH H

oxidation

DMPO-•OH H3C H3C

N O

OH H

DMPO-•OH

H3C H3C

N O

OH H

oxoammonium cation reduction

H3C H3C

N OH

(118)

OH H

hydroxylamine

(119)

The fluorescent probes HPF and APF have been reported to be oxidized to fluorescein by OH (reactions 120 and 121), with propensity of the latter to undergo oxidation also by HOCl [6]. •

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COOH

COOH + O

O

•OH

O

HO

O

HPF

O

O

fluorescein

O

(120)

OH

COOH

COOH + O

O APF

O

•OH

O

HO

NH

O

O

fluorescein

NH2

(121)

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The conversion of probes to product is based on the deprotection of the hydroxyl substituent of the fluorescein moiety, which is accompanied by ca. 100-fold increase in the fluorescence quantum yield. Although the mechanism of the conversion has not been thoroughly investigated, the HPF probe exhibits selectivity towards •OH, with the additional possibility of oxidation in peroxidase-catalyzed reactions with H2O2. 3.6.4. Inhibitors and Scavengers In cell-free systems addition of hydroxyl radical scavengers (e.g. DMSO, EtOH) is the simplest and most effective method to remove •OH. However, secondary, scavenger-derived radicals will be formed, which upon reaction with oxygen may generate peroxyl-type radicals. An alternate approach may entail use of less selective scavengers (e.g. ABTS, Fe(CN)64-) which yield less reactive products in reaction with •OH. Due to high concentration of endogenous intracellular scavengers of •OH (most intracellular organic matter) it is practically impossible to lower significantly •OH levels by addition of drug which acts solely as •OH scavenger. To decrease •OH concentration by a factor of 2, one would need to add an amount of drug equivalent to that of total intracellular organic matter! This also implies that attribution of the protective effects of any compound to • OH scavenging is questionable. Thus, the only practical method to decrease intracellular levels of •OH is to prevent its formation. In the case of Fenton‟s reaction, curtailment of the availability of metal ions and H2O2 should diminish •OH production. Decreases in intracellular H2O2 concentrations can be accomplished by inhibition of its production (e.g. inhibition of flavoproteins by diphenylene iodonium, DPI) or stimulation of its removal (e.g. by supplying the cells with reduced glutathione). Administration of potent, cell-permeable iron chelators may also effectively attenuate Fenton‟s reaction. In fact, the chelator desferoxamine is widely used clinically to ameliorate iron-dependent pathologies.

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3.7. Organic Peroxyl Radicals (ROO•) 3.7.1. Physicochemical Properties Peroxyl radicals are formed as products of oxygen addition to carbon- or heteroatomcentered radicals (reaction 122) [65]. R• + O 2

ROO•

(122)

In most cases the addition of oxygen is fast (k ~ 109 M-1s-1) and irreversible. Compounds with highly delocalized spin density and thiyl radicals (RS•) are the exception to this rule. Also, in the case of highly reducing radicals (pyridynyl radicals, Pyr•, including paraquat radical cation and the radical of nicotinamide adenine dinucleotide, NAD•) peroxyl radicals have not been detected and direct electron transfer to oxygen has been proposed (reaction 123) [66]. +

O2

+

N R

N R

Pyr•

Pyr+

O2•-

(123)

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Peroxyl radicals are relatively strong one-electron oxidants, with reduction potential E(ROO•, H+/ROOH) in the range of 0.7 – 1.2 V for aliphatic peroxyl radicals. In the case of radicals R• of reducing character, the corresponding peroxyl radicals may undergo spontaneous or base-catalyzed elimination of HO2• or O2•-, e.g. -hydroxyperoxyl radicals (reaction 124) [67].

R1

OH OO• R2

O R1

R2

+

O2•-

+ H+

(124)

Peroxyl radicals may also react with other molecules by addition to unsaturated bonds or oxygen atom transfer with the formation of alkoxyl (RO•) radical. 3.7.2. Generation 3.7.2.1. Cell-free Systems Aerobic Generation of Peroxyl Radicals A typical method for generation of peroxyl radical is the production of the oxygenreactive radical (R•) from substrate molecule (S) in the presence of O2 (reactions 125-126) [65, 68]. S

R•

R• + O 2

(125) ROO•

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Jacek Zielonka and Balaraman Kalyanaraman

Various methods of production of ROO• (radiolytic, photolytic, and thermal radical production) differ mainly in the way the initial radical R• is generated. Radiolysis Radiolysis of aqueous (or organic) solutions is the most universal method for the preparation of radical species from diverse organic compounds. One-electron oxidation of the substrate S will lead to the formation of the corresponding radical cation S•+ (reaction 127). S•+

S - e-

(127)

Because radical cations are in most cases strong acids, they undergo deprotonation in aqueous solution with the formation of neutral radical •S(-H) (equivalent to R•, reaction 128). S•+ + H2O

R• + H3O+

(128)

Direct abstraction of hydrogen atom may be used to generate the radicals. In the case of alkyl radicals, the substrate reacts with •OH (reaction 129) and H• (reaction 130) to form the corresponding radical R•. S + •OH

R• + H2O

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S + H•

(129)

R• + H 2

(130)

Substrate radical may also arise by addition of •OH to an unsaturated bond. For example, addition of •OH to aromatic rings yields carbon-centered cyclohexadienyl-type radical (first step of reaction 105). One-electron reduction of organic compounds may also yield neutral radicals, precursors of peroxyl radicals, via intermediate formation of radical anion S•(reaction 131). The latter is typically a weak acid and undergoes protonation to form neutral radical •S(+H) (equivalent to R•, reaction 132). S + e-

S•-

S•- + H2O

(131) R• + -OH

(132)

A relevant example is provided by carbonyl compounds which upon one-electron reduction and protonation yield carbon-centered ketyl radicals (reaction 133). O R1

O

1-e reduction R2

R1

OH

H+ R2

R1

R2

(133)

One-electron reduction of alkyl halides (RX, where X is a halogen atom) is another convenient way of generating alkyl radicals (reaction 134). RX + e-

R• + X -

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species …

247

An example is the radiolytic generation of trichloromethylperoxyl radical, widely used as a highly-reactive model of biologically-relevant peroxyl radicals. The radiolysis of the O2saturated mixture of water and 2-propanol (48% by vol., each) and tetrachloromethane (CCl4, 4% by vol.) generates CCl3OO• radical in high yield. In this system, both solvated electron and 2-PrOH-derived ketyl radicals reduce CCl4 with the formation of •CCl3 and subsequently CCl3OO• radical (reactions 135-137) [69]. •

CCl4 + e-

CCl3 + Cl-

CCl4 + (CH3)2C•OH •

CCl3 + O2

(135)

•

CCl3 + Cl- + H+ + (CH3)2CO

CCl3OO•

(136) (137)

One-electron reduction of aryl bromides may also serve as a convenient method for production of arylperoxyl radicals (reactions 138-139) [70, 71]. Br +

eaq-

+

bromobenzene

Br-

phenyl radical

(138)

OO• +

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phenyl radical

O2 phenylperoxyl radical

(139)

Similar to the radiolysis system, Fenton‟s reagent may be used to generate substratederived radicals as precursors of peroxyl radicals. Azo Compounds (Free Radical Initiators) The use of the azo compounds, which upon thermal decomposition yield carbon-centered radicals, is a distinct and convenient approach for generation of alkylperoxyl radicals in a controlled manner [72]. In the first step the substrate decomposes unimolecularly to two radicals and nitrogen molecule (reaction 140). R-N=N-R

2 R• + N 2

(140)

The radicals formed (R•) may undergo recombination within the solvent cage or escape producing “free” radical in bulk solvent. In the presence of oxygen the radical will give rise to ROO• via reaction 126. Two azo initiators are most frequently employed: (a) hydrophilic AAPH (2,2‟-azobis(2-amidinopropane) dihydrochloride, reaction 141) and (b) lipophilic AMVN (2,2‟-azobis(2,4-dimethylvaleronitrile, reaction 142).

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Jacek Zielonka and Balaraman Kalyanaraman NH H2N

NH N

N

NH2

2

+

H2N

N2

NH AAPH

(141)

NC N

+

2

N

CN

N2

CN

AMVN

(142)

The rate of the decomposition of AAPH is controlled mostly by the temperature and to a lesser extent by type of solvent and pH of the solution. At 37 oC the initial flux f of the generated radicals R• is given by the equation 7. f (M/s) = 1.36 × 10-6 × [AAPH]

(Equation 7)

where [AAPH] is the concentration of AAPH in M (moles/dm3). Upon photolysis many azoalkanes undergo decomposition with the formation of carboncentered radicals (reaction 143). R-N=N-R + h

2 R• + N 2

(143)

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Anaerobic Generation of Peroxyl Radicals In situations where the presence of oxygen is not desirable, peroxyl radicals can be formed from compounds bearing peroxy moieties, including peroxynitrates and hydroperoxides. In the former case, peroxynitrate (ROONO2) exists in solution in equilibrium with corresponding peroxyl radical and nitrogen dioxide radical (reaction 144). ROONO2 ⇄ ROO• + •NO2

(144)

Here, the formation of ROO• is accompanied by the release of •NO2, and scavenging of any of these radicals will shift the equilibrium towards greater radical production. The major limitation of this method is the requirement for preparation and isolation of the corresponding peroxynitrate substrate. In the case of hydroperoxides (ROOH), the corresponding peroxyl radical is formed by hydrogen abstraction from the hydroperoxide (reaction 145). ROOH + X

ROO• + HX

(145)

In this reaction X denotes a hydrogen atom acceptor, which in practice can be •OH, F• or Cl . The simplest example of this reaction is the formation of hydroperoxyl radical (HO2•) when •OH is formed in the presence of H2O2 (reaction 41). •

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249

3.7.2.2. Biological Systems Peroxyl radicals are believed to be crucial intermediates formed upon oxidative insult to the cellular environment [1, 73]. The abstraction of hydrogen atom from lipid molecules by any of the radical species mentioned above will ultimately lead to the formation of lipidderived free radicals (reaction 146) which upon reaction with oxygen will generate peroxyl radicals (reaction 147). Due to their highly-oxidizing character, lipid peroxyl radicals can abstract hydrogen from other lipid molecules (reaction 148) leading to self-propagating events known as lipid peroxidation (Chapter 7). LH + R1• L• + O2 LO2• + LH

L• + R1H LO2• LOOH + L•

(146) (147) (148)

In the course of auto-catalytic lipid peroxidation, a wide range of free radical species may be generated including O2•-, NO2•, CO3•-, •OH and lipid peroxyl radicals. Direct stimulation of intracellular lipid peroxidation can also be achieved by treatment with lipophilic free radical initiators (e.g. AMVN, reaction 142). In this case alkyl radical generated from AMVN will react with oxygen to form peroxyl radical which, in turn, may initiate lipid peroxidation.

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3.7.3. Detection 3.7.3.1. Cell-free Systems With the exception of vinyl, aryl and thiyl peroxyl radicals, peroxyl radicals are generally not easy to detect by fast kinetic methods due to their spectral characteristics, exhibiting only weak absorption in the UV region. Vinyl and aryl peroxyl radicals are characterized by absorption bands with maxima between 400 - 600 nm and extinction coefficients within the range of (1 - 2) × 103 M-1cm-1. In addition, thiyl peroxyl radicals absorb in the visible light region with maxima close to 550 nm, although with lower extinction coefficients (ca. (2 - 4) × 102 M-1cm-1). In most cases the detection of peroxyl radicals is based on their relatively strong oxidizing nature, and probes which upon oxidation yield easily detectable products can be implemented. ABTS2- is commonly used because of the spectral properties and stability of the product, ABTS•-. In the case of chain reactions involving peroxyl radicals (e.g. lipid peroxidation), measurement of the rate of oxygen consumption in solution has been successfully applied for interrogation of the kinetics of those systems. It has been assumed that the step involving oxygen consumption is the formation of peroxyl radical(s), typically from carbon-centered radicals.

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250

Jacek Zielonka and Balaraman Kalyanaraman In addition, EPR spin-trapping can effectively detect peroxyl radicals (reaction 149). H3C H3C

+ ROO• N O

H

H3C H3C

N O

OOR H

DMPO-•OOR

DMPO

(149)

Because the spin traps can also efficiently trap the carbon-centered radicals, EPR spintrapping of peroxyl radical is used when the radical is formed via pathways not involving alkyl radicals, for example during oxidation of organic hydroperoxides. Due to the oxidizing properties of ROO•, in the presence of suitable reductants the corresponding hydroperoxide ROOH is formed (reaction 150). R1OO• + R2H

R1OOH + R2•

(150)

Thus, detection of hydroperoxide is a good indication of the formation of peroxyl radical(s). ROOH can be detected by HPLC equipped with electrochemical detectors. Furthermore, reaction of hydroperoxides with diphenyl-1-pyrenephospine (DPPP, reaction 151) may be used for ROOH detection, as the product, DPPP oxide (DPPP=O) fluoresces with excitation at 351 nm and emission at 380 nm [74, 75].

P

P O

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+ ROOH

DPPP

+ ROH

DPPP=O

(151)

This reaction can be used for monitoring total pools of hydroperoxides, or can be applied for post-column derivatization in HPLC-fluorescence based detection of individual hydroperoxides. Biological Systems Due to the lack of specific probes for peroxyl radicals, their formation in biological systems is usually investigated by following the formation of various markers of ROO•, including hydroperoxides, and non-specific oxidation products such as aldehydes (e.g. 4hydroxynonenal, MDA), isoprostanes etc. Also, fluorogenic probe for hydroperoxides (DPPP, reaction 151) can be used as an indirect way of monitoring peroxyl radical formation. Inhibitors and Scavengers The method of inhibition of peroxyl radical formation includes inhibition or scavenging of precursor radicals (e.g. carbon-centered radical) and/or deoxygenation of the solution. Due to the high rate constant of the reaction between carbon-centered radicals (R•) and oxygen, scavengers of R• should be added in high excess relative to O2. In biological systems the

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species …

251

inhibition of formation of R• radicals is the most effective way to prevent ROO• generation. For that purpose substrates for the Fenton reaction should be suppressed by iron chelation and by preemption or scavenging of O2•- and H2O2. In cell-free systems scavenging of peroxyl radical can be accomplished with most antioxidants, including reduced glutathione (GSH, reaction 152), ascorbic acid (AscH2, reaction 153) and phenolic compounds (PhOH, e.g. tocopherols, polyphenols, reaction 154). ROO• + GSH

ROOH + GS•

(152)

ROO• + AscH2

ROOH + AscH•

(153)

ROO• + PhOH

ROOH + PhO•

(154)

As discussed above, in biological systems peroxyl radicals are classically formed within membranes during lipid peroxidation. Therefore, the most common strategy to scavenge intracellular peroxyl radicals is to supplement cells with lipophilic antioxidants such as and -tocopherol (Vol. II, Chapter 3). It must be borne in mind, however, that radicalscavenging is not the sole activity of tocopherols (vitamin E) and the observed effects of treatment with tocopherols should not be relied upon as exclusively indicating peroxyl radical formation.

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4. Mechanistic Aspects and Difficulties Encountered with the Use of Probes for Radical Detection Spectroscopic probes offer the possibility of non-invasive, often real-time monitoring of oxidant production and are thus widely employed in cell-free, tissue culture and in vivo experiments. However, the exact mechanism of conversion of probe to detectable product is not always understood. Situations whereby probe may yield false information concerning rates of oxidant production are presented below.

4.1. The Probe Requires “Activation” to React with the Radical In many cases the probe by itself is unreactive towards the radical or the reaction is too slow for efficient competition with other pathways of radical decay in biological systems. In those cases the probe needs firstly to be “activated” by its conversion into a radical-type derivative. The consecutive step involves a radical-radical reaction and is usually very fast. The need for activation introduces additional variables to quantitative analysis, as the changes in the rate of probe activation will affect the rate of the reaction with the radical detected, and thus will impact the magnitude of the signal measured. In other words, at constant flux of radical, changes in the rate of probe activation may give rise to spurious observations

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Jacek Zielonka and Balaraman Kalyanaraman

concerning rates of radical production. Examples of probes requiring activation include luminol (LumH2), lucigenin (Luc2+) and diaminofluorescein (DAF-2). Luminol Luminol (LumH2) is a chemiluminescent probe which has been widely used for the detection of O2•- and H2O2 (Chapter 12). LumH2 does not react directly with O2•-, and needs to be activated by one-electron oxidation to the radical LumH• (reaction 155, step a). It is the radical LumH• which reacts with O2•- to form hydroperoxide (LumH-OOH, step b) which undergoes conversion to peroxyacid (step c). Peroxyacid isomerizes to endoperoxide (step d) which upon elimination of N2 (step e) yields 3-aminophthalate in the electronically excited state (step f), followed by emission of light during relaxation to the ground state (step g). NH2 O

NH2 O N NH

1-e oxidation (155a)

O LumH2

NH2 O O2

N N

•-,

H+

NH2 O N N

(155b)

HO OOH

OH LumH•

N NH OOH

(155c)

O peroxyacid

LumH-OOH

(155d) NH2 O

NH2 OH OH OH

+

O

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aminophthalic acid in ground state

h (155g)

O O

*

NH2 OH

(155f)

OH aminophthalic acid in excited state

O O

NH2 OH

N NH O O

- N2 (155e)

OH

O endoperoxides

(155)

Lucigenin Lucigenin (Luc2+) is another widely used chemiluminescent probe for O2•-; however, in contrast to luminol, the first step of the reaction involves one-electron reduction (reaction 156, step a). The second step involves the reaction of Luc•+ with O2•- with the formation of a dioxetane ring linking two acridane moieties (step b). Dioxetane derivative undergoes dissociation with the formation of two molecules of N-methylacridone: one in electronic ground state and one in the excited state (step c). The relaxation of the excited state to the ground state is accompanied by the emission of light (step d). One-electron reduction of Luc2+ to Luc•+ can be accomplished by flavoproteins as well as •O2 . However, as the estimated rate constant of the reaction between Luc2+ and O2•- is relatively low (ca. 6 × 104 M-1s-1) in biological systems, in the presence of other scavengers of O2•- (e.g. SOD) the occurrence of direct reduction of Luc2+ by O2•- is highly unlikely. As discussed below, the occurrence of the reverse reaction, reduction of oxygen by Luc•+, is much more probable.

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species … CH3 N

CH3 N

253

CH3 N

1-e reduction

O2•-

(156a)

(156b)

O

O

N CH3

N CH3

N CH3

Luc2+

Luc•+

dioxetane (156c)

O

*

O h

N CH3 N-methylacridone in excited state

(156d)

N CH3 N-methylacridone in ground state

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(156)

Diaminofluorescein As discussed in paragraph 3.3.3, diaminofluorescein (DAF-2) is a fluorogenic probe used for monitoring of intracellular •NO production. The probe does not react directly with •NO, but with the product(s) of its oxidation. Of the two possible pathways for the formation of fluorescent product, one involves one-electron oxidation of the aromatic amine moiety to aminyl radical (reaction 65, step b). In the subsequent step •NO reacts with DAF-2-derived radical to form nitrosoamine (step c), which subsequently undergoes conversion into the triazole derivative, DAF-2T (step d). As it has been argued that N2O3-dependent nitrosation of DAF-2 may be of negligible importance in cellular systems, the activation of DAF-2 by one-electron oxidation to iminyl radical may be one of the limiting factors in the detection of intracellular •NO. Hydroethidine Hydroethidine oxidation to 2-hydroxyethidium (reaction 49) is a multistep process involving the reaction of O2•- with HE-derived radical. HE-derived radical is formed by oneelectron oxidation of HE (step a). In contrast to LumH2 probe, the first step of the reaction can also be accomplished by O2•-. There is, however, the possibility of HE oxidation to its radical by other oxidants (including heme proteins) which would be reflected in the yield of additional, dimeric products (reaction 157) formed upon one-electron oxidation of HE (step a), followed by radical dimerization (step b) and further oxidation (steps c and d).

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Jacek Zielonka and Balaraman Kalyanaraman H2N

H2N

1-e oxidation (157a)

H N C2H5

NH2

+

H2N -H

H N C 2H 5

+

H

+H+

NH2

N C2H5

HE•+

HE

NH

HE(•NH) dimerization (157b)

C2H5

H2N

H 2N

oxidation (157c)

H N C2H5

C2H5

H2N

H2N

N

N H

H

NH2 NH2

HE-E+

N C2H5

NH2 NH2

HE-HE

oxidation (157d)

C2H5

H 2N

H2N

N

N C 2H 5

NH2 NH2

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(157) The O2•--independent oxidation of HE may significantly increase its efficiency for O2•scavenging, as it has been estimated that the rate of the reaction with O2•- is three orders of magnitude higher for HE radical than for HE. Nitrotyrosine Formation of nitrotyrosine from tyrosine in the presence of •NO2 is a multistep process (reaction 158) with the oxidation of tyrosine to tyrosyl radical as the first step (step a). This step can be accomplished by •NO2, but in biological systems this reaction is rather unlikely due to the relatively low reactivity of •NO2 towards tyrosine, while scavenging of •NO2 by GSH will be considerably faster. Other oxidants capable of oxidation of tyrosine involve CO3•- and compound I and II of peroxidases. After formation of the tyrosyl radical, its reaction with •NO2 will eventually lead to the formation of nitrotyrosine (step b).

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Methods of Investigation of Selected Radical Oxygen/Nitrogen Species … OH

OH

O •NO 2

1-electron oxidation (158a) COOH NH2 tyrosine

255

NO2

(158b) COOH

COOH

NH2

NH2 tyrosyl radical

3-nitro-tyrosine

(158)

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4.2. Probe-derived Transient Radical Can Induce the Formation of Radical Species The need to “activate” the probe to detect a specific radical is a confounding factor in the quantitative analysis of radical production but does not prevent the analyst from concluding that the targeted radical was formed in the investigated system, as long as the product is specific for the radical. A different situation arises when the probe, the product or any intermediate can lead to the production of the radical of interest and increase the yield of the detected product. This “self-fulfilling prophecy” disqualifies the probe for the purpose of detecting the targeted radical. In case of such probes, the use of radical-specific scavengers will inhibit the probe oxidation, leading to the false conclusion on the identity of the primary species reacting with the probe. The actual action of the scavenger is due to elimination of the secondary radicals involved in the chain reaction, not the primary radicals responsible for initiation of probe oxidation. Thus, not only the quantification but also the identification of the radicals may be confounded by the undesired reactivity of probe-derived radicals. Despite these disqualifying features, there are many probes belonging to this class which are commercially available and widely employed in research in the field of redox biology. Some examples of the phenomena of stimulation of radical/oxidant production by the probes are presented below. 4.2.1. The Probe-derived Radical Reduces Oxygen to O2•The reaction of the probe-derived radical with oxygen may lead to the formation of O2•-. Because O2•- is a precursor of other biologically-relevant radicals, this reaction may elicit a chain reaction for production of the detected radical even if the early event in the formation of probe-derived radical was due to different oxidizing species (e.g. oxidized by heme proteins). Examples can be found among spectrophotometric (nitroblue tetrazolium), chemiluminescent (lucigenin) and fluorogenic (dichlorodihydrofluorescein, dihydrorhodamine) probes. Nitroblue Tetrazolium Nitroblue tetrazolium (NBT2+) is a probe used for the detection of O2•-, as it undergoes one-electron reduction by O2•- to form the tetrazolinyl radical NBT•+ (reaction 159), which upon additional reduction (e.g. via dismutation) generates highly colored formazan (reaction 160). However, NBT2+ can be also reduced by O2•--independent pathways producing NBT•+, which upon reaction with O2 may undergo re-oxidation with the formation of O2•- (reverse reaction 159).

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Jacek Zielonka and Balaraman Kalyanaraman O

O

O

N N N

N N

N

NO2

N N

O

N + O2

•-

N N

N N

O2N

N

NO2

N N

+ O2

O2N NBT•+

NBT2+

(159) O

O

O

N N N

N N

N

1-e reduction

N N

O N

N N

N

N N

N NH

NO2

O2N

NO2

NBT•+

O2N

mono-formazan

(160)

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Lucigenin As discussed above, the conversion of lucigenin into light-emitting product requires oneelectron reduction to Luc•+, followed by its reaction with O2•-. In the presence of O2, however, Luc•+ may undergo re-oxidation to form O2•- and Luc2+ (reaction 161). CH3 N

CH3 N

+

O2

+

N CH3

N CH3

Luc•+

Luc2+

O2•-

(161)

Because the estimated equilibrium constant for the reaction 161 is ca. 50, and as there may be other pathways of O2•- decay (e.g. reaction with SOD), the probability of the forward reaction is much higher than the reverse one. One of the approaches to limit Luc2+-dependent O2•- formation in biological systems is to lower the rate of O2•--independent reduction of Luc2+ to Luc•+. It has been proposed that lowering the concentration of Luc2+ below 5 µM will eliminate the enzyme (flavoprotein)-dependent Luc2+ reduction and thus make the assay applicable for quantitative study of O2•- formation. However, decreasing the concentration of Luc2+ will decrease not only enzymatic but also O2•--dependent reduction, which is not efficient even at higher concentrations of Luc2+. More importantly, lowering the concentration of Luc2+ shifts the equilibrium state (reaction 161) further to the right, actually increasing the yield of O2•-.

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Dichlorodihydrofluorescein and Dihydrorhodamine Both DCFH2 and RhH2 react with radical oxidants with the formation of the intermediate radicals DCFH• and RhH•, respectively (see paragraph 3.4.3.1). It has been shown that both radicals react relatively rapidly with O2 to generate fluorescent products (DCF and Rh) as well as O2•- (reactions 162 and 163) [17, 54, 55].

COOH Cl

Cl

+ HO

+

O2

OH

O

COOH Cl

Cl HO

O

DCFH•

DCF

O

O + O

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RhH•

(162)

O

O

H2N

O2•- + H+

O

NH2

+

O2 H2N

O Rh

O2•-

NH2

(163)

A simple example of how these reactions may lead to misinterpretation of experimental data is the study of radical/oxidant formation during apoptotic cell death. The increase in the fluorescence intensity during apoptosis of cells loaded with fluorogenic probes has been interpreted as indicative of oxidant production during the event. However, one of the critical steps in the apoptotic pathway is the release of cytochrome c from mitochondria to cytosol. Cytochrome c is a heme protein capable of oxidizing numerous probes directly as well as in a peroxidase-like manner. One-electron oxidation of the probe DCFH2 or RhH2 will lead to the formation of the corresponding radicals, which in the presence of oxygen will generate O2•-. This will lead not only to further oxidation of the probe, but also may affect the biological pathways investigated (in this case the apoptotic signaling cascade). 4.2.2. Probe-derived Radical Oxidizes Cellular Low-molecular-Weight Reductants Apart from the reducing properties of many probe-derived radicals, some of them also exhibit oxidative properties and can react with bioreductants (GSH, NAD(P)H, AscH2) to generate reductant-derived radicals, and in most cases regenerating the probe molecules. Formation of GSH- and NAD(P)H-derived radicals in the presence of oxygen will lead to formation of O2•- (see paragraph 2.2.). A known example of such a probe is DCFH2 [76]. The reactivity of radicals formed from many radical probes towards bioreductants remains to be established.

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Jacek Zielonka and Balaraman Kalyanaraman

4.2.3. The Detection Event is Accompanied by Free Radical Production Another confounding aspect of the use of some probes is the induction of probe conversion into the product by the stimuli associated with the detection method. Relevant examples are fluorogenic probes, whose fluorescent product may act as photosensitizers of probe oxidation. It is well known that the longer one monitors the formation of fluorescent probe, the more intense is the observed signal due to photoreactivity of the fluorescent products. It has been shown that illumination of the solution of DCF in the presence of GSH or NADH leads to the reduction of DCF to DCFH•, with concomitant oxidation of the reductant (reactions 164-166) [77]. DCF + h

(DCF)*

(DCF)* + GSH

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(DCF)* + NADH

DCFH• + GS• DCFH• + NAD•

(164) (165) (166)

The formation of DCFH•, GS• and NAD• radicals in the presence of oxygen will lead to the generation of O2•- (reactions 162, 16-17 and 19, respectively), which may be transformed into additional species capable of oxidation of DCFH2. Thus, the excitation light used for fluorescence monitoring will actually stimulate oxidation of the probe. The extent of productdependent probe oxidation may be manipulated by changing the intensity of the excitation light and/or decreasing sample exposure time. Another example of photoreactivity of the fluorescent product is the photosensitized oxidation of hydroethidine (HE) to ethidium (E+) by 2-hydroxyethidium (2-OH-E+). This reactivity, which is unrelated to GSH or NADH but dependent on the presence of O2, has been postulated to explain O2•--dependent formation of E+ in biological systems. The confusion due to the photo-reactivity of 2-OH-E+ can be avoided if the HPLC-based quantitative analysis is performed, which is the preferred method for quantitation of 2-OH-E+.

4.3. The Product Detected May be Formed via Different Pathways Most of the probes used for the detection of radicals lack selectivity towards the specific radical as well as specificity for the product detected. For instance, most of the so-called leuco-dyes (e.g. DCFH2, RhH2, HE), which upon oxidation yield fluorescent product(s), can be oxidized by a variety of oxidants including heme proteins. Moreover, the product formed from different oxidants is typically the same, with the exception of HE which yield two fluorescent products, one of them, 2-OH-E+, being specific for O2•-. Furthermore, spectrophotometric probes such as cytochrome c(Fe3+) and NBT2+ can be reduced to highly colored products by a variety of reductants including enzymatic systems. In the case of cytochrome c the enzymatic reduction can be minimized by modification of the protein by acetylation or succination rendering it a poorer substrate for the enzyme(s). Another example of the lack of specificity is the conversion of tyrosine to nitrotyrosine. Although TyrNO2 has been used as a specific marker of •NO2 (formed from ONOO- or

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MPO/H2O2/NO2- system), it may be also formed in the reaction of tyrosyl radical with •NO. Similar to the reaction with NO2, the initial step involves one-electron oxidation of tyrosine (Tyr(OH)) to Tyr(O•) (reaction 167, step a), followed by radical-radical recombination between Tyr(O•) and •NO (step b). Under oxidative conditions, 3-nitrosotyrosine can be oxidized to 3-nitrotyrosine (step c) yielding the same product as from the reaction with •NO2. OH

OH

O •

1-e oxidation (167a) COOH NH2 tyrosine

COOH

COOH

oxidation (167c)

NO2

COOH

NH2

NH2

3-nitroso-tyrosine

3-nitro-tyrosine

NH2 tyrosyl radical

OH NO

NO (167b)

(167)

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Conclusions The detection of radicals and other short-lived reactive species is not an easy task in biological systems and in cell-free chemical samples. Nevertheless, understanding the basic physicochemical properties and reactivity both of the radicals investigated and of the probes used, enables in many cases reliable detection of the radicals and confirmation of their involvement in intracellular signaling. This Chapter by no means covers the whole range of radical species which may be generated in biological systems and all of the probes used for their detection. The probes selected for discussion herein intends to illustrate diverse approaches to detect radicals as well as various pitfalls associated with the application of commonly used reagents in free radical research. Rational approaches for development of new probes to detect radical species in biological systems will undoubtedly yield new, better tools lacking many of the drawbacks associated with some of the currently used probes. An understanding of the reactivity of the probe towards different radical and non-radical oxidants, including the kinetics of the reaction, is the first step towards its application for detection purposes. Also, the chemistry of any transient(s) and product(s) of the reaction of the probe with radicals needs to be carefully examined. Critical evaluation of the probe in cell-free and cellular systems is always a prerequisite for its proper use in free radical biology and medicine.

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[21] Von Sonntag C, Schuchmann HP. Pulse radiolysis. Methods Enzymol 1994; 233:320. [22] Wishart JF, Nocera DG. Photochemistry and radiation chemistry (complementary methods for the study of electron transfer). American Chemical Society; 1998. [23] Buxton GV, Greenstock CL, Helman WP, Ross AB. Critical review of rate constants for reactions of hydrated electrons, hydrogen atoms and hydroxyl radicals. Phys Chem Ref Data 1988; 17:513-886. [24] Weil JA, Bolton JR. Electron paramagnetic resonance: elementary theory and practical applications. Wiley; 2007. [25] Gore MG. Spectrophotometry and spectrofluorimetry: a practical approach. Oxford University Press, USA; 2000. [26] Lakowicz JR. Principles of fluorescence spectroscopy. 3rd ed. Springer; 2006. [27] Snyder LR, Kirkland JJ, Dolan JW. Introduction to modern liquid chromatography. Wiley; 2009. [28] Bielski BH, Cabelli DE. Highlights of current research involving superoxide and perhydroxyl radicals in aqueous solutions. Int J Radiat Biol 1991; 59(2):291-319. [29] Valentine JS, Curtis AB. Convenient preparation of solutions of superoxide anion and the reaction of superoxide anion with a copper(II) complex. J Am Chem Soc 1975; 97(1):224-226. [30] Ingold KU, Paul T, Young MJ, Doiron L. Invention of the first azo compound to serve as a superoxide thermal source under physiological conditions: Concept, synthesis, and chemical properties. J Am Chem Soc 1997; 119(50):12364-12365. [31] Bielski BHJ, Cabelli DE, Arudi RL, Ross AB. Reactivity of HO2/O2•- Radicals in Aqueous Solution. J Phys Chem Ref Data 1985; 14(4):1041-1100. [32] Fridovich I. Quantitative aspects of the production of superoxide anion radical by milk xanthine oxidase. J Biol Chem 1970; 245(16):4053. [33] Butler J, Koppenol WH, Margoliash E. Kinetics and mechanism of the reduction of ferricytochrome c by the superoxide anion. J Biol Chem 1982; 257(18):10747. [34] Frejaville C, Karoui H, Tuccio B et al. 5-(Diethoxyphosphoryl)-5-methyl-1-pyrroline N-oxide: a new efficient phosphorylated nitrone for the in vitro and in vivo spin trapping of oxygen-centered radicals. J Med Chem 1995; 38(2):258-265. [35] Zielonka J, Kalyanaraman B. Hydroethidine- and MitoSOX-derived red fluorescence is not a reliable indicator of intracellular superoxide formation: another inconvenient truth. Free Radic Biol Med 2010; 48(8):983-1001. [36] Gardner PR. Aconitase: sensitive target and measure of superoxide. Methods Enzymol 2002; 349:9-23. [37] Rebouças JS, Spasojević I, Batinić-Haberle I. Pure manganese (III) 5, 10, 15, 20tetrakis (4-benzoic acid) porphyrin (MnTBAP) is not a superoxide dismutase mimic in aqueous systems: a case of structureûactivity relationship as a watchdog mechanism in experimental therapeutics and biology. J Biol Inorg Chem 2008; 13(2):289-302. [38] Ignarro LJ. Nitric oxide: biology and pathobiology. Academic Pr; 2000. [39] Hughes MN. Chemistry of nitric oxide and related species. Methods Enzymol 2008; 436:3-19. [40] Stamler JS, Feelisch M. Methods in Nitric Oxide Research. Wiley; 1996.

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[41] Wang PG, Xian M, Tang X et al. Nitric oxide donors: chemical activities and biological applications. Chem Rev 2002; 102(4):1091-1134. [42] Keefer LK, Nims RW, Davies KM, Wink DA. " NONOates"(1-Substituted Diazen1-ium-1, 2-dio-lates) as Nitric Oxide Donors: Convenient Nitric Oxide Dosage Forms. Methods Enzymol 1996;281-292. [43] Li Q, Lancaster Jr JR. Calibration of nitric oxide flux generation from diazeniumdiolate NO donors. Nitric Oxide 2009; 21(1):69-75. [44] Christodoulou D, Kudo S, Cook JA et al. Electrochemical methods for detection of nitric oxide. Methods Enzymol 1996; 268:69-83. [45] Murphy ME, Noack E. [24] Nitric oxide assay using hemoglobin method. Methods Enzymol 1994; 233:240-250. [46] Kojima H, Nakatsubo N, Kikuchi K et al. Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Anal Chem 1998; 70(13):24462453. [47] Sun J, Zhang X, Broderick M, Fein H. Measurement of nitric oxide production in biological systems by using Griess reaction assay. Sensors 2003; 3(8):276-284. [48] Rodriguez J, Specian V, Maloney R, Jourd'heuil D, Feelisch M. Performance of diamino fluorophores for the localization of sources and targets of nitric oxide. Free Radic Biol Med 2005; 38(3):356-368. [49] Augusto O, Bonini MG, Amanso AM, Linares E, Santos CC, De Menezes SL. Nitrogen dioxide and carbonate radical anion: two emerging radicals in biology. Free Radic Biol Med 2002; 32(9):841-859. [50] Wardman P. Methods to measure the reactivity of peroxynitrite-derived oxidants toward reduced fluoresceins and rhodamines. Methods Enzymol 2008; 441:261-282. [51] Neta P, Huie RE, Ross AB. Rate constants for reactions of inorganic radicals in aqueous solution. J Phys Chem Ref Data 1988; 17(3):1027-1284. [52] Burner U, Furtmuller PG, Kettle AJ, Koppenol WH, Obinger C. Mechanism of reaction of myeloperoxidase with nitrite. J Biol Chem 2000; 275(27):20597. [53] Goldstein S, Lind J, Merenyi G. Chemistry of peroxynitrites as compared to peroxynitrates. Chem Rev 2005; 105(6):2457-2470. [54] Folkes LK, Patel KB, Wardman P, Wrona M. Kinetics of reaction of nitrogen dioxide with dihydrorhodamine and the reaction of the dihydrorhodamine radical with oxygen: Implications for quantifying peroxynitrite formation in cells. Arch Biochem Biophys 2009; 484(2):122-126. [55] Wrona M, Wardman P. Properties of the radical intermediate obtained on oxidation of 2', 7'-dichlorodihydrofluorescein, a probe for oxidative stress. Free Radic Biol Med 2006; 41(4):657-667. [56] Cope VW, Chen SN, Hoffman MZ. Intermediates in the photochemistry of of carbonato-amine complexes of cobalt (III). Carbonate (-) radicals and the aquocarbonato complex. J Am Chem Soc 1973; 95(10):3116-3121. [57] Chen SN, Cope VW, Hoffman MZ. Behavior of carbon trioxide (-) radicals generated in the flash photolysis of carbonatoamine complexes of cobalt (III) in aqueous solution. The J Phys Chem 1973; 77(9):1111-1116. [58] Richardson DE, Yao H, Frank KM, Bennett DA. Equilibria, kinetics, and mechanism in the bicarbonate activation of hydrogen peroxide: oxidation of sulfides by peroxymonocarbonate. J Am Chem Soc 2000; 122(8):1729-1739.

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[59] Ferrer-Sueta G, Radi R. Chemical biology of peroxynitrite: kinetics, diffusion, and radicals. ACS Chemical Biology 2009; 4(3):161-177. [60] Alvarez MN, Peluffo G, Folkes L, Wardman P, Radi R. Reaction of the carbonate radical with the spin-trap 5,5-dimethyl-1-pyrroline-N-oxide in chemical and cellular systems: pulse radiolysis, electron paramagnetic resonance, and kinetic-competition studies. Free Radic Biol Med 2007; 43(11):1523-1533. [61] Winterbourn CC. Toxicity of iron and hydrogen peroxide: the Fenton reaction. Toxicol Letters 1995; 82:969-974. [62] Goldstein S, Meyerstein D, Czapski G. The fenton reagents. Free Radic Biol Med 1993; 15(4):435-445. [63] Halliwell B, Gutteridge J. Biologically relevant metal ion-dependent hydroxyl radical generation. An update. FEBS Letters 1992; 307(1):108-112. [64] Dawn-Linsley M, Ekinci FJ, Ortiz D, Rogers E, Shea TB. Monitoring thiobarbituric acid-reactive substances (TBARs) as an assay for oxidative damage in neuronal cultures and central nervous system. J Neurosci Methods 2005; 141(2):219-222. [65] Neta P, Huie RE, Ross AB. Rate constants for reactions of peroxyl radicals in fluid solutions. J Phys Chem Ref Data 1990; 19:413. [66] Zielonka J, Marcinek A, Adamus J, Gębicki J. Direct observation of NADH radical cation generated in reactions with one-electron oxidants. J Phys Chem A 2003; 107(46):9860-9864. [67] Bothe E, Schuchmann MN, Schulte-Frohlinde D, Von Sonntag C. HO2 Elimination from -hydroxyalkylperoxyl radicals in aqueous solution. Photochem Photobiol 1978; 28(4-5):639-643. [68] Von Sonntag C, Dowideit P, Fang X et al. The fate of peroxyl radicals in aqueous solution. Water Science and Technology 1997; 35(4):9-15. [69] Shen X, Lind J, Eriksen TE, Merenyi G. Reactivity of the trichloromethylperoxo radical: Evidence for a first-order transformation. J Phys Chem 1989; 93(2):553-557. [70] Alfassi ZB, Khaikin GI, Neta P. Arylperoxyl radicals. Formation, absorption spectra, and reactivity in aqueous alcohol solutions. J Phys Chem 1995; 99(1):265-268. [71] Fang X, Mertens R, Sonntag C. Pulse radiolysis of aryl bromides in aqueous solutions: Some properties of aryl and arylperoxyl radicals. J Chem Soc, Perkin Transactions 2 1995; 1995(6):1033-1036. [72] Niki E. [3] Free radical initiators as source of water-or lipid-soluble peroxyl radicals. Methods Enzymol 1990; 186:100-108. [73] Niki E, Yoshida Y, Saito Y, Noguchi N. Lipid peroxidation: mechanisms, inhibition, and biological effects. Biochem Biophys Res Comm 2005; 338(1):668-676. [74] Akasaka K, Suzuki T, Ohrui H, Meguro H. Study on Aromatic Phosphines for Novel Fluorometry of Hydroperoxides (II)-The Determination of Lipid Hydroperoxides with Diphenyl-1-Pyrenylphosphine. Analytical Letters 1987; 20(5):797-807. [75] Okimoto Y, Watanabe A, Niki E, Yamashita T, Noguchi N. A novel fluorescent probe diphenyl-1-pyrenylphosphine to follow lipid peroxidation in cell membranes. FEBS Letters 2000; 474(2-3):137-140. [76] Wrona M, Patel KB, Wardman P. The roles of thiol-derived radicals in the use of 2', 7'-dichlorodihydrofluorescein as a probe for oxidative stress. Free Radic Biol Med 2008; 44(1):56-62.

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[77] Marchesi E, Rota C, Fann YC, Chignell CF, Mason RP. Photoreduction of the fluorescent dye 2'-7'-dichlorofluorescein: a spin trapping and direct electron spin resonance study with implications for oxidative stress measurements. Free Radic Biol Med 1999; 26(1-2):148-161.

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Chapter 11

Enzymatic Generation of Hypoxia and Steady-state H2O2 Gabi N. Waite1, Gunda Millonig2, Lee R. Waite3, Helmut K. Seitz2 and Sebastian Mueller2,* 1

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Department of Cellular and Integrative Physiology, Indiana University School of Medicine, Terre Haute, IN 47809, U.S. 2 Department of Medicine and Center for Alcohol Research, Liver Disease and Nutrition, Salem Medical Center, University of Heidelberg, Zeppelinstraße 11 – 33 69121 Heidelberg, Germany 3 Department of Applied Biology and Biomedical Engineering, Rose-Hulman Institute of Technology, Terre Haute, IN 47803, U.S.

1. Introduction The objective of this Chapter is to present a flexible enzymatic system to study the in vitro effects of hypoxia and H2O2 on cells. Such studies aim at better understanding a wide variety of inflammatory diseases such as rheumatoid arthritis, heart disease or diabetes. In addition, they are important for all pathologies that involve redox reactions under hypoxic conditions, e.g. as occurs in all cancer tissues. The presented cell culture system can independently control O2 and H2O2 concentrations; in section 2 we focus on applications for generating hypoxia, while section 3 describes how to generate steady state concentrations of H2O2. Each section briefly introduces the biological and clinical relevance of O2/H2O2 studies, followed by the presentation of the enzymatic system and recommendations for its use. The system can also be employed to investigate O2 and H2O2 in combination. A few examples will be provided in section 4. The reader is referred to references 12 and 23 for more detailed information and protocols.

*

Corresponding author. Email: [email protected]

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2. Control of Oxygen Concentration 2.1. The Role of Hypoxia in Normal and Pathological Conditions

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2.1.1. Normoxia and Hypoxia Oxygen is essential for animal life. Hence, virtually all animal cells sense O2 levels and respond to O2 concentrations that differ from normal. Normal O2 concentrations, or “normoxic conditions”, of mammalian cells in the body typically range between 2 and 9% O2, which is significantly lower than the 21% O2 of ambient air at sea level. Nine percent O2 in this case means an O2 concentration that is in equilibrium with a partial pressure of 760 x 0.09 = 68 mm Hg. The highest O2 concentration detected in the human body is around 16% in the lung alveoli. On the other hand, many body cells can exist normally at O2 conditions as low as 1% O2, amongst them cells from the thymus, the kidney medulla, the liver, and the bone marrow (Figure 1). Normoxic conditions fluctuate with time. For instance, during the development of the embryo in the mother‟s womb, the norm is hypoxia when compared to the O2 requirement of the corresponding cells after birth. In fact, without hypoxia, embryonic organs do not develop normally [1]. Another example is the normal change in O2 conditions during the lifetime of adult cells. White blood cells typically experience a dramatic drop in O2 pressure when leaving the blood vessel and entering tissue. On the other hand, some cells, among them heart and brain cells, have a consistently high O2 requirement to function normally and cannot easily adapt to changes. Even transient hypoxia can cause enormous damage as evidenced by irreversible brain injury after cardiac resuscitation [2].

Figure 1. Partial O2 pressures (in percent or mm Hg) normally vary in human tissues, from partial pressures in equilibrium with 21% O2 for tissues in direct contact with the ambient air, to partial pressure in equilibrium with 16% in lung alveoli and 1% in organs such as the liver. Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

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2.1.2. The Hypoxia Sensor HIF How cells actually sense hypoxia and control their oxygen supply has been a subject of continuous debate for many years. One strong candidate among others is the transcription factor hypoxia-inducible factor 1 (HIF-1) which is stabilized under low oxygen conditions and orchestrates the activation of numerous genes [3] (for details see Vol. II, Chapter 22). HIF-1 is only found in multicellular organisms and consists of two subunits, HIF-1 and HIF-1 is degraded by proteasomes. The initial trigger for degradation is its hydroxylation by HIF prolyl hydroxylases, which require O2 to create an –OH group on proline residues. Hence, in the presence of low or no oxygen, the lack of hydroxylation stabilizes HIF-1 so that it can bind, together with HIFelements in DNA. This activates the expression of a wide range of genes whose protein products can lead to protective and adaptive responses to hypoxia. These include systemic changes, such as increased formation of erythrocytes and blood vessels, and cellular changes, such as shifts to the much less efficient anaerobic energy metabolism. Pharmacological stabilization of HIF-1 offers an attractive approach to protect heart and brain cells from hypoxia and to provide a potentially therapeutic benefit in the case of myocardial infarction or stroke [4]. On the other hand, enhanced degradation of HIF-1 has become a hot topic in cancer research. Solid tumors have a low supply of oxygen so that HIF1 is over-expressed in many cancers, such as brain or breast tumors. Moreover, mutations of key oncogenes, as well as free radical formation as a consequence of tumor hypoxia or immune cell infiltration, further increase HIF activity. Tumor cells that survive radiation and chemotherapy show even higher HIF activity than the untreated cancer cells due to their successful adaptation to hypoxia. Hence, the inhibition of HIF activity is believed to confer therapeutic benefit in cancer [5].

2.2. Oxygen Conditions In Vitro The previous section briefly introduced the variability of O2 concentration in tissues and the protective cellular and physiological responses implemented when O2 levels fall. Thus, normoxia and hypoxia studies may contribute substantially to the advent of effective therapies targeting a wide range of afflictions including cancer and cardiovascular disease. However, the majority of in vitro studies in these areas are performed using classic cell culture techniques wherein the surface of the cell media forms an interface with ambient air. The 21% oxygen in ambient air reaches equilibrium with the media, limited by its solubility in water (about 220 x10-6 M at the air media interface). Thus, classic routine cell culture methods, developed over decades and employed all around the world, almost always simulate hyperoxic conditions as compared to the conditions that cells experience inside the body, in health or disease. Several indirect methods are available to investigate hypoxia-related question in vitro. For instance, it is possible to chemically induce HIF-1 by exposure to cobalt chloride, or to block mitochondrial respiration by cyanide poisoning. Most often, to lower directly the partial oxygen pressure, cell culture plates are placed in gas impermeable containers, in which ambient air is replaced by a defined, low oxygen gas mixture [6]. This method has several

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obvious limitations. First, once inside the airtight container, the cells cannot be manipulated or microscopically inspected. Second, if factory mixed, pre-analyzed gas tanks are used, for each tested O2 condition a different tank must be purchased. To circumvent these limitations, sophisticated and expensive incubators with air locks are required. From a practical perspective, perhaps the most serious limitation is the fact that it takes many hours until the cells are exposed to the final O2 concentration; 2-24 hours, depending on the system [7]. This limits the use of such hypoxic chambers to mimic slow body O2 changes.

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2.3. The Enzymatic GOX/CAT System to Study Hypoxia In Vitro 2.3.1. Overview It is the objective of this Chapter to introduce an enzymatic hypoxia research system, called the “GOX/CAT system”. It is inexpensive, flexible and can rapidly create a wide range of O2 conditions. The GOX/CAT system uses two well known enzymes, glucose-1-oxidase (GOX) [8] and catalase (Cat) [9]. GOX and CAT are added to buffered solutions containing at least 5 mM D-glucose. With glucose present, GOX consumes O2 to produce H2O2, and CAT removes H2O2 while replenishing half of the consumed oxygen (Figure 2). Thus, the overall reaction consumes O2, the prerequisite for generating hypoxia. From the overall stoichiometry of the GOX/CAT system, it becomes evident that, apart from depleting O2, varying the ratios of GOX and CAT can also control H2O2 levels. This allows one to study the impact of hypoxia in the presence or absence of physiological steadystate H2O2 concentrations, better mimicking inflammatory or ischemic conditions. Further details are provided in section 3 below. Here, we concentrate on the control of hypoxia. To create hypoxia for cultured cells, commercially available purified GOX and CAT are added to the medium of regular culture vessels that contain cells at the bottom of the dishes (Figure 3). As regular culture vessels have an air-medium interphase, the degree of hypoxia that the cells are exposed to depends mainly on two factors: i) the O2 diffusion distance (or the volume of the medium), and ii) the specific GOX activity KGOX (or the amount of GOX added to the medium). First, let us address the GOX activity.

Figure 2. The stoichiometry of the GOX/CAT system indicates that GOX consumes O2 to produce H2O2, and CAT removes H2O2 while partly replenishing half of the O2. By varying the GOX and CAT activities and the culture conditions, steady-state (ss) O2 and H2O2 concentrations can be maintained independently.

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Figure 3. The GOX/CAT hypoxia system can be used in regular culture vessels with cells at the bottom of the vessel and access to 21% O2 of ambient air at the top. Hypoxia at the cell level develops under conditions where GOX-dependent O2 consumption in medium (corrected by the CAT activity) is larger than its replacement from air by diffusion.

Figure 4. The modeled and measured O2 partial pressures of the GOX/CAT system are in good agreement. Higher GOX concentration leads to greater hypoxia and faster O2 steady-state (A, B), and the same can be accomplished by increasing the height of the media column above the cells (C, D). Principles of Free Radical Biomedicine, Nova Science Publishers, Incorporated, 2011. ProQuest Ebook Central,

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2.3.2. Impact of GOX Activity GOX is naturally present in some fungi and bacteria and in the hypopharyngeal gland of the honey bee, but absent from other animal cells, which remain unresponsive to the addition of GOX to the medium. GOX is stable at 37 C and works well at physiological pH 7.4 (although its optimal activity is between pH 4.5 and 6). GOX is highly substrate-specific and at the typical D-glucose concentrations of cell culture media (5-25 mM), it operates under near-saturated conditions (KM of 9.8 mM) [10], allowing an almost constant turnover independently of glucose levels. As a side note, the accumulation of gluconolactone can acidify the medium, which should be avoided when using the system. With temperature, pH, and salt concentration constant and with the substrate glucose present in excess, O2 consumption is proportional to the GOX enzyme activity. Simply adding more enzyme to the medium will lead to stronger hypoxia. However, the developing hypoxia will inhibit the enzyme activity because O2 is the second substrate of GOX, and an enzyme only works maximally with unlimited substrate. Hence, to predict the O2 environment of the cultured cells, KGOX is multiplied by the ratio of the O2 concentration at the depths of the cells and the O2 concentration at the air-medium interface. Using a luminol/hypochlorite dependent H2O2 assay [11] (see also Chapter 12), we determined the KGOX of Aspergillus Niger as 4.5 x10-2 mol/L/s, at 37 C and 25 mM glucose. For the O2 concentration at the air-media interface, we used 220 x 10-6 moles/ L, the O2 concentration in saturated water at 37 ºC and atmospheric pressure. 2.3.3. Impact of O2 Diffusion Next, we want to predict the natural O2 concentration at the bottom of the well. Because it depends on oxygen consumption and on O2 diffusion, we used Fick‟s law to calculate the diffusion. This introduces the second factor besides KGOX that affects hypoxia in our system, the media volume. The higher the column of medium above the cells, the longer it takes O2 from ambient air to reach the cells by diffusion. Calculations of actual resulting hypoxia conditions have been been done exemplarily using Excel [12] and by solving the partial differential equations in Matlab [13]. Figure 4 demonstrates the impact of GOX concentration and diffusion distance on the development of hypoxia at the bottom of cell culture dishes. Oxygen concentrations are given as dissolved oxygen, with 220 μM O2 corresponding to 21% ambient O2. Data on the left are from the Matlab model; data on the right are obtained with an oxygen electrode. In the top set of modeled/measured data (Figures 4A and B), it can be seen that higher GOX concentrations lead to greater hypoxia. In the lower set of modeled/measured data (Figures 4 C and D), a longer diffusion distance results in increased hypoxia. Figure 4 includes another layer of information, the time to reach steady-state hypoxia: The higher the GOX concentration, the faster the steady-state O2 concentrations are reached. Figure 5 shows that steady-state hypoxia can also be reached quickly by using a shallow fluid layer above the cells. In a well of a 12-well culture plate, stable hypoxia is reached within 10 minutes at a medium height of 2 mm, compared to 40 minutes with 5.6 mm liquid height. Consequently, the 2 mm setting would be chosen for the design of experiments with fast hypoxia/re-oxygenation cycles, while the 5.6 mm setting would be more appropriate for longterm experiments, aimed at minimizing GOX substrate depletion and product accumulation.

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Figure 5. In the GOX/CAT system, a combination of high GOX activity and large volume of medium can be used to induce hypoxia within minutes. This system is ideal for fast hypoxia/re-oxygenation cycles, since it cannot be maintained for long due to the conversion of glucose to acidic gluconolactone.

2.3.4. Impact of CAT Activity In our discussion thus far, we neglected CAT which also influences oxygen flux as seen in Figure 2. Unlike most other enzymes, CAT cannot be saturated up to millimolar concentrations (see also Vol. II, Chapter 7), concentrations that far exceed cell toxic levels. Thus, at all experimental conditions, CAT degrades H2O2 to O2 in a process governed by first order kinetics, with the rate of H2O2 breakdown to O2, proportional to the H2O2 concentration. As a result of this breakdown, CAT always replaces one half of the O2 that is used up by GOX. This O2 source is important to consider in any modeling, but it is not necessary for the principle understanding of the GOX/CAT system as an oxygen-consuming, hypoxia-generating system.

2.4. Getting Started with the GOX/CAT Hypoxia System In the GOX/CAT system, hypoxia at the cell level develops under conditions where GOXdependent O2 consumption in media is larger than oxygen replacement from air by diffusion. The system‟s advantage is that it is relatively easy to use. Furthermore, it is affordable and can be used with almost all cell cultures and conditions. This allows investigators to inexpensively test whether a cellular response under study depends on the partial pressure of O2. The disadvantage of the system is that the O2 gradient across the culture vessel is precarious, so that slight disturbances may lead to short term re-oxygenations. Hence, the system is not applicable for studies in which the cells must be totally shielded from ambient air. Examples of GOX and CAT combinations that lead to O2 concentrations corresponding to 18-0.5% O2 at the bottom of a well of a 12-well plate, filled with 1 ml medium are given in more detail elsewhere [12]. The information can be adapted to other culture conditions with

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the use of tables that relate culture media volumes to O2 diffusion between air and the bottom of the wells. The Appendix of this article contains interactive Excel tables to facilitate these calculations. The Appendix furthermore contains example protocols that explain in a stepwise fashion the procedure on how to expose cells to hypoxia. To allow for more flexibility in adapting the GOX/CAT system to a particular in vitro situation, we modeled the system in Matlab [13]. This program permits prediction of the development of O2 and H2O2 gradients for any GOX and CAT activity and for any experimental setting which may vary as per media volume, culture vessel growth area and cellular O2 consumption. We further solved the inverse problem, thus predicting the necessary enzyme activities to establish experimentally the desired O2/ H2O2 concentrations.

3. Control of H2O2 Concentration 3.1. The Role of H2O2 in Normal and Pathological Conditions

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Like oxygen, H2O2 has both beneficial and deleterious effects in the body. At cell concentrations of about 10 M and above, H2O2 becomes toxic for most cell types. All cells can sense these concentrations and respond with an increased activity of the enzymes that eliminate the molecule and thus protect the organism from this type of oxidative stress [14, 15]. On the other hand, at cell concentrations of about 10 M and below, H2O2 can modify the activity of redox-sensitive proteins. The altered proteins might directly act as signaling messengers or indirectly change the cells‟ redox potential, which in both cases can influence numerous cellular actions [16, 17]. Let us first consider H2O2 as a toxin. 3.1.1. H2O2 as Toxin H2O2 has been established as important cellular toxin for many decades and numerous defense systems have been identified across the phylogenetic spectrum that protect from H2O2 toxicity, such as catalases, peroxidases, glutaredoxins and peroxiredoxins (Vol. II, Chapters 7-10). Life on our planet had to cope from the very beginning with oxygen-derived H2O2, which was generated through cosmic irradiation and UV light exposure (Chapter 1). The concomitant presence of increasing ambient oxygen concentrations and transition metals (mostly iron) was particularly toxic (Chapter 5). H2O2 itself is relatively stable and weakly reactive. Protected from light and in the absence of catalysts (e.g. in pure deionized water), H2O2 stock solutions can be stably maintained for many years. However, in the presence of minuscule amounts of iron, H2O2 is readily degraded leading to one of the most reactive and damaging oxygen species, the hydroxyl radical. Hydroxyl radicals are typically generated by the Fenton and Haber-Weiss reactions (Chapter 5) that essentially involve H2O2 (Figure 6). Commercially, this oxidizing reactivity is used to bleach hair and textiles, and clinically, it is used to kill microorganisms. On the other hand, H2O2 is continuously produced as a byproduct of normal oxidative metabolism. H2O2 is further produced by some biosynthetic (e.g. thyroid hormone synthesis) or degradation (e.g. liver cytochrome P450 detoxification) pathways. In addition, mammalian cells contain peroxisomes (Vol. II, Chapter 17) that harbor more than 20 H2O2-producing oxidases. Another important class of H2O2-producing enzymes includes the NADPH-

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dependent oxidases, now classified as NOX or DUOX family members. Their most widely known representative is NOX2, the H2O2-producing enzyme of inflammatory cells that directly kills bacteria in concert with other enzymes, such as myeloperoxidase. Under healthy conditions, the rates of H2O2 production and clearance are normally well balanced. The balance can be disturbed by excessive H2O2 production (e.g. due to chronic inflammatory activity), by unexpected H2O2 sources (e.g. from cigarette smoke), or by diminished H2O2 clearance (e.g. decline in the efficiency of antioxidant mechanisms with advancing age). Local H2O2 concentrations can reach toxic levels due to its production by white blood cells as a first line of defense against microorganisms. Especially during inflammation (Vol. II, Chapter 21), excessive quantities of H2O2 are produced, leading to destruction of body tissues and disease,. Chronic inflammation contributes to a large array of human illnesses, including atherosclerosis, Alzheimer disease, and certain cancers. 3.1.2. H2O2 as Beneficial Cell Regulator Although H2O2 contributes to oxidative stress, it can also have a positive impact on cells. With the evolution of organisms with increasing complexity, the number of H2O2 /redox sensing proteins adjusted accordingly as a defence against increasing burdens of oxidative and metabolic stress. The activity of many cellular proteins came under regulation by oxidative stress including transcription factors such as activator protein 1 (AP-1; Vol. II, Chapter 14) and nuclear factor-kappa B (NF-κB, Vol. II, Chapter 12). Other intracellular targets of H2O2 were the protein tyrosine phosphatases (PTP); binding of H2O2 promotes their inhibition which shifts the cells‟ phosphate status, thereby influencing certain growth factor and hormone signaling pathways. Although the significance of H2O2 as a cell signaling molecule has achieved wide acceptance, many questions remain unanswered. For instance, it is not clear whether the regulatory function of H2O2 is restricted to certain cells, especially those involved in inflammatory events, or whether H2O2 plays a more ubiquitous role akin to nitric oxide and calcium. Although it was initially believed that H2O2 can freely pass through biological membranes, the existence of H2O2 gradients across membranes has been documented [18]. Due to its stability and diffusion properties, leading to a rather large “mean displacement constant”, H2O2 can reach distant locations within the cells or between cells. The exact molecular actions of H2O2 are still poorly understood and debatable [19, 20]. From a therapeutic perspective, low concentrations of H2O2 may have salutary effects on cellular function not unlike the benefits conferred by NO [21].

3.2. H2O2 Conditions In Vitro To ascertain the medical potential of H2O2, it is necessary to understand better how cells discriminate between beneficial and harmful H2O2 effects. Many of these studies require exposing cells to a wide variety of H2O2 concentrations in vitro, while simulating inflammatory tissue conditions such as varying O2 concentrations.

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Figure 6. Hydrogen peroxide, formed during the reduction of oxygen to water, is not very toxic by itself, but may give rise to the extremely toxic hydroxyl or hydroperoxyl radicals. Cat: Catalase, GSH-PX: glutathione peroxidase, Prx: peroxiredoxin, SOD: superoxide dismutase.

We already mentioned the limitations in the ability of many hypoxia chambers to manipulate the cells once they are inside the chamber, for instance by H2O2 bolus administration. Even under atmospheric O2 pressures, adding H2O2 as bolus is less than optimal [22]. To expose cell proteins to meaningful H2O2 concentrations of about 1 to 20 M, one has to typically add 5 to 50-fold higher concentrations to the medium. The effects of these artificially high concentrations on cell membrane proteins are largely unknown. The second drawback of using H2O2 boluses is their limited lifetime (few minutes) under cell culture conditions, which prevents studies of long-term H2O2 exposure.

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Figure 7. By monitoring H2O2 in real-time, it is evident that H2O2 decreases after addition of CAT and is generated upon addition of GOX. In the presence of GOX and CAT, a steady-state level of H2O2 ( 0.5 µM) is maintained.

Figure 8. In the GOX/CAT model, the absolute H2O2 levels are similar when the ratio between GOX and CAT is maintained, as indicated by the dashed line of the modeled data. The time to reach steady-state is controlled by the GOX concentration and is faster at higher GOX concentrations.

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3.3. Generation of Steady-state H2O2 by the GOX/CAT System

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The GOX/CAT system offers the possibility of exposing cells for 24 hours or longer to steady-state H2O2 concentrations. Steady-state is reached when the rates of H2O2 production and H2O2 removal are equal. The H2O2 production rate depends on the GOX activity, at conditions when O2, glucose and gluconolactone are not limiting. We have already discussed this in the context of O2 removal by GOX, but it holds equally true for the production of H2O2 due to the 1:1 stoichiometry for the conversion of O2 to H2O2 (Figure 2). Hence, for the mathematical model of the system, we modeled the O2 flux as described above in Section 1.3 as a basis for developing the H2O2 flux. The H2O2 removal rate depends on the CAT activity, which is proportional to the amount of H2O2, up to millimolar concentrations. This first order kinetics of CAT is the prerequisite for the formation of steady-state levels because it means that at any time the H2O2 removal rate is proportional to the H2O2 production rate. The luminol-hypochlorite assay depicted in Figure 7 shows the logarithmic decrease of H2O2 after addition of CAT, the production of H2O2 by GOX, and the formation of steady-state H2O2 in the presence of both enzymes. Not shown is the fact that this steady-state can be maintained for as long as glucose is not falling below levels affecting GOX activity and for as long as the accumulating gluconolactone is not acidifying the medium. To avoid the latter, in typical experiments, the medium should be exchanged about once every 24 hours [12]. The steady-state H2O2 concentration is determined by the ratio of GOX and CAT activities. This is demonstrated in Figure 8, which shows plots of H2O2 concentrations over time at the bottom of a well of a 96-well culture plate, filled with 0.2 ml liquid. It can be seen that for a given GOX concentration, the H2O2 steady-state level will solely depend on the activity of CAT. Steady-state H2O2 concentration of approximately 7 M H2O2 will be obtained when combining 1X GOX and 0.1X CAT.

3.4. Getting Started with the H2O2 Steady-state System Recommendations on how to apply the steady-state H2O2 generating GOX/CAT system to the individual experimental set-up are similar to those presented above for its use in a hypoxia system. First, consult the recently published review article for practical tips and example protocols [12]. Examples of GOX and CAT dilutions from stock are given to obtain 1, 5 and 10 µM steady-state H2O2 concentrations at the bottom of a 12-well plate, filled with 1 ml medium. As for hypoxia studies, bench protocols and tables facilitate adaptation to conditions other than 12-well plates. The second recommendation is to use our models [12, 13]. Using the mathematical model of O2 and H2O2 flux, we are able to create programs in which values of O2 and H2O2 concentrations are stored in vectors as a function of time and well depth. This allows the creation of graphs using any of the system‟s parameters. For instance, Figure 9 relates O2 to H2O2 concentrations at three different GOX/CAT combinations. From these graphs, it can be seen that H2O2 equilibrium will always be reached first (triangles), before O2 concentration starts to decrease substantially towards steady-state (filled circles). It indicates that hypoxic

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cells will only be exposed to the final H2O2 equilibrium value, and not to a range of H2O2 concentrations. In most cases, the cells‟ ability to degrade extracellular H2O2 can be neglected for the use of the GOX/CAT system. However, taking the cells‟ activity into account for the overall CAT activity will increase the accuracy of the system. On average, 105 cells are able to degrade extracellular H2O2 with a KCAT of 5 x 10-4/s, which can be increased up to 2.5 fold for cells implicated in redox metabolism, such as macrophages. Last, the accompanying Chapter 12 of this book introduces a highly sensitive H2O2 detection assay based on the H2O2–dependent oxidation of luminol by sodium hypochlorite. This assay is not necessary for the GOX/CAT system, but it is very helpful for validation of the system under individual experimental culture conditions, as it can be used to determine the enzyme activities of GOX and CAT and the steady-state H2O2 concentration.

4. Example Results of the GOX/CAT System

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4.1. Macrophage Activities at Low O2 and H2O2 Macrophages are white blood cells that derive from circulating monocytes when they reach sites of infection. There, one major task is to help destroy microorganisms and to eliminate cell debris. As part of this lifecycle, macrophages might encounter a wide range of O2 concentrations, from roughly 13% O2 in arterial blood to less than 1% O2 in infected tissue. A second major task of macrophages is the use of H2O2 as a bactericidal defense chemical, a phenomenon known as the oxidative burst. As part of the oxidative burst, molecular O2 is enzymatically reduced to generate superoxide, which spontaneously, or enzymatically, converts to H2O2. The GOX/CAT system can be employed to study the behavior of macrophages under various O2 and H2O2 conditions. Figure 10A shows that THP-1 macrophages stimulated with PMA for 48 h tolerate hypoxia down to 10%, but beyond that they respond with a change in metabolism. This is reflected in their decreased capacity to reduce the dye MTT to a spectroscopically quantifiable chemical. While the decrease in cellular reducing activity does not specify the relevant biochemical reactions, it indicates that macrophages markedly change their behavior during hypoxia. This is necessary because they face the daunting task of increasing their energy-consuming activities, phagocytosis and oxidative burst, under hypoxic conditions when aerobic energy production becomes limited. Figure 10B shows that macrophages are well adapted to functioning under hypoxic conditions and even increase phagocytosis of E. coli particles with decreasing O2 concentrations. These data imply that low oxygen itself is a strong macrophage activator, in line with reported hypoxia-dependent changes in gene expression profiles [23, 24].

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Figure 9. Oxygen versus H2O2 plots at three different GOX/CAT combinations show similar kinetics because H2O2 steady-state (triangles) is always reached before O2 concentrations start to drop significantly towards O2 steady-state (circles).

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4.2. A New Role for HIF? The HIF system as biological regulator of hypoxia has been introduced above. The system includes prolyl-hydroxylases (PHDs) that target HIF-1 for destruction under normoxia and allow its stabilization under hypoxia. Stable HIF activates a wide variety of hypoxic stress responses, amongst them metabolic adjustments to hypoxia. In experiments applying the GOX/CAT method, we showed that the HIF system responds to oxygen changes, not hypoxia per se, and also to oxidative stress [25].

Figure 10. Macrophage behavior under infectious/ inflammatory conditions, simulated with the GOX/CAT hypoxia/ steady-state H2O2 system. A: Cells alter their metabolism at O2 pressures below 10% (OD 560 indicates the absorbance of reduced MTT). B: Cells respond to hypoxia with increased phagocytosis.

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Figure 11. Insight into the HIF/PHD system using the GOX/CAT hypoxia/steady-state H2O2 system. A: During hypoxia HIF1- is transiently increased, but disappears again after the upregulation of PHD2, implicating an HIF/PHD loop. B: In the presence of H2O2, HIF-1 remains increased after 12 h of steady hypoxia, indicative of HIF/PDH loop disruption (right panel).

A few pieces of evidence to support these views are presented here. Figure 11A shows HIF-1 expression as detected by Western blots in Huh-7 hepatoma cells exposed to hypoxia by the GOX/CAT system. HIF-1 is upregulated within minutes of hypoxia, but disappears again after the upregulation of PHD2. This indicates that PHD2 initiated the degradation of HIF-1 despite maintained hypoxia. The explanation is that PHD2 and PHD3 themselves are target genes of HIF [26] so that the picture of a PHD/HIF feedback loop emerges. Hypoxia transiently decreases PHD activity (which increases HIF), but PHD activity increases after HIF-induced de novo synthesis of PHD (which decreases HIF). This loop could resolve the long-standing conundrum why HIF is not present in tissues with stable and low O2 pressures. In other studies, not discussed in detail here, we show that the loop starts when the O2 concentration falls below the pre-existing O2 concentration, as indicated by transiently increased HIF-1 and by the disappearance of HIF-1 as soon as the new situation stabilizes. This indicates that the HIF/PHD loop is designed to compensate for fast tissue fluctuations of O2 rather than to reflect the existing O2 conditions. Furthermore, by using the GOX/CAT system, we can show that oxygen-independent signals such as H2O2 can upregulate HIF-1 in a dose-response fashion up to 10 M H2O2.Figure 11B indicates that HIF-1 does not disappear as expected under stable hypoxia for 12 hours in the presence of H2O2. These data indicate that HIF-1 responds both to H2O2 and hypoxia. The GOX/CAT system is the first experimental setting that permits interrogation of the effect of both hypoxia and H2O2 independently, on a redox-sensitive transcription factor such as HIF.

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Conclusions Studies of the crucial cell regulators O2 and H2O2 have been hampered by the lack of appropriate experimental tools to control independently both molecules in cultured cells which would better simulate typical inflammatory conditions. This Chapter presents an enzymatic system that can generate physiologically relevant steady-state dissolved O2 and steady-state H2O2 concentrations at the bottom of cell culture vessels, independently and in combination. The system uses the enzymes glucose-1-oxidase (GOX) and catalase (CAT). GOX consumes O2 and glucose to produce H2O2, and CAT removes H2O2 while partly replenishing O2. The GOX activity and the O2 diffusion distance determine the concentrations of O2 in the culture medium. By adjusting the amount of GOX and the media volume, stable hypoxic levels of oxygen concentration between 0.01 and 21% can be generated. The main determinant for the steady-state concentrations of H2O2 is the ratio between the GOX and CAT activities. Biologically and clinically relevant H2O2 steadystate levels ranging between 1 and 20 M H2O2 can be maintained for hours or days. The GOX/CAT system can create hypoxia faster than a hypoxia chamber and it can create biologically relevant steady-state H2O2 concentrations for hours or days, without or with hypoxia. Thus, the GOX/CAT system is a powerful novel tool to study redox-sensitive signaling cascades and cellular functions under physiological low oxygen levels.

References [1]

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[2] [3] [4] [5] [6]

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Simon MC, Keith B. The role of oxygen availability in embryonic development and stem cell function. Nature Reviews Mol Cell Biol 2008; 9:285-196. Mueller PS. The Terri Schiavo saga: ethical and legal aspects and implications for clinicians [review]. Polskie Archiwum Medycyny Wewnetrznej 2009; 119(9):574-81. Yee Koh M, Spivak-Kroizman TR. Powis G. HIF-1 regulation: not so easy come, easy go. [Review] [85 refs]. Trends in Biochem Sci 2008; 33(11):526-34. Marx J. Cell biology. How cells endure low oxygen. Science 2004; 303(5663):14546. Poon E. Harris AL. Ashcroft M. Targeting the hypoxia-inducible factor (HIF) pathway in cancer. [Review]. Expert Reviews in Molecular Medicine 2009; 11:e26 Jenkins S. Making hypoxia happen. Just how do researchers produce low oxygen environments and measure tissue oxygen concentration. The Scientist 2002; 16(16): 44. Allen CB, Schneider BK, White CW. Limitations to oxygen diffusion and equilibration in in-vitro cell exposure systems in hyperoxia and hypoxia. Am J Physiol Lung Cell Mol Physiol 2001, 281(4): L1021-1027. Bankar SB. Bule MV. Singhal RS. Ananthanarayan L. Glucose oxidase--an overview. Biotechnology Advances 2009; 27(4):489-501. Zamocky M. Furtmuller PG. Obinger C. Evolution of catalases from bacteria to humans. Antioxidants & Redox Signaling 2008; 10(9):1527-48.

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[10] Mueller S, Weber A, Fritz R, Mütze S, Rost D, Walczak H, Völkl A, Stremmel W. Sensitive and real-time determination of H2O2 release from intact peroxisomes. Biochem J 2002; 1;363(Pt 3):483-91. [11] Mueller S. Sensitive and nonenzymatic measurement of hydrogen peroxide in biological systems. Free Radic Biol Med 2000 Sep; 29(5):410-5. [12] Mueller S, Millonig G, Waite GN. The GOX/CAT system: A novel enzymatic method to independently control hydrogen peroxide and hypoxia in cell culture. Adv Med Sci. 2009 Nov 27: 1-15. [13] Waite GN, Waite LR, Millonig G, Muller S. Model for in vitro control of H2O2 and O2 concentration in cell culture. Biomedical Engineering Society Annual Proceedings, October 2009; Pittsburgh, PA: PS 10A-143. [14] Rhee SG. Yang KS. Kang SW. Woo HA. Chang TS. Controlled elimination of intracellular H2O2: regulation of peroxiredoxin, catalase, and glutathione peroxidase via post-translational modification. Antioxidants & Redox Signaling 2005; 7(56):619-26. [15] Janssen-Heininger YMW. Mossman BT. Heintz NH. Forman HJ. Kalyanaraman B. Finkel T. Stamler JS. Rhee SG. van der Vliet A. Redox-based regulation of signal transduction: principles, pitfalls, and promises. Free Radical Biology & Medicine 2008; 45(1):1-17.Waite GN, Balcavage WX. From redox homeostasis to protein structure modulation and redox signaling therapy. Cell Science Reviews 2009; 5(3): 95-127. [16] Pedroso N. Matias AC. Cyrne L. Antunes F. Borges C. Malho R. de Almeida RF. Herrero E. Marinho HS. Modulation of plasma membrane lipid profile and microdomains by H2O2 in Saccharomyces cerevisiae. Free Radic Biol Med 2009; 46(2):289-98. [17] Carreras MC. Poderoso JJ. Mitochondrial nitric oxide in the signaling of cell integrated responses. American Journal of Physiology - Cell Physiology 2007; 292(5):C1569-80. [18] Abbas K. Breton J. Drapier JC. The interplay between nitric oxide and peroxiredoxins. Immunobiology 2008; 213(9-10):815-22. [19] Linnane AW. Kios M. Vitetta L. The essential requirement for superoxide radical and nitric oxide formation for normal physiological function and healthy aging. Mitochondrion 2007; 7(1-2):1-5. [20] Mueller S. Sensitive and nonenzymatic measurement of hydrogen peroxide in biological systems. In: Pryor WA, editor. Bio-assays for oxidative stress status (BOSS). New York: Elsevier; 2001. p. 170-5. [21] Elbarghati L. Murdoch C. Lewis CE. Effects of hypoxia on transcription factor expression in human monocytes and macrophages. Immunobiology 2008; 213(910):899-908. [22] Fang HY, Hughes R, Murdoch C, Coffelt SB, Biswas SK, Harris AL, Johnson RS, Imityaz HZ, Simon MC, Fredlund E, Greten FR, Rius J, Lewis C. Hypoxia-inducible factors 1 and 2 are important transcriptional effectors in primary macrophages experiencing hypoxia. Blood 2009; 114(4):844-59. [23] Millonig G, Hegeduesch S, Becker L, Seitz HK, Schuppan D, Mueller S. Hypoxiainducible factor 1 alpha under rapid enzymatic hypoxia: cells sense decrements of oxygen, but not hypoxia per se. Free Radic Biol Med 2009, 46(2), 182-191.

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[24] Metzen E. Stiehl DP. Doege K. Marxsen JH. Hellwig-Burgel T. Jelkmann W. Regulation of the prolyl hydroxylase domain protein 2 (phd2/egln-1) gene: identification of a functional hypoxia-responsive element. Biochem J 2005; 387(Pt 3):711-7.

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In: Principles of Free Radical Biomedicine. Volume 1 ISBN: 978-1-61209-773-2 Editors: K. Pantopoulos and H. M. Schipper © 2012 Nova Science Publishers, Inc.

Chapter 12

Chemiluminescence Detection of H2O2 Sebastian Mueller1,*, Gunda Millonig1 Helmut K. Seitz1 and Gabi N. Waite2 1

Department of Medicine and Center for Alcohol Research, Liver Disease and Nutrition, Salem Medical Center, University of Heidelberg, Zeppelinstraße 11 – 33 69121 Heidelberg, Germany 2 Department of Cellular and Integrative Physiology, Indiana University School of Medicine, Terre Haute, IN, U.S.

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1. Introduction The objective of this Chapter is to introduce the reader to an ultrasensitive chemoluminescence method for the detection of H2O2 in biological systems, based on luminol and hypochlorite. H2O2 is increasingly attracting attention not only as a potentially toxic cellular metabolite but also as an important signaling molecule comparable to calcium or nitric oxide. Many important basic cellular functions such as the cell cycle, programmed cell death (apoptosis), wound healing, inflammatory responses and others, unambiguously involve H2O2 as the central reactive oxygen species (ROS). Since H2O2 coexisted with living organisms early in our planet‟s history (Chapter 1), defense strategies against toxic H2O2 levels were developed that allow its rapid intracellular removal (within milliseconds). This extremely short half-live is responsible for the difficulties in studying H2O2 in biological systems. The hypochlorite/luminol assay allows very sensitive detection in the extracellular fluid, but is not able to detect intracellular H2O2. However, it is unsurpassed in terms of sensitivity and rapidity, which allows for real-time analysis of very low H2O2 levels. These facts and its cost-effectiveness make it an ideal exploratory tool to enter the area of H2O2 research and to learn more about this fascinating molecule. In addition, the luminol/hypochlorite assay has been instrumental in developing the glucose oxidase/catalase system that allows for the independent control of hypoxia and H2O2 in cultured cells (see *

Corresponding author. Email: [email protected]

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Chapter 11). This Chapter will first introduce the reader to functions of H2O2 in living organisms. It will then discuss conventional approaches and strategies to detect H2O2 under various conditions. Finally, the luminol/hypochlorite assay and some of its applications will be described in more detail.

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2. Physiology and Pathophysiology of H2O2 H2O2 is regarded as a major oxygen metabolite [1] (see also Chapter 2). Although not a radical by definition and rather stable compared to other oxygen metabolites, it is the source of other highly reactive oxygen species (ROS), e.g. the hydroxyl radical assumed to be generated in the Fenton and Fenton-like reactions by transition metal ions (Chapter 5). Due to the relatively high stability of H2O2 compared to other ROS, it can diffuse far from its origin with a mean displacement of 10-15 µm within the cell or in the extracellular space. This is one of the prerequisites for serving as a signaling molecule. H2O2 becomes very unstable and dangerous in the presence of transition metals, e.g. reduced iron, because it gives rise to toxic hydroxyl radicals via the Fenton reaction. Hydroxyl radicals react at nearly diffusion-controlled rates with lipids, carbohydrates, proteins and other biomolecules, thus, causing general damage to structural and functional components of cells and tissues. Patients with inherited disorders of copper and iron metabolism (Wilson‟s disease and hemochromatosis) have increased concentrations of these transition metals especially in their livers leading to liver fibrosis, cirrhosis and liver cancer [2, 3] (see also Vol. III, Chapters 2-4). Oxidative damage by H2O2 and H2O2-metabolites is not restricted to mammals but poses a threat to all pro- and eukaryotic cells including plants and bacteria. In mammalian organisms, white blood cells are one of the main sources of H2O2 release. Human polymorphonuclear neutrophils (PMN) and macrophages generate H2O2 during a respiratory burst of non-mitochondrial oxygen uptake. After stimulation, the membraneassociated NADPH-oxidase (now termed NOX2) reduces molecular oxygen to superoxide radicals (O2 -) [4 ]that dismutate to H2O2 either spontaneously or catalytically by superoxide dismutase (SOD; see Vol. II, Chapter 5). Hypochlorous acid (HOCl) is another toxic metabolite generated from H2O2 in neutrophils. In the seventies, myeloperoxidase was identified as a major chlorinating compound of neutrophils producing HOCl from H2O2 and chloride [5]. The oxidizing potential of HOCl, however, is intentionally used by leukocytes to kill bacteria as part of their immunological defense cascade. Hence, H2O2 serves under these conditions as a physiological substrate. In addition, H2O2 is an essential part of many biochemical pathways. It is enzymatically generated by oxidases, e.g. during fatty acid oxidation in peroxisomes [6] (see also Vol. II, Chapter 17), the monoamine oxidase of the outer membrane of mitochondria, membrane bound NADPH oxidases of the NOX and DUOX families, xanthine oxidase, and during the reduction of oxygen to water in the respiratory chain. H2O2 is also essential for the synthesis of some hormones, e.g. thyroid hormones, and melanin [1, 7-10].

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Figure 1. Major characteristics of an „ideal‟ H2O2 assay for cells and tissues.

Apart from its metabolic and toxic properties, H2O2 is increasingly being considered as a biological signaling molecule. In addition to inflammatory cells, a wide variety of normal and malignant cell types generate and release O2 - and H2O2 in vitro either in response to specific cytokine/growth factor stimuli or constitutively, as in the case of tumor cells. These ROS appear to act at submicromolar levels as novel intra- and intercellular "messengers", capable of promoting growth responses in culture. The mechanisms are thought to involve direct interaction with specific receptors or oxidation of growth signal transduction molecules such as protein kinases, protein phosphatases, transcription factors or transcriptional inhibitors. It is also possible that H2O2 modulates the redox state and activity of these important signal transduction proteins indirectly through changes in cellular levels of GSH and GSSG. More recently, direct posttranslational modifications of sulfhydryl residues mediated by still unknown peroxidases have been proposed [11]. Critical balances appear to exist in relation to cell proliferation on one hand and lipid peroxidation and cell death on the other. At low levels H2O2 promotes cell proliferation while in excess it leads to cell death [12-19]. In Escherichia coli, many genes are transcriptionally activated when challenged with H2O2 [20]. These responses are mediated by a central regulatory protein, OxyR, which stimulates transcription of many target genes when activated under oxidative stress. Finally, an interesting regulatory link between H2O2 and mammalian iron metabolism, which is mediated by iron regulatory protein 1 (IRP1), has been identified (Vol. II, Chapter 19). Sustained low levels of H2O2 are able to activate this central iron regulator. This finding is striking considering the role of H2O2 in iron toxicity (Fenton chemistry). Activation of IRP1 by H2O2 results in enhanced expression of transferrin receptor and increased cellular iron uptake from transferrin, while it also leads to a significant reduction in synthesis of the iron storage protein ferritin [21-26].

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Figure 2. Overview of various strategies to detect H2O2 in biological systems. The assays require either enzymatic or non-enzymatic reactions to oxidize substrates by H2O2 in a specific manner. Changes in fluorescence or luminescence by H2O2-dependent oxidation are then quantified.

H2O2 seems to be highly compartmentalized at the cellular and even subcellular level [6, 27]. Although H2O2 diffuses through lipid bilayers, cellular membranes, but also packed transmembrane proteins, appear to form a diffusion barrier for H2O2 that readily affects its effective concentrations [6]. In addition, water-traffic controlling aquaporins also influence H2O2 diffusion and may be essential in determining effective cellular levels of H2O2 [28]. The low concentration, the short half-live and the specific redox environment make it challenging to measure H2O2 within tissues and cells. An ideal assay should detect H2O2 in real time down to nanomolar concentrations within a specific cellular compartment of 10 cubic nanometer, and without any interferences by factors such as oxygen, pH etc. Unfortunately, we are still far from that, and must rely on a combination of approaches. While the luminol- hypochlorite assay achieves real-time qualities and ultrahigh sensitivity at the nanomolar scale, it does not allow for intracellular H2O2 detection. On the other hand, recent molecular approaches such as the H2O2-sensitive fusion protein OxyR-YFP [29] or the redoxsensitive fusion protein consisting of human glutaredoxin-1 and roGFP2 [30] allow for compartment-specific monitoring of redox changes within cells and tissues but are limited by sensitivity (micromolar range) and specificity.

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3. H2O2 Detection in Biological Systems: A Critical Review This section summarizes all currently available methods for H2O2 detection and highlights specific challenges and limitations in biological systems that still prevent thorough in vivo investigation of this important molecule. Figure 1 defines the five criteria that should be met by any ideal H2O2 assay with regard to specificity, sensitivity, real-time and compartment-specific properties and potential interferences. For over 100 years, various methods have been developed to detect H2O2; for a historic background the reader is referred to the extensive review by Schumb and colleagues [31]. First gravimetric and volumetric assays consisted of the direct chemical titration with permanganate, ceric or iodide ions of a weighed sample solution. More recently, a non-enzymatic assay was successfully applied to measure H2O2 removal by cultured cells, although the assay is not specific for H2O2 but also detects other hydroperoxides [32]. Conversion of H2O2 to oxygen and subsequent oxygen measurement using an oxygen electrode is another traditional approach. This gasometric method is still commonly employed and, under proper circumstances, is reliable; for everyday practice, however, it is less convenient than other procedures. The method involves catalytic decomposition of H2O2, e.g. by catalase, leading to a stoichiometric conversion of H2O2 to molecular oxygen that is detected by an oxygen electrode [33]. An interesting but rarely employed H2O2 detection method utilizes the H2O2 electrode, although it does not reach the sensitivity of fluorescence techniques [34]. The direct spectrophotometric determination of H2O2 in the UV range (specific absorption coefficient 230 = 74 M-1 cm-1) is needed to monitor highly concentrated stock solutions [35]. Due to the relatively low H2O2 sensitivity of the assay, high concentrations (ca. 100-1000 µM) must used that may rapidly inactivate enzymes [36-38]. The majority of assays, however, involve either enzymatic or non-enzymatic conversion of substrates by H2O2 (Figure 2). H2O2 assays that are most often employed to assess H2O2 metabolism of cells and enzymes are peroxidase-based methods. In these assays, indicator substances are oxidized by H2O2 in the presence of a peroxidase. Changes in fluorescence, absorption or luminescence intensity are quantified and directly related to the amount of H2O2. The high sensitivity and convenience of using fluorescent dyes have resulted in the wide application of peroxidase-based assays. A common assay employs the H2O2-dependent oxidation of scopoletin (7-hydroxy-6-methoxy-coumarin) to a non-fluorescent product by horseradish peroxidase. Loss of scopoletin fluorescence is measured at 460 nm [39]. Other peroxidase substrates as homovanillic acid [40] or Ampex Red [41] have also been used. An example of a peroxidase-based chemiluminescence assay is the HRPO-luminol assay [42] that allows the direct monitoring of H2O2 turnover. Because of their wide use, it seems worthy to discuss peroxidase-based H2O2 assays in more detail. There are several general caveats about peroxidase-based H2O2-assays: First, they constitute “trapping” methods requiring an incubation of several minutes to “trap” H2O2 , and they are not specifically designed to determine H2O2 concentrations but rather H2O2 turnover. This very basic principle has important consequences: H2O2 metabolism is significantly influenced by such assays leading to artifacts in evaluating cellular regulatory pathways. More simply, H2O2 might easily be underestimated, as only one part of the H2O2 pool is “trapped” in deference to other H2O2 metabolizing pathways. Another problem is that some compounds such as ascorbic acid or thiols are also substrates for the peroxidase and can

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compete with scopoletin to cause artifactual inhibition. Second, peroxidase-based assays are not specific for H2O2 for several reasons. One-electron transfers inherent in such assays may cause redox-cycling reactions or reduction of oxygen to superoxide. This is one of the reasons why peroxidase-based assays usually give lower signals in the presence of SOD. The radical generation by the analytical method itself is a severe artifact that might lead to misinterpretations [43-45]. Third, inhibitors that are often used in biological experiments may directly interfere with the peroxidase reaction as the basic principle of the assay. For instance, competitive inhibitors of heme enzymes such as sodium azide will also inhibit the peroxidase used in the assay, resulting in falsely low H2O2 concentrations. The non-fluorescent dye 2‟,7‟-dichlorodihydrofluorescin diacetate (H2DCF-DA) is widely used to estimate the intracellular production of oxidants during stress conditions. The oxidation of intracellularly trapped H2DCF-DA requires the removal of diacetate by esterases. Such activated H2DCF is intracellularly oxidized to the highly fluorescent derivative 2‟,7‟dichlorofluorescein (DCF). It is mistakenly assumed that DCF detects H2O2. H2DCF is oxidized to DCF by neither O2 - nor H2O2. Instead, only oxidants derived from H2O2 in the presence of catalytically active metal ions or peroxidases are able to oxidize H2DCF [46-48]. On the other hand, other oxidants such as cholesterol hydroperoxides, lipid hydroperoxides (both in the presence of hemin as iron source) as well as phenoxy radicals are also known to oxidize H2DCF [49, 50]. Thus, this assay detects non-specifically oxidant production in cells under stress situations. Although this approach is not specific for H2O2, its popularity is rationalized by to the lack of alternatives and may be justified with appropriate controls [5153]. Very recently, intracellular biological H2O2- or redox-sensors have been introduced. These approaches allow, for the first time, a space- and compartment-specific detection of H2O2 levels or redox changes. Belousov and coworkers fused the bacterial H2O2-sensor OxyR to circularly permutated yellow fluorescent proteins (YFP) in such an elegant way that conformational changes of OxyR in the presence of H2O2 lead to fluorescence changes of YFP [29]. The OxyR-YFP fusion protein is a ratiometric dye, meaning that at least two fluorescence values change in opposite directions in response to H2O2, and a mere ratio of both wavelengths is sufficient for H2O2 measurements. As a result, measurements are not dependent on the expression level of the dye which is usually hard to control in the laboratory. The novel dye has been successfully explored in other laboratories and has lead to exciting, novel insights regarding basic mechanisms of wound healing [54]. Figure 3 demonstrates the mitochondrial detection of H2O2 by mitochondrially targeted OxyR-YFP. The basic principle of fusing H2O2 or redox-responsive proteins to fluorescence proteins is now widely explored. Thus, a glutaredoxin 1-roGFP2 fusion protein allows dynamic live imaging of the glutathione redox potential in different cellular compartments with high sensitivity and temporal resolution. Altogether, these molecular fusion probes are very promising. However, there are still limitations to consider with regard to potential interferences of the dye with local conditions such as pH, sensitivity (e.g. of OxyR-YFP is limited to micromolar concentrations), delayed response (real-time) and reversibility. We will now discuss in more detail a previously introduced non-enzymatic H2O2 assay based on the non-enzymatic oxidation of luminol by the two-electron oxidant sodium hypochlorite. The luminol/hypochlorite assay allows for ultra-sensitive detection of H2O2 in biological systems down to nanomolar concentrations in real-time [55, 56].

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Figure 3. Example of the first compartment-specific H2O2 assay based on the OxyR-YFP fusion protein. Huh7 hepatoma cells were transfected with an OxyR-YFP expressing plasmid containing a mitochondrial targeting sequence. Changes in fluorescence allow for measurement of mitochondrial H2O2. Nuclei are counterstained (blue) with DAPI.

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4. Luminol Chemistry and Light Emission: General Mechanisms Luminol (5-amino-2,3-dihydro-1,4-phthalazinedione) and its chemiluminescence were first described by H. O. Albrecht in 1928 [57]. Albrecht‟s paper provided considerable information about blue luminol chemiluminescence. Several oxidants including hypochlorite and potassium ferricyanide were found to generate chemiluminescence in the presence of low H2O2 concentrations. It was further observed that H2O2 elicited a very weak generation of light, while the addition of catalysts such as peroxidases markedly enhanced the luminescence intensity. The importance of alkaline conditions was underscored and diazaquinone was postulated as a possible intermediate. Finally, an intermediate was regarded as the light emitter in lieu of luminol itself. Subsequently, important milestones in our perception of the inherent chemistry were achieved by the laboratories of White, Rauhut, Gundermann, and most recently, by Merenyi, Erikson and Lind [58-65]. For a comprehensive review, the article by Merenyi et al is recommended [63]. Meanwhile, luminol chemiluminescence has found many applications in different fields such as forensic medicine (imaging blood analysis) and molecular biology (e.g. antigen detection by western blotting) . The overall stoichiometry was shown to involve the consumption of 1 mole each of luminol and O2 to yield 1 mole of dicarboxylate and nitrogen. In each case, the corresponding dicarboxylates were proved to be the light-emitting species. Furthermore, the reaction of diazaquinone – the two-electron oxidized luminol – with H2O2 was shown to yield chemiluminescence as well [58, 66].

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5. Two-electron Oxidation of Luminol by Hypochlorite The analytical potential of the luminol/hypochlorite oxidation system became first evident during detailed luminescence studies designed to understand better the so-called luminol-dependent chemiluminescence of phagocytes [55, 67-70]. Figure 4A shows the typical blue chemiluminescence flash after the injection of hypochlorite into a luminol solution containing traces of H2O2. Figure 4B demonstrates the brevity of the light reaction: chemiluminescence is terminated within 2 seconds. Although oxidation of luminol by NaOCl was one of the first reactions discovered to yield luminescence [57] it did not attract further attention because of the advent of more efficient catalysts and technical limitations in measuring light [71]. With the exception of some historical studies [72, 73], diazaquinone as obligatory intermediate in the luminol oxidation by NaOCl was postulated [74] and later demonstrated [44, 59]. While HOCl reacts with luminol, OCl- does not [75, 76]. This fact, coupled with the pKa value of HOCL (7.3), explains the apparent pKa of luminol oxidation [44]. The two-electron generation of diazaquinone by hypochlorite is depicted in Figure 4. The initial reactants luminol and hypochlorite are shown in red, while the light reaction is shown in blue. Generally, the complete chemistry of luminol can be subdivided into i) pathways leading to the -hydroxy-hydroperoxide and ii) its decomposition pattern. The first part is heavily dependent on the exact composition of the reacting system (concentrations and nature of oxidants, additives, pH and buffer). In contrast, the decomposition pattern of the key intermediate depends only on the pH of the system. As mentioned earlier, luminol can be oxidized in one-and two electron oxidations, both leading to the diazaquinone. One-electron oxidation systems such as H2O2-peroxidase based assays directly generate luminol radical. Following redox cycling reactions, this luminol radical is able to reduce molecular oxygen (that is usually present in biological samples under aerobic conditions) to the O2 - that might further react with luminol or intermediates and complicate the reaction chemistry. It is clear from an analytical point of view that such a complex reaction mixture is unfavorable. Indeed, the generated O2 - can be detected by removal of oxygen or addition of SOD in luminolperoxidase based assays [65]. The major advantage of the luminol/hypochlorite oxidation system is that luminol is directly oxidized to diazaquinone while non-specific reactions are minimized. Once the diazaquinone is generated, the subsequent reaction cascade is very specific for H2O2. In the absence of H2O2, diazaquinone rapidly hydrolyses in a “dark” reaction. The decomposition of the hydroperoxide eventually yields aminophthalate, but only the mono-anion decomposition leads to chemiexcitation. The light yield increases with pH, having an apparent pKa at pH 8.2. The pH dependence is inherent to all luminol chemiluminescence and independent of the reaction mixtures

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Figure 4. Short chemiluminescence peak after injection of 1 µM NaOCl into a phosphate buffered aqueous solution at pH 7.4. Figure 4A demonstrates the blue chemiluminescence at 430 nm. Figure 4B shows the respective light intensities over time for two different H2O2 concentrations.

A direct proportional relation between H2O2 concentration and luminescence intensity was established for the NaOCl-luminol system [55, 68-70]. At very low H2O2 concentrations, the luminescence of this system apparently became independent of H2O2 [70]. Further careful experiments on the various inhibition mechanisms revealed two major groups of inhibitors of the light intensity in the luminol/NaOCl system [69]: Group A inhibits the chemiluminescence by directly reacting with hypochlorite and scavenging it. This group includes substances that react with hypochlorite, e.g. amino acids containing amino or sulfhydryl groups. Group B represents molecules that decompose H2O2, e.g. catalase, peroxidases or permanganate. These findings strongly supported the general scheme of luminol oxidation as demonstrated in Figure 5: Luminol is oxidized by NaOCl in a twoelectron-oxidation directly to the diazaquinone. Diazaquinone hydrolyses rapidly to dark products in aqueous solutions. In the presence of H2O2, however, hydroperoxide is formed. According to our studies, most of the chemiluminescence in a buffered aqueous luminol solution is due to small quantities of H2O2 that are present in these aerobic solutions and that are generated by photochemical reactions. Its contribution becomes apparent in the presence of catalase which efficiently removes even traces of H2O2. The detection of these low H2O2 levels establishes the luminol-hypochlorite assay as a sensitive non-enzymatic H2O2 assay for biological systems[ 55, 56, 70]. Figure 5 provides the rationale for the assay. Thus, all additional molecules that react with NaOCl will inhibit the luminescence. Optimal assay conditions include NaOCl concentrations of 1-10 µM and luminol concentrations of 50 µM (range 10-100 µM). At higher luminol concentrations, kinetics become faster and less efficient as the intramolecular amino group of luminol reacts competitively with the hydrazide group for NaOCl. The pH needs to be strictly maintained at 7.4 as it is crucial for luminol chemiluminescence. The major advantages of this luminescence reaction for biomedical sciences are the high sensitivity for H2O2 at physiological pH.

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Figure 5. Reaction scheme of the two-electron oxidation of luminol by hypochlorite. The importance of pH in luminol chemistry is taken into consideration by vertically aligning the different protonated species (acidic pH on top). The blue arrows and pathways mark the two-electron oxidation pathway of luminol by hypochlorite that specifically detects H2O2.

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6. Determination of H2O2 by Using Luminol and NaOCl Determination of H2O2 by the luminol/NaOCl system can be performed by two technical approaches either using a flow system or an injection device. Both of them require different preconditions and have different potentials and limitations. The flow system allows for continuous H2O2 determination in larger sample volumes and is most suitable for following rapid enzyme-dependent changes in H2O2 concentrations. The injection technique permits the end point determination of H2O2 in small sample volumes and allows a high sample throughput. Both techniques are now discussed in detail by demonstrating typical applications.

6.1. Determination of fast H2O2 Kinetics in Enzymatic Reactions with a Flow System In the flow system, the sample is continuously pumped out from a reaction reservoir and luminol and NaOCl are continuously added, allowing real-time registration of H2O2, e.g. during fast enzyme kinetics (Figure 6). An advantage is that the sample solution in the reservoir is not in contact with any of the detection reagents and thus, H2O2 can be measured independently of the detection system. Conditions can be controlled by temperature adjustment, magnetic stirring or even adding substrates or inhibitors during the experiment. Additional parameters can be determined in parallel (e.g. oxygen measurement). On the other hand, the flow technique requires a large sample volume (up to 100 ml) as the sample is

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continuously removed for H2O2 detection. This is usually not limiting since enzyme solutions can be highly diluted due to the sensitivity of the assay.

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Figure 6. Luminol/hypochlorite assay used in a flow system that allows for the continuous determination of H2O2 in real time. The sample solution can be independently controlled (injection of additional compounds, temperature control, magnetic stirring, measurement of additional parameters e.g. oxygen). The flow system has been successfully employed in studying H2O2 decomposition and generation by enzymes such as catalase, glutathione peroxidases and oxidases.

Figure 7. Determination of liver catalase activity using the luminol/hypochlorite assay in the flow mode. For details see text.

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Equipment (see Figure 4) luminometer (any luminometer can be used which allows the installation of a flow cell in front of the photomultiplier, e.g. the AutoLumat LB 953 from Berthold EG&G, Wildbad, Germany). The luminometer is controlled by a computer equipped with software for further processing of time/luminescence intensity data perfusion pump for NaOCl and luminol peristaltic pump for sample aspiration flow cell to allow separate and continuous addition of luminol and NaOCl in phosphate buffered saline (PBS) at pH 7.4 and the continuous addition of the sample. In the authors‟ laboratory, a peristaltic pump is used with a 3 mm polyethylene pipeline to aspirate continuously the sample solution (ca. 4 ml/min). Black 50 ml plastic syringes are loaded with luminol and NaOCl work solutions and both reactions are continuously pumped via the same perfusion pump into the polyethylene pipeline (ca. 12 ml/h). graduated 100 ml cylinder for sample solution magnetic stirrer to continuously mix sample solution temperature control unit (if necessary) Reagents

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50 ml 10-4 M luminol in 10 mM PBS at pH 7.4 (working solution) 50 ml 10-4 M NaOCl in tridistilled water (working solution) 100 ml 10-2 M H2O2 in tridistilled water for calibration Procedure The syringes are loaded with the working solutions of NaOCl and luminol and 50-100 ml PBS are added to the graduated cylinder. All pumps are switched on and the system is allowed to equilibrate for about 5 minutes. The optimal measuring range is found by adjusting the perfusion pump and calibrated by addition of 10-5 M H2O2 and catalase, respectively. Example 1: Determination of Catalase Activity at Physiological H2O2 Concentrations [77] (Figure 7) The luminol/hypochlorite assay was first successfully applied to ascertain catalase activity. An example is now described in more detail demonstrating calibration, data processing and data analysis. In Figure 7, a typical catalase experiment is shown. First, 10 µM final H2O2 is added to 100 ml 10 mM PBS pH 7.4. Liver homogenate is added at a later time point. A magnetic stirrer is used to ensure rapid mixing of the enzyme substrate solution. An exponential decay of the chemiluminescence can be observed that reaches background levels after several minutes. At this time, no changes in luminescence are detectable. The “background luminescence” has several causes and is H2O2-dependent (caused by small H2O2 impurities in the reagents) or -independent. Factors that contribute to H2O2-independent background chemiluminescence are the purity of the NaOCl solution and other side reactions of the luminol/hypochlorite system that also lead to the excited aminophthalate, e.g. in association with transition metals in the sample. The concentration of the NaOCl solution

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clearly defines the extent of the H2O2 independent background luminescence and should be as low as possible. Very often, the background luminescence is negligible. If not, it can be determined, e.g. in the presence of catalase, and subtracted from the data. As H2O2 concentration is directly proportional to the chemiluminescence intensity, one calibration concentration is usually sufficient. In contrast to most other enzymes, catalase does not follow Michaelis-Menten kinetics. It is not saturable by H2O2 and catalase activity is therefore described by the rate constant k = ln (S1/S2)/dt where dt is the measured time interval; S1 and S2 are H2O2 concentrations at time t1 and t2, respectively. In Figure 7, catalase activity is calculated from the exponential decay of H2O2 by linear regression analysis. In the above equation, the ratio (S1/S2) rather than absolute values of H2O2 concentrations is important so that k can be calculated directly from the luminescence intensities: k = ln (I1/I2)/dt where dt is the measured time interval; I1 and I2 are luminescence integrals at time t1 and t2. The constant k can be used as a direct measure of catalase concentration. The specific catalase activity k'1 is obtained by dividing k by the molar concentration of catalase (e): k'1 = k/e. k'1 is known for many catalases from different cell types. The value k'1 for purified catalase from human erythrocytes is 3.4 x 107 M-1s-1. This value is used to calculate the absolute concentration of enzyme in blood and tissues [36, 78]. The hypochlorite/luminol technique provides several advantages in comparison to conventional spectrophotometric and titrimetric catalase assays: i) due to the low H2O2 concentrations used, molecular oxygen is completely dissolved and not liberated in gaseous form that causes artifacts; ii) since maximal extracellular H2O2 concentrations are known to reach only micromolar levels, determinations of catalase activity at submicromolar concentrations reflect physiological conditions; iii) repetitive measurements are possible without loss of enzyme activity or cell viability. The assay has been successfully used to compare catalase activity of intact and homogenized cells/organelles [77].

Figure 8. Generation of H2O2 steady-state concentrations with catalase and glucose/glucose oxidase (GOX). The steady-state concentration of H2O2 is determined by the GOX/catalase ratio and can be maintained over hours. This tool appears to be very useful in studying signal functions of H 2O2 on a quantitative basis in various biological systems.

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Figure 9. H2O2 removal by cultured B6 fibroblasts (monolayer) in different volumes of culture medium. After bolus addition, H2O2 is decomposed by the cells within minutes. The short half-life of H2O2 should be considered when studying its effects on cellular functions.

Example 2: Steady-state Generation of H2O2 with a Glucose/glucose Oxidase/catalase System to Study H2O2–dependent Signaling Pathways [53, 79] The glucose/glucose oxidase system was recently developed in conjunction with the luminol/hypochlorite assay and appears to be a powerful tool for studying signal functions of H2O2 and hypoxia on a quantitative basis. More details are provided in a review article [79]. During the oxidation of glucose by glucose oxidase, H2O2 is generated following a zero-order kinetic with dH2O2/dt = kGOX (kGOX = rate constant) if O2 and glucose are maintained at constant concentrations. Accumulation of H2O2 can be controlled by adding appropriate amounts of catalase. H2O2 degradation rate by catalase is described by dH2O2/dt = kCAT x [H2O2]. Thus, steady-state levels of H2O2 are generated when kGOX = kCAT x [H2O2], and at a constant glucose and O2 concentration [H2O2] = kGOX / kCAT. By varying the enzyme activities, the H2O2 concentration can be adjusted and maintained over hours. The luminol/hypochlorite assay assists by measuring this steady-state as shown in Figure 8. Steady-state generation in turn allows exact time- and dose-dependent studies on redoxsensitive signaling pathways instead of simply adding H2O2 as bolus. The GOX/catalase system has been successfully employed to study the regulation of iron protein 1 (IRP1) by H2O2. It was shown that 10 µM H2O2 (steady state) suffice to activate IRP1 within 20 minutes by a still unknown signaling cascade [24, 53, 80]. It is also the first enzymatic system to generate well-defined hypoxia in cultured cells [79, 81].

6.2. End Point Determination of H2O2 Using an Injection System In this procedure, luminol is premixed with the sample (e.g. culture medium or tissue perfusate). At appropriate times, NaOCl is added and the luminescence intensity is measured immediately. The measurement is fast and only a small sample volume is required. Theoretically, luminol can be added just before the NaOCl injection to avoid any interferences of sample and detection system. Thus far, no such interferences have been observed by us, and we usually premix luminol with the samples for practical reasons. An

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injection device in measuring position is necessary because the luminescence reaction reaches completion within less than 2 seconds. As an advantage, less sample volume is needed and the procedure can be fully automated. Equipment luminometer with injection device in measuring position (e.g. AutoLumat LB 953 from Berthold EG&G, Wildbad, Germany). Other injection devices are helpful for complete automatization of the experiment, e.g. addition of cell stimulators. The luminometer should be controlled by a computer equipped with software allowing automated performance. polystyrene tubes for luminometer. Reagents

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stock solution of 10-3 M luminol in 10 mM PBS at pH 7. 4 (final concentration of luminol between 10-5 – 10-4 M) stock solution of 10-4 M NaOCl in tridistilled water (final concentration of NaOCl between 10-6 – 10-5 M) 10-3 M H2O2 in tridistilled water for calibration. Procedure The injector in measuring position is loaded with NaOCl solution and primed. For optimal measuring range, samples with PBS and luminol are loaded containing catalase (e.g. 10-7 M final concentration) and 10-5 or 10-6 M H2O2. If necessary, the NaOCl concentration may be adjusted. In a typical experiment, the injection device adds 50 µl of NaOCl (10-6 - 10-5 M final concentration) to 950 µl sample with luminol (5x10-5 M final concentration). Usually, samples are measured together with an H2O2 calibration solution at the beginning and end of each batch. Example 3: H2O2 Removal from Culture Medium by B6 Fibroblasts [53, 79] Figure 9 shows the removal of H2O2 from culture medium (three different volumes) by fibroblasts growing in monolayer culture. B6 fibroblasts were cultured in RPMI medium in 10 cm culture dishes at 37° C. 10-4 M H2O2 (final concentration) is added to the medium and 500 µl of the medium is transferred at different time points into polystyrene tubes. 10 µl luminol stock solution is added. After addition of 50 µl NaOCl (5x10-6 M final concentration), the luminescence intensity is recorded for 2 seconds. The culture medium should not contain serum. The determination of H2O2 removal in cell culture medium is important for all conditions where H2O2 is applied to cells in order to study its signaling functions.

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Conclusions This article describes a sensitive and non-enzymatic H2O2 assay which is based on the chemiluminescence reaction of luminol with hypochlorite. Actual H2O2 concentrations can be measured in monolayer cultures, perfusates, suspensions of intact cells and crude homogenates. One of the strengths of this assay is that it may be used to assess fast enzyme kinetics at very low H2O2 concentrations. These unique characteristics rendered the luminol/hypochlorite assay instrumental in developing novel enzymatic H2O2 and hypoxia models such as the GOX/CAT system as described in Chapter 11. Studies are under way that extend its application towards peroxidase-based H2O2 decomposition and peroxisomal H2O2 metabolism. In summary, the luminol/hypochlorite assay opens new niches in studying the functions and metabolism of H2O2 in biological systems.

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[32] Dringen R, Kussmaul L, Hamprecht B. Detoxification of exogenous hydrogen peroxide and organic hydroperoxides by cultured astroglial cells assessed by microtiter plate assay. Brain Res Protoc 1998 Mar;2(3):223-8. [33] del Rio LA, Ortega MG, Lopez AL, Gorge JL. A more sensitive modification of the catalase assay with the Clark oxygen electrode. Application to the kinetic study of the pea leaf enzyme. Anal Biochem 1977;80(2):409-15. [34] Test ST, Weiss SJ. Quantitative and temporal characterization of the extracellular H2O2 pool generated by human neutrophils. J Biol Chem 1984;259(1):399-405. [35] Beers RF, Sizer IW. A spectrometric method for measuring the breakdown of hydrogen peroxide by catalase. J Biol Chem 1952;195:133-40. [36] Aebi H. Catalase in vitro. Methods Enzymol 1984;105:121-6. [37] Bonnichsen R. Blood catalase. Methods Enzymol 1955;2:781-4. [38] Deisseroth A, Dounce AL. Catalase: physical and chemical properties, mechanism of catalysis, and physiological role. Physiol Rev 1970;50 (3):319-75. [39] Corbett JT. The scopoletin assay for hydrogen peroxide. A review and a better method. J Biochem Biophys Meth 1989;18:297-309. [40] Zhou M, Diwu Z, Panchuk-Voloshina N, Haugland RP. A stable nonfluorescent derivative of resorufin for the fluorometric determination of trace hydrogen peroxide: applications in detecting the activity of phagocyte NADPH oxidase and other oxidases. Anal Biochem 1997;253(2):162-8. [41] Baggiolini M, Ruch W, Cooper PH. Measurement of hydrogen peroxide production by phagocytes using homovanillic acid and horseradish peroxidase. Methods Enzymol 1986;132:395-400. [42] Wymann MP, Tscharner Vv, Deraulean DA, Baggiolini M. Chemiluminescence detection of H2O2 produced by human neutrophils during the respiratory burst. Anal Lett 1987;165:317-78. [43] Lock R, Johansson A, Orselius K, Dahlgren C. Analysis of horseradish peroxidaseamplified chemiluminescence produced by human neutrophils reveals a role for the superoxide anion in the light emitting reaction. Anal Biochem 1988;173(2):450-5. [44] Eriksen TE, Lind J, Merényi G. Oxidation of luminol by chlorine dioxide: formation of 5-aminophthalazine-1, 4-dione. J Chem Soc Faraday Trans 1981;77(9):2125-35. [45] Faulkner K, Fridovich I. Luminol and lucigenin as detectors for O2.-. Free Rad Biol Med 1993;15(4):447-51. [46] LeBel CP, Ischiropoulos H, Bondy SC. Evaluation of the probe 2',7'dichlorofluorescin as an indicator of reactive oxygen species formation and oxidative stress. Chem Res Toxicol 1992;5(2):227-31. [47] Rothe G, Valet G. Flow cytometric analysis of respiratory burst activity in phagocytes with hydroethidine and 2',7'-dichlorofluorescin. J Leukoc Biol 1990;47(5):440-8. [48] Zhu H, Bannenberg GL, Moldeus P, Shertzer HG. Oxidation pathways for the intracellular probe 2',7'- dichlorofluorescein. Arch Toxicol 1994;68(9):582-7. [49] Kalinich JF, Ramakrishnan N, McClain DE. The antioxidant Trolox enhances the oxidation of 2',7'- dichlorofluorescin to 2',7'-dichlorofluorescein. Free Radic Res 1997;26(1):37-47. [50] Cathcart R, Schwiers E, Ames BN. Detection of picomole levels of hydroperoxides using a fluorescent dichlorofluorescein assay. Anal Biochem 1983;134(1):111-6.

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[51] Jakubowski W, Bartosz G. 2,7-dichlorofluorescin oxidation and reactive oxygen species: what does it measure? Cell Biol Int 2000;24(10):757-60. [52] Ubezio P, Civoli F. Flow cytometric detection of hydrogen peroxide production induced by doxorubicin in cancer cells. Free Radic Biol Med 1994;16(4):509-16. [53] Mueller S, Pantopoulos K, Hentze MW, Stremmel W. A chemiluminescence approach to study the regulation of iron metabolism by oxidative stress. In: Stanley PE, editor. Bioluminescence and chemiluminescence: molecular reporting with photons. Baffins Lane, Chichester, Sussex: John Wiley & Sons Ltd; 1997. p. 338-41. [54] Niethammer P, Grabher C, Look AT, Mitchison TJ. A tissue-scale gradient of hydrogen peroxide mediates rapid wound detection in zebrafish. Nature 2009 Jun 18;459(7249):996-9. [55] Mueller S, Arnhold J. Fast and sensitive chemiluminescence determination of H2O2 concentration in stimulated human neutrophils. J Biolumin Chemilumin 1995;10(4):229-37. [56] Mueller S. Sensitive and nonenzymatic measurement of hydrogen peroxide in biological systems. Free Radic Biol Med 2000;29(5):410-5. [57] Albrecht HO. Über die Chemiluminescenz des Aminophthalsäurehydrazids. Z. Physikal. Chem 1928;136:321-30. [58] Gundermann KD. Chemiluminescence of luminol and related compounds. In: Cormier MJ, editor. Chemiluminescence and bioluminescence. New York: Plenum Press; 1973. [59] Gundermann KD. Grundlagen der Anwendungsmöglichkeiten von Chemilumineszenz, der Umwandlung von chemischer Energie in Licht. Vorträge der Rheinisch Westfälischen Akademie der Wissenschaften. Opladen: Westdeutscher Verlag; 1975. [60] Gundermann KD, McCapra F. Chemiluminescence in organic chemistry. Heidelberg, New York, Toronto: Springer Verlag; 1987. [61] White EH, Roswell DF. Luminol chemiluminescence. Chemiluminescence and bioluminescence. New York: Marcel Dekker, Inc.;1985. p. 215-44. [62] Roswell DF, White EH. The chemiluminescence of luminol and related hydrazides. Methods Enzymol 1978;57:409-23. [63] Merényi G, Lind J, Eriksen TE. Luminol chemiluminescence: Chemistry, Excitation, Emitter. J Biolumin Chemilumi 1990;5:53-6. [64] Lind J, Merényi G, Erikson TE. Chemiluminescence mechanism of cyclic hydracides such as luminol in aquous solutions. J Am Chem Soc 1983;105:7655-61. [65] Merényi G, Lind J, Eriksen TE. The reactivity of superoxid (O2-) and its ability to induce chemiluminescence and luminol. PhotochemPhotobiol. 1985 1985;41:203-8. [66] White EH, Nash EG, Roberts DR, Zafiriou OC. A chemiluminescent diazaquinone. J Am Chem Soc 1968;90(21):5932-3. [67] Brestel EP. Co-oxidation of luminol by hypochlorite and hydrogen peroxide: Implications for neutrophil chemiluminescence. Biochem Biophys Res Commun 1985;126(1):482-8. [68] Arnhold J, Mueller S, Arnold K, Grimm E. Chemiluminescence intensities and spectra of luminol oxidation by sodium hypochlorite in the presence of hydrogen peroxide. J Biolumin Chemilumin 1991;6(3):189-92.

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[69] Arnhold J, Mueller S, Arnold K, Sonntag K. Mechanisms of inhibition of chemiluminescence in the oxidation of luminol by sodium hypochlorite. J Biolumin Chemilumin 1993;8(6):307-13. [70] Mueller S. Untersuchungen zur Chemolumineszenz im System LuminolHypochlorsäure: Eine Möglichkeit zur Bestimmung von Wasserstoffperoxid an aktivierten neutrophien Granulozyten: Universität Leipzig; 1994. [71] Langenbeck W, Ruge U. Einige Versuche mit Luminol. Berichte der Deutschen Chemischen Gesellschaft 1937;2:367-9. [72] Bremer T. Le mécanisme de la chimiluminescence en solution - II Oxydation du 3aminophthalhydrazide. Bull Soc Chim Belges 1953;62:369-610. [73] Harris L, Parker AS. The chemiluminescence of the 3-aminophthalhydrazide. J Am Chem Soc 1935;57:1939-42. [74] Seitz WR. Chemiluminescence from the reaction between hypochlorite and luminol. J Phys Chem 1975;79(2):101-6. [75] Vorobeva TP, Kozlov YN, Koltypin YV, Purmal AP, Rusin BA, Talroze VL, et al. Oxidation of luminol associated with chemiluminescence. Communication 1. Mechanism of oxidation by hypochlorite. Izv Akad Nauk SSSR 1976;10:2187-93. [76] Isacsson K, Wettermark G. The determination of inorganic chlorine compounds by chemiluminescence reactions. Anal Chim Acta 1976;83:227-39. [77] Mueller S, Riedel HD, Stremmel W. Direct evidence for catalase as the predominant H2O2 -removing enzyme in human erythrocytes. Blood 1997;90(12):4973-8. [78] Aebi H. Catalase. In: Bergmeyer HU, Bergmeyer J, Graál M, editors. Methods of enzymatic analysis. 3rd ed. Weinheim, Deerfield Beach, Florida, Basel: Verlag Chemie; 1983. p. 273-86. [79] Mueller S, Millonig G, Waite GN. The GOX/CAT system: A novel enzymatic method to independently control hydrogen peroxide and hypoxia in cell culture. Adv Med Sci. 2009 Nov 27:1-15. [80] Pantopoulos K, Mueller S, Atzberger A, Ansorge W, Stremmel W, Hentze MW. Differences in the regulation of IRP-1 (iron regulatory protein-1) by extra- and intracelullar oxidative stress. J Biol Chem. 1997;272:9802-8. [81] Millonig G, Hegedusch S, Becker L, Seitz HK, Schuppan D, Mueller S. Hypoxiainducible factor 1 alpha under rapid enzymatic hypoxia: cells sense decrements of oxygen but not hypoxia per se. Free Radic Biol Med. 2009 Jan 15;46(2):182-91.

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Index

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A absorption spectra, 85, 202, 263 absorption spectroscopy, 213, 215, 216 abstraction, 32, 50, 63, 80, 83, 89, 93, 114, 116, 139, 140, 141, 168, 171, 172, 187, 191, 246, 249 access, 79, 83, 222, 269 accessibility, 83 accounting, 47, 55 acetylation, 258 acidic, 5, 29, 31, 33, 34, 49, 51, 53, 61, 66, 99, 100, 101, 127, 128, 162, 171, 180, 194, 227, 229, 230, 232, 238, 241, 271, 292 acidity, 173 activation energy, 26, 33 active compound, 29, 43 active site, 21, 63, 96, 123, 125, 126, 132, 222 adaptability, 14 adaptation, 16, 152, 267, 276 additives, 290 adduction, 158, 183 adenine, 150, 189, 192, 206, 245 adhesion, 176, 177, 178 adipocyte, 178 adipose, 158, 164 adipose tissue, 158, 164 adjustment, 98, 292 ADP, 4, 5, 7 aerobic bacteria, 8 aerobic capacity, 13, 14 age, 2, 7, 14, 93, 124, 273 age-related diseases, 14, 93 aggregation, 60, 122, 126, 177 aging studies, 15 alanine, 89, 90, 116 albumin, 92, 123

alcohols, 36, 63, 80, 89, 116, 138, 148 aldehydes, 36, 41, 118, 124, 131, 138, 142, 143, 145, 149, 150, 151, 153, 207, 250 algae, 6, 8, 9 aliphatic compounds, 161 alkane, 171, 172 alkenes, 35, 173, 180 alkylation, 178, 183 alpha-tocopherol, 73, 153, 182 alters, 142 alveoli, 266 ambient air, 266, 267, 269, 270, 271 amine, 156, 253, 262 amines, 46, 65, 73, 149 amino, 29, 31, 32, 34, 36, 40, 55, 59, 60, 61, 63, 66, 71, 75, 80, 81, 84, 85, 88, 89, 90, 95, 114, 117, 121, 122, 123, 124, 127, 128, 129, 130, 131, 132, 133, 138, 148, 149, 151, 165, 169, 187, 188, 189, 190, 191, 192, 197, 199, 289, 291 amino acid, 29, 31, 32, 34, 36, 40, 55, 59, 60, 61, 63, 66, 71, 75, 80, 81, 88, 89, 90, 114, 117, 121, 122, 123, 124, 127, 128, 129, 130, 131, 132, 133, 169, 190, 192, 199, 291 amino groups, 32, 82, 149, 165 ammonium, 46, 243 amyloid beta, 90 ancestors, 8 anhydrase, 70 antibody, 36, 41, 113, 130 antigen, 71, 126, 128, 176, 289 antigen-presenting cell, 71 antioxidant, 1, 2, 23, 24, 48, 60, 66, 73, 101, 130, 139, 153, 156, 166, 170, 175, 179, 273, 300 aorta, 69 apoptosis, 2, 8, 12, 13, 14, 16, 99, 126, 128, 133, 134, 150, 151, 153, 158, 159, 257, 283, 298

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Index

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aqueous solutions, 31, 37, 51, 67, 68, 104, 106, 186, 220, 238, 261, 263, 291 Argentina, 137 arginine, 24, 46, 67, 116, 117, 191, 231 argon, 225, 227 aromatic compounds, 30 aromatic rings, 32, 34, 240, 246 aromatics, 121 arrest, 124 arteries, 41 arthritis, 43 ascorbic acid, 236, 238, 251, 287 aspartic acid, 116 aspirate, 294 aspiration, 294 assessment, 141, 154, 182, 185 assimilation, 3, 6, 8 atherosclerosis, 43, 70, 93, 147, 164, 176, 273 atherosclerotic vascular disease, 177 atmosphere, 7, 8, 19, 93 atmospheric pressure, 270 atomic orbitals, 20 atoms, 20, 21, 37, 49, 58, 163, 165, 171 ATP, 3, 4, 5, 6, 7, 12, 13, 109, 127 attachment, 127, 208 attribution, 244 autoimmune diseases, 99 autoimmunity, 113 automatization, 297 autooxidation, 165, 171, 219, 233

B bacteria, 2, 3, 4, 6, 7, 8, 9, 10, 11, 16, 30, 53, 97, 270, 273, 280, 284 bacterium, 16 banks, 4 base, 4, 34, 65, 67, 121, 186, 187, 188, 189, 191, 192, 193, 198, 199, 200, 202, 210, 218, 220, 236, 237, 238, 245 base pair, 65 basement membrane, 63 basic research, 131 benefits, 2, 14, 273 bicarbonate, 27, 31, 39, 68, 236, 237, 262 biliverdin, 179 bioavailability, 24, 57, 169 biochemistry, 15, 38, 51, 75, 96, 105, 131, 152, 153, 164, 175, 182 bioenergy, 46 biological activities, 37, 150 biological activity, 48, 55, 68, 105 biological consequences, 60, 131, 198

biological fluids, 199 biological media, 25 biological processes, 23, 31, 43, 128, 207 biological responses, 197 biological roles, 164 biological samples, 179, 180, 207, 223, 290 biological stability, 173 biological systems, 29, 30, 35, 36, 41, 43, 49, 50, 51, 52, 53, 54, 57, 60, 65, 66, 71, 75, 113, 114, 121, 154, 156, 169, 171, 179, 201, 204, 206, 207, 208, 209, 230, 231, 232, 235, 236, 237, 238, 239, 240, 243, 250, 251, 252, 254, 256, 258, 259, 262, 281, 283, 286, 287, 288, 291, 295, 298, 299, 301 biologically active compounds, 70 bioluminescence, 301 biomarkers, 67, 131, 145, 193, 198 biomass, 7 biomaterials, 104 biomolecules, 20, 21, 22, 23, 24, 26, 30, 31, 32, 34, 36, 43, 44, 48, 49, 50, 52, 64, 65, 66, 95, 99, 113, 138, 139, 145, 151, 162, 169, 175, 207, 284 biosphere, 7 biosynthesis, 67, 138, 139 birds, 13, 14 blood, 43, 52, 54, 57, 69, 95, 101, 107, 176, 179, 181, 183, 184, 266, 267, 277, 289, 295 blood plasma, 179, 181, 184 blood pressure, 43 blood vessels, 54, 267 bloodstream, 99, 177 body size, 14 bonding, 20, 47, 55, 90 bonds, 20, 21, 35, 47, 79, 80, 81, 83, 89, 90, 123, 124, 131, 132, 140, 146, 165, 172, 238, 245 bone, 266 bone marrow, 266 brain, 41, 93, 100, 146, 152, 155, 156, 157, 266, 267 Brazil, 19, 38 breakdown, 13, 17, 145, 271, 300 by-products, 113

C calcium, 4, 177, 181, 226, 273, 283 calcium carbonate, 4 calibration, 195, 226, 294, 297 calorie, 2 cancer, 14, 154, 176, 213, 265, 267, 280, 301 cancer cells, 267, 301 candidates, 97 cannabinoids, 165

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Index capillary, 199 carbohydrate, 80 carbohydrates, 30, 80, 89, 114, 139, 284 carbon, 3, 6, 8, 19, 30, 32, 35, 45, 51, 67, 69, 75, 76, 77, 79, 80, 82, 89, 94, 104, 105, 114, 116, 117, 124, 139, 140, 142, 149, 150, 158, 162, 164, 165, 171, 172, 173, 174, 179, 211, 220, 245, 246, 247, 248, 249, 250, 262 carbon atoms, 142, 164 carbon dioxide, 3, 19, 51, 68, 69, 94, 104, 105, 211 carbon monoxide, 179 carbon-centered radicals, 32, 76, 77, 80, 82, 116, 117, 150, 158, 220, 247, 248, 249, 250 carbonyl groups, 117, 118, 130, 131 carboxyl, 82, 84 carboxylic acid, 164, 205 carboxylic acids, 164, 205 carcinogenesis, 200 cardiovascular disease, 14, 60, 267 carotenoids, 36 case study, 16 caspases, 8 casting, 36 catabolism, 66 catabolized, 114 catalysis, 34, 88, 96, 240, 300 catalyst, 99, 125 catalytic activity, 56, 128 catalytic properties, 111 cation, 21, 45, 48, 61, 84, 86, 90, 122, 162, 187, 188, 189, 190, 191, 210, 223, 225, 230, 234, 239, 243, 245, 246, 263 CBS, 79 cell culture, 209, 224, 238, 260, 265, 267, 270, 271, 274, 280, 281, 297, 302 cell culture method, 267 cell cycle, 2, 124, 126, 158, 159, 283 cell death, 8, 35, 109, 110, 208, 257, 283, 285 cell division, 11 cell membranes, 165, 263 cell metabolism, 162 cell signaling, 38, 138, 162, 164, 165, 174, 273 cell size, 11 cell surface, 154 cellular homeostasis, 1 cellular regulation, 103 cellular signaling pathway, 60 central nervous system, 143, 149, 263 cerebrospinal fluid, 100, 111 ceruloplasmin, 92, 101 CH3COOH, 230 chain propagation, 170 challenges, 184, 287

305

changing environment, 14 chelates, 98, 108, 239 chemical, 3, 6, 8, 15, 20, 21, 22, 25, 33, 35, 36, 37, 44, 45, 46, 47, 49, 50, 51, 66, 80, 89, 105, 114, 122, 132, 144, 162, 173, 177, 182, 193, 207, 208, 209, 211, 212, 213, 216, 239, 259, 261, 262, 263, 277, 287, 300 chemical bonds, 122 chemical properties, 20, 37, 45, 261, 300 chemical reactions, 8, 22, 162 chemical reactivity, 47, 207, 208, 209, 239 chemical structures, 50 chemicals, 7 chemiluminescence, 152, 156, 227, 231, 287, 289, 290, 291, 294, 298, 300, 301, 302 chemokines, 178 chemotherapy, 267 chlorination, 53, 61, 70 chlorine, 31, 34, 40, 300, 302 chlorophyll, 6, 8 chloroplast, 16 cholesterol, 35, 36, 41, 71, 93, 103, 142, 288 choline, 144 chromatographic technique, 196, 202, 213 chromatography, 141, 194, 199, 218 chronic granulomatous disease, 299 chymotrypsin, 127 cigarette smoke, 161, 273 circulation, 70 cirrhosis, 284 citrulline, 46, 169 City, 5 classes, 25, 34, 36, 144, 204, 212 classification, 114 cleavage, 51, 77, 79, 118, 122, 127, 128, 131, 132, 140, 165 clinical application, 131 cloning, 134 clusters, 5, 6, 7, 29, 30, 55, 57, 58, 114, 123, 240 CO2, 3, 4, 5, 6, 7, 19, 27, 28, 31, 51, 69, 86, 94, 95, 170, 172, 211, 220, 232, 237, 238, 240 cobalt, 262, 267 coding, 125 coenzyme, 231 collateral, 176 colon, 151, 159 colon cancer, 151, 159 combustion, 21 commercial, 200 communication, 93 community, 96, 97 compatibility, 12 compensation, 7

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competition, 11, 83, 171, 251, 263 compilation, 39 complexity, 4, 7, 11, 16, 30, 149, 180, 273 complications, 60, 156 composition, 53, 65, 92, 99, 218, 290 compounds, 25, 30, 32, 37, 39, 46, 50, 56, 63, 66, 84, 88, 116, 130, 135, 143, 145, 149, 154, 155, 176, 185, 186, 190, 201, 202, 208, 209, 212, 218, 219, 222, 224, 225, 226, 228, 229, 231, 232, 240, 246, 247, 248, 287, 293, 301, 302 computer, 294, 297 conception, 1, 4, 161, 177 concordance, 12 conductivity, 77, 85, 86, 202 configuration, 5, 172, 211 conflict, 2 conjugated dienes, 207 consensus, 60, 120, 129 conservation, 21, 22 constituents, 164, 165, 173, 182, 189, 207, 243 Constitution, 106 consumption, 7, 39, 53, 54, 205, 218, 219, 249, 269, 270, 271, 272, 289 containers, 267 controversial, 3, 46, 195 COOH, 205 coordination, 55, 57, 95, 99, 100 copper, 33, 55, 58, 59, 72, 73, 90, 91, 92, 93, 94, 96, 97, 98, 100, 101, 102, 103, 107, 108, 110, 111, 141, 240, 261, 284 coronary artery disease, 70 cost, 11, 100, 283 covalent bond, 5, 20, 21, 30 crown, 219 crust, 5 crystals, 88 CSF, 100 culture, 251, 268, 269, 270, 271, 272, 276, 277, 280, 285, 296, 297 culture conditions, 268, 271, 277 culture media, 272 culture medium, 280, 296, 297 cyanide, 267 cyanide poisoning, 267 cycles, 15, 19, 270, 271 cycling, 5, 92, 94, 98, 100, 142, 209, 221, 288, 290 cyclooxygenase, 36, 138, 143, 165, 176 cystathionine, 79 cysteine, 26, 59, 61, 70, 73, 75, 88, 89, 90, 99, 116, 119, 120, 125, 130, 132, 133, 134, 149, 174, 175, 178, 183 cystine, 76, 77, 89, 119, 120

cytochrome, 2, 8, 12, 13, 15, 33, 36, 41, 55, 57, 58, 66, 92, 100, 102, 122, 141, 150, 154, 158, 163, 222, 243, 257, 258, 272 cytochromes, 92 cytokines, 46, 176, 177, 178, 226 cytoplasm, 125 cytoplasmic inheritance, 11 cytosine, 65, 69, 187, 191, 192, 198 cytoskeleton, 9 cytotoxicity, 41, 57, 71, 177

D D-amino acids, 82 danger, 8 data analysis, 294 data processing, 294 decay, 53, 95, 96, 97, 173, 204, 212, 216, 217, 219, 222, 230, 251, 256, 294 decomposition, 30, 31, 51, 54, 61, 86, 96, 97, 105, 106, 115, 117, 138, 145, 149, 151, 165, 167, 170, 171, 173, 185, 186, 187, 188, 189, 190, 191, 197, 219, 225, 232, 233, 236, 237, 240, 248, 287, 290, 293, 298 decomposition reactions, 185, 187, 188, 189, 191, 197 defence, 94, 101, 273 deficiency, 7, 8 degradation, 60, 108, 124, 126, 127, 128, 129, 130, 132, 133, 134, 148, 149, 162, 179, 185, 186, 190, 207, 241, 267, 272, 279, 296 degradation rate, 296 dehydration, 188 deoxyribonucleic acid, 110 deoxyribose, 80, 187, 191, 192, 207, 241 depolarization, 7 depression, 88 depth, 276 deregulation, 103 derivatives, 63, 72, 118, 141, 161, 169, 171, 172, 174, 177, 178, 179, 181, 183, 194, 205, 231 desorption, 144, 175 destruction, 101, 114, 273, 278 detectable, 27, 201, 202, 213, 237, 249, 251, 294 detection, 36, 60, 77, 104, 107, 129, 130, 144, 158, 174, 179, 193, 194, 195, 199, 200, 201, 202, 203, 204, 207, 209, 213, 216, 218, 221, 222, 223, 224, 227, 229, 230, 231, 233, 235, 237, 238, 240, 241, 242, 243, 249, 250, 252, 253, 255, 258, 259, 260, 262, 277, 283, 286, 287, 288, 289, 291, 292, 296, 300, 301 detection system, 292, 296 detoxification, 60, 75, 272

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Index diabetes, 14, 93, 147, 152, 164, 176, 178, 265 dienes, 35 diet, 24, 53, 138 diffusion, 26, 28, 29, 30, 35, 47, 50, 53, 54, 55, 65, 69, 93, 95, 96, 99, 105, 162, 166, 172, 181, 263, 268, 269, 270, 271, 272, 273, 280, 284, 286, 299 diffusion rates, 47 diffusivity, 170 digestion, 129, 193, 194, 198, 235 dimerization, 30, 53, 61, 116, 119, 120, 235, 253 dimethylsulfoxide, 242 dipeptides, 71 direct measure, 295 discrimination, 25 disease progression, 60 diseases, 10, 14, 16, 23, 33, 50, 92, 101, 130, 138, 157, 175, 176 disorder, 157 displacement, 273, 284 dissociation, 51, 55, 56, 76, 83, 84, 211, 252 dissolved oxygen, 270 distribution, 20, 47, 107, 133, 134, 154, 169 diversity, 151, 154 DNA, 2, 11, 23, 25, 30, 36, 37, 40, 44, 53, 65, 66, 67, 73, 74, 95, 105, 106, 110, 111, 113, 114, 138, 143, 151, 162, 178, 185, 186, 188, 189, 190, 191, 192, 193, 194, 195, 196, 197, 198, 199, 200, 207, 267 DNA damage, 2, 30, 67, 110, 113, 114, 192, 195, 197, 199 DNA lesions, 185, 190, 191, 192, 193, 194, 195, 196, 197, 198 DNA repair, 113, 195 DNA strand breaks, 65, 195 docosahexaenoic acid, 137, 139, 140, 143, 144, 145, 155, 156 donors, 3, 6, 7, 19, 60, 72, 77, 86, 123, 161, 163, 179, 219, 225, 226, 262 double bonds, 35, 36, 50, 63, 79, 89, 138, 139, 141, 149, 164, 165 down-regulation, 177, 178

E editors, 302 eicosapentaenoic acid, 143, 154 electricity, 3 electrochemistry, 110 electrodes, 93, 226 electromagnetic, 211, 214, 215 electron pairs, 5

307

electron paramagnetic resonance, 56, 107, 173, 202, 260, 263 Electron Paramagnetic Resonance, 213 electrons, 3, 4, 5, 6, 7, 8, 19, 20, 21, 22, 26, 46, 47, 49, 92, 93, 99, 104, 213, 261 electrophoresis, 129, 195 ELISA, 130, 135, 200 ELISA method, 135 emission, 40, 203, 209, 216, 217, 227, 234, 250, 252 employment, 23, 149, 150, 196 encoding, 10, 11, 12, 13 endogenous synthesis, 161 endothelial cells, 144, 146, 151, 176, 177, 178 endothelium, 38, 66, 176, 177, 178 endothermic, 85 energy, 2, 3, 4, 5, 6, 7, 11, 15, 16, 19, 20, 21, 22, 30, 51, 138, 157, 164, 178, 203, 210, 213, 214, 227, 267, 277 England, 16, 155 environment, 5, 49, 55, 83, 92, 99, 116, 128, 131, 138, 145, 165, 172, 174, 197, 209, 236, 240, 249, 270, 286 enzyme, 23, 46, 54, 55, 58, 60, 63, 66, 88, 89, 97, 122, 123, 125, 130, 134, 135, 152, 156, 165, 169, 196, 208, 213, 223, 224, 226, 256, 258, 270, 272, 273, 277, 292, 294, 295, 296, 298, 300, 302 enzyme immunoassay, 135 enzyme inhibitors, 213 enzyme-linked immunosorbent assay, 130 enzymes, 8, 21, 24, 27, 33, 34, 36, 100, 113, 120, 126, 128, 129, 133, 138, 141, 143, 148, 152, 157, 176, 178, 195, 222, 223, 231, 268, 271, 272, 276, 280, 287, 288, 293, 295 epithelial cells, 67 epithelial lining fluid, 71 EPR, 38, 56, 57, 58, 104, 109, 152, 173, 202, 204, 213, 214, 215, 221, 222, 224, 229, 230, 231, 238, 241, 242, 243, 250 equilibrium, 29, 31, 34, 49, 50, 51, 53, 76, 79, 80, 81, 83, 93, 172, 174, 210, 218, 231, 238, 248, 256, 266, 267, 276 equipment, 213 erythrocyte membranes, 69 erythrocytes, 54, 69, 143, 153, 154, 267, 295, 302 ESR, 70, 99, 108 ESR spectra, 70 essential fatty acids, 164 ester, 142, 144, 153, 173, 205, 231 ethanol, 23, 242 ethers, 80, 89 ethylene, 62, 107 eukaryote, 8, 9, 16

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eukaryotic, 8, 9, 10, 11, 16, 127, 144, 284 eukaryotic cell, 9, 10, 11, 127, 284 evidence, 2, 9, 10, 12, 16, 23, 24, 25, 37, 46, 72, 80, 81, 85, 92, 95, 109, 120, 141, 149, 150, 151, 157, 162, 173, 175, 179, 191, 279, 302 evolution, 2, 4, 6, 7, 8, 9, 11, 12, 13, 14, 15, 16, 20, 23, 25, 37, 38, 44, 49, 130, 132, 273 excitation, 209, 217, 226, 250, 258 excretion, 100 exons, 9 experimental condition, 204, 225, 271 experimental design, 180, 239 exposure, 40, 49, 53, 62, 77, 79, 127, 135, 185, 210, 211, 213, 225, 258, 267, 272, 274, 280 extinction, 9, 203, 216, 218, 221, 222, 249 extracellular matrix, 63, 72 extraction, 193, 194, 195, 196, 198

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F fabrication, 15 FAD, 46 families, 145, 284 family members, 273 fatty acids, 32, 35, 36, 61, 63, 66, 74, 75, 79, 80, 88, 138, 141, 144, 145, 149, 152, 161, 162, 164, 165, 169, 172, 173, 174, 175, 176, 177, 178, 179, 180, 182, 183 fermentation, 2, 7, 12 ferric ion, 106, 107, 108 ferritin, 92, 99, 101, 110, 285 ferrous ion, 106, 109, 157, 239 fertility, 12, 13, 14 fertilization, 13 fibrin, 134 fibroblasts, 133, 296, 297, 298, 299 fibrosis, 284 fidelity, 113 films, 5 fish, 153 fission, 30 fitness, 12, 13, 14, 20, 102 flexibility, 139, 272 flight, 175 fluctuations, 279 fluid, 240, 263, 270, 283 fluorescence, 69, 101, 181, 202, 203, 207, 212, 213, 217, 218, 223, 229, 235, 241, 244, 250, 257, 258, 260, 261, 286, 287, 288, 289 fluorophores, 262 force, 3, 17, 85 Ford, 70, 108 formamide, 186, 187

formula, 21, 165 fragments, 145 France, 185 free radicals, 2, 20, 21, 22, 23, 24, 41, 43, 47, 68, 75, 82, 104, 106, 108, 131, 138, 143, 153, 154, 201, 204, 207, 249 fructose, 63 fungi, 9, 10, 12, 125, 270 fusion, 286, 288, 289

G gamma radiation, 199 gamma-tocopherol, 73 gel, 129, 195, 196 gene expression, 1, 16, 138, 143, 151, 156, 159, 164, 277 gene regulation, 179 genes, 8, 9, 10, 11, 12, 13, 30, 125, 176, 267, 279, 285, 299 genetic code, 4 genome, 10, 11, 12, 16, 17 geometry, 55, 215 Germany, 113, 265, 283, 294, 297 gland, 270 global warming, 5 glucose, 3, 63, 178, 268, 270, 271, 276, 280, 283, 295, 296 glucose oxidase, 283, 295, 296 glutamate, 117 glutathione, 23, 26, 33, 36, 39, 41, 44, 59, 67, 70, 77, 81, 99, 101, 106, 109, 126, 130, 131, 138, 148, 155, 156, 158, 163, 174, 175, 183, 206, 207, 222, 236, 238, 244, 251, 274, 281, 288, 293, 299 glycerol, 63, 144, 164 glycine, 89, 90, 116 glycol, 208, 224 glycolysis, 2 glycosaminoglycans, 63, 72 grants, 37, 66 grouping, 8, 9 growth, 3, 4, 7, 25, 139, 151, 159, 165, 178, 272, 273, 285 growth factor, 25, 178, 273, 285 guanine, 30, 65, 69, 73, 106, 188, 189, 190, 191, 192, 193, 197, 198, 199 guidance, 96

H habitats, 9

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Index hair, 272 half-life, 54, 66, 122, 192, 296 halogen, 246 halogenation, 34 harmful effects, 29 healing, 176 health, 20, 39, 70, 103, 134, 154, 164, 267 heart disease, 265 heart failure, 176 height, 127, 269, 270 heme, 27, 33, 36, 39, 46, 49, 55, 56, 57, 58, 59, 66, 95, 162, 177, 179, 181, 183, 184, 253, 255, 257, 258, 288 heme oxygenase, 177, 183, 184 hemochromatosis, 92, 103, 284 hemoglobin, 33, 52, 54, 56, 57, 70, 95, 102, 133, 163, 230, 262 hemostasis, 177 hepatocytes, 151 hepatoma, 279, 289 heterogeneity, 28, 144 histidine, 56, 59, 61, 63, 116, 120, 123, 132, 149, 158, 174, 238 historical overview, 38 history, 97, 283 homeostasis, 1, 4, 39, 157, 281, 298 homocysteine, 79 homolytic, 30, 51, 76, 77, 162, 169, 172 homovanillic acid, 287, 300 hormone, 2, 151, 159, 272, 273 hormone levels, 2 hormones, 25, 164, 284 host, 9, 11, 33, 94, 143, 176 human, 23, 36, 40, 41, 62, 63, 67, 70, 71, 72, 74, 88, 103, 105, 110, 111, 114, 132, 134, 143, 153, 154, 156, 159, 161, 175, 177, 179, 181, 183, 184, 199, 200, 266, 273, 281, 286, 295, 298, 300, 301, 302 human body, 266 human health, 103 human neutrophils, 161, 181, 298, 300, 301 hybrid, 13, 17, 127 hydrazine, 129 hydrocarbons, 39 hydrogen abstraction, 30, 31, 32, 77, 80, 89, 115, 141, 171, 172, 180, 191, 192, 220, 248 hydrogen atoms, 30, 50, 89, 104, 116, 140, 171, 215, 261 hydrogen gas, 4 hydrogen peroxide, 15, 38, 39, 40, 41, 61, 62, 74, 90, 91, 102, 106, 107, 108, 109, 110, 111, 132, 133, 210, 221, 260, 262, 263, 281, 298, 299, 300, 301, 302

hydrolysis, 5, 86, 87, 115, 120, 165, 172, 188, 191, 194, 205, 226, 230 hydroperoxides, 30, 32, 35, 36, 40, 41, 116, 138, 142, 145, 148, 149, 151, 152, 154, 155, 156, 157, 170, 186, 207, 248, 250, 287, 288, 300 hydrophobicity, 44, 164, 218 hydrothermal process, 7 hydroxide, 4, 97, 101, 174 hydroxyl, 22, 38, 51, 65, 68, 72, 77, 82, 84, 88, 90, 91, 93, 104, 105, 107, 108, 109, 114, 122, 123, 133, 139, 141, 153, 172, 185, 186, 191, 193, 208, 210, 211, 220, 222, 232, 236, 238, 239, 240, 241, 244, 261, 263, 272, 274, 284 hyperfine interaction, 215 hypothesis, 10, 12, 13, 15, 16, 99, 161, 175 hypoxia, 265, 266, 267, 268, 269, 270, 271, 272, 274, 276, 277, 278, 279, 280, 281, 282, 283, 296, 298, 302 hypoxia-inducible factor, 267, 280 hypoxic cells, 277

I ideal, 271, 283, 285, 286, 287 identification, 25, 134, 144, 161, 177, 179, 184, 186, 199, 214, 215, 255, 282 identity, 207, 209, 235, 238, 239, 240, 241, 242, 255 idiopathic, 103 illumination, 258 immune regulation, 24 immune response, 43, 46, 93, 128, 134 immune system, 113 immunohistochemistry, 70, 130 impurities, 294 in transition, 61 in vitro, 60, 73, 95, 100, 101, 102, 128, 139, 141, 143, 155, 156, 157, 161, 169, 177, 178, 197, 260, 261, 265, 267, 272, 273, 281, 285, 299, 300 in vivo, 2, 24, 25, 30, 33, 37, 50, 57, 59, 60, 62, 69, 72, 92, 93, 95, 97, 99, 100, 101, 102, 113, 120, 122, 123, 128, 143, 145, 154, 155, 156, 162, 163, 165, 166, 169, 172, 174, 175, 176, 177, 179, 180, 183, 185, 193, 197, 198, 199, 209, 230, 231, 251, 260, 261, 287 incidence, 14 incubation time, 227 indirect effect, 197 individuals, 161, 176 inducer, 153, 155 inducible enzyme, 177 induction, 177, 178, 184, 198, 226, 232, 258 infancy, 25

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infection, 176, 277 infertility, 13, 14 inflammation, 36, 41, 43, 46, 60, 72, 99, 161, 169, 175, 176, 177, 178, 180, 183, 198, 273 inflammatory cells, 178, 273, 285 inflammatory disease, 63, 177, 180, 265 inflammatory mediators, 176 inflammatory responses, 179, 283 inheritance, 14 inherited disorder, 284 inhibition, 57, 67, 95, 133, 134, 140, 145, 153, 167, 171, 176, 177, 180, 182, 208, 236, 238, 244, 250, 263, 267, 273, 288, 291, 302 inhibitor, 62, 72, 145, 170, 183, 184 initiation, 72, 96, 139, 147, 150, 154, 169, 170, 176, 255 insertion, 145 insulin, 82, 90 integration, 226 integrin, 177, 181 integrity, 126 interface, 133, 267, 270 interference, 129, 205, 207, 208, 209, 239 interphase, 268 intervention, 23, 24 intima, 40, 178 introns, 9 inventions, 15 inversion, 21, 22 investment, 20 ion transport, 140, 157 ionization, 76, 144, 175, 189, 195, 210 ionizing radiation, 79, 191, 193, 195, 198, 210 ions, 4, 21, 33, 34, 58, 75, 88, 89, 92, 93, 97, 100, 101, 106, 107, 108, 111, 139, 144, 182, 239, 240, 287 iron, 5, 6, 7, 8, 15, 21, 29, 33, 55, 57, 58, 70, 91, 92, 93, 94, 95, 96, 97, 98, 99, 100, 101, 103, 106, 107, 108, 109, 110, 111, 123, 163, 179, 223, 229, 230, 239, 240, 244, 251, 263, 272, 284, 285, 288, 296, 298, 299, 301, 302 iron transport, 103 irradiation, 22, 30, 88, 157, 239, 272 ischemia, 53, 169, 176, 183 isolation, 151, 195, 248 isomerization, 51, 58, 63, 72, 75, 79, 80, 89, 105, 126, 145, 165, 172 isomers, 125, 145, 153, 165, 172, 179, 190, 191, 243 isotope, 39, 105, 184, 198, 199 issues, 144 Italy, 180

J Jordan, 184

K ketones, 36 kidney, 156, 183, 266 kill, 8, 15, 272, 284 kinetic constants, 58 kinetic methods, 202, 234, 240, 249 kinetic model, 39 kinetics, 3, 7, 21, 26, 27, 28, 38, 39, 55, 69, 89, 102, 103, 108, 110, 171, 182, 249, 259, 262, 263, 271, 276, 278, 291, 292, 295, 298 Krebs cycle, 3

L L-arginine, 46, 67, 169, 231 LDL, 155 lead, 24, 34, 43, 46, 48, 56, 60, 61, 63, 65, 66, 79, 87, 95, 99, 101, 113, 114, 121, 124, 162, 177, 206, 207, 208, 209, 211, 224, 225, 233, 237, 239, 246, 249, 254, 255, 257, 258, 267, 270, 271, 288, 294 lens, 165, 170, 181 lesions, 63, 120, 177, 185, 190, 191, 192, 193, 194, 195, 196, 197, 198 leucine, 116 life cycle, 7 lifetime, 35, 95, 123, 201, 210, 219, 221, 225, 266, 274 ligand, 33, 55, 85, 92, 98, 151, 174, 178 light, 4, 6, 8, 14, 15, 22, 40, 161, 175, 203, 209, 211, 215, 216, 217, 227, 249, 252, 256, 258, 272, 289, 290, 291, 300 linear dependence, 215 linoleic acid, 40, 137, 145, 153, 157, 161, 164, 172, 174, 175, 176, 177, 178, 179, 181, 184 lipid oxidation, 50, 73, 132, 166, 169, 170, 171, 182 lipid peroxidation, 2, 26, 32, 35, 41, 48, 51, 61, 63, 72, 114, 118, 124, 138, 139, 140, 141, 142, 143, 145, 146, 147, 148, 149, 150, 151, 152, 153, 154, 155, 156, 157, 158, 159, 165, 166, 169, 170, 172, 181, 182, 249, 251, 263, 285 lipids, 12, 23, 25, 30, 32, 35, 36, 37, 44, 49, 65, 71, 89, 93, 132, 138, 139, 140, 144, 145, 149, 151, 154, 161, 162, 164, 165, 170, 173, 177, 179, 180, 181, 182, 207, 284 lipooxygenase, 165 lipoproteins, 61, 165, 170, 173, 174, 179, 180, 184

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Index liposomes, 39, 70, 149, 154, 156, 157 liquid chromatography, 198, 199, 218, 261 liquid phase, 227 liver, 100, 101, 107, 133, 141, 145, 152, 153, 154, 155, 156, 266, 272, 284, 293, 299 liver cancer, 284 local conditions, 288 localization, 134, 196, 262 longevity, 2, 152 low temperatures, 8 low-density lipoprotein, 141, 157 LTD, 199 lumen, 53, 129 luminescence, 40, 286, 287, 289, 290, 291, 294, 296, 297 lycopene, 36 lysine, 32, 117, 118, 132, 149, 158, 174, 191, 198 lysosome, 99, 129 lysozyme, 39, 88

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M machinery, 11, 129 macromolecules, 129, 139 macrophages, 43, 71, 94, 176, 177, 178, 183, 277, 281, 284, 299 magnesium, 4 magnetic field, 214, 215 magnetic moment, 215 magnetic resonance, 230 magnetic resonance imaging, 230 magnitude, 2, 7, 10, 11, 12, 32, 81, 98, 130, 195, 251, 254 majority, 11, 96, 116, 267, 287 mammalian cells, 43, 125, 129, 131, 144, 199, 266, 272 mammalian tissues, 46, 144 mammals, 13, 14, 23, 24, 153, 164, 165, 284 man, 41 management, 103, 298 manganese, 58, 59, 60, 92, 97, 132, 224, 261 mantle, 4 MAPK/ERK, 179 Marx, 103, 280 mass, 82, 103, 141, 144, 175, 181, 194, 197, 198, 199, 202, 218, 235 mass spectrometry, 82, 103, 141, 144, 175, 181, 194, 197, 198, 199, 218 materials, 203 matrix, 63, 72, 97, 144, 175 matter, 6, 244 measurement, 57, 70, 130, 135, 195, 196, 197, 198, 199, 200, 203, 210, 224, 227, 229, 231, 249,

311

259, 260, 281, 287, 289, 292, 293, 296, 299, 301 measurements, 40, 77, 85, 86, 132, 203, 207, 222, 230, 264, 288, 295 media, 26, 27, 28, 52, 154, 166, 267, 269, 270, 271, 272, 280 medical, 2, 213, 273 medicine, 14, 38, 41, 102, 132, 259, 289, 298 medulla, 266 meiosis, 9 melanin, 284 melatonin, 153, 155, 156 melting, 139 melting temperature, 139 membrane permeability, 149, 208, 221, 222 membranes, 4, 5, 6, 7, 10, 11, 15, 39, 47, 50, 53, 54, 61, 65, 67, 69, 70, 125, 138, 139, 141, 144, 146, 147, 148, 149, 150, 151, 152, 154, 157, 165, 166, 170, 173, 174, 180, 181, 251, 273, 286, 299 menadione, 221 messengers, 138, 145, 150, 151, 272, 285, 299 Metabolic, 175 metabolic disorder, 178 metabolic disorders, 178 metabolism, 3, 6, 16, 19, 23, 38, 46, 52, 66, 102, 103, 124, 130, 138, 154, 155, 156, 157, 178, 267, 272, 277, 278, 284, 285, 287, 298, 299, 301 metabolites, 20, 26, 39, 138, 141, 260, 284 metabolized, 141, 145, 175, 226 metabolizing, 287 metabolome, 183 metal complexes, 55, 58, 110, 224, 236, 237 metal ion, 21, 33, 92, 101, 108, 137, 154, 240, 244, 263, 288 metal ions, 21, 33, 92, 101, 108, 137, 154, 244, 288 metalloenzymes, 237, 240 metals, 91, 92, 101, 102, 110, 140, 167, 181, 284 metastasis, 111, 260 metazoa, 16 meter, 93 methodology, 184, 201 methyl group, 186, 187, 192 mice, 22 microcalorimetry, 88 microorganisms, 16, 272, 273, 277 microsomes, 139, 141, 152, 153, 155, 156 microwave radiation, 214 migration, 28, 122, 176, 177, 178 miniaturization, 231 miscarriage, 13

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mitochondria, 7, 8, 9, 10, 11, 12, 13, 14, 16, 52, 73, 107, 125, 128, 139, 140, 152, 153, 155, 156, 158, 179, 221, 257, 284 mitochondrial damage, 150 mitochondrial DNA, 2, 10, 16, 111, 260 mitogen, 178, 183 mitosis, 9 mixing, 96, 202, 210, 212, 294 model system, 105, 132, 149 models, 56, 80, 100, 276, 298 modifications, 36, 46, 49, 61, 63, 64, 65, 66, 114, 117, 121, 124, 130, 132, 144, 148, 149, 154, 207, 235, 285 molar ratios, 93 mole, 289 molecular biology, 208, 289 molecular mass, 125, 134 molecular oxygen, 7, 19, 20, 21, 32, 40, 41, 43, 48, 58, 59, 80, 82, 83, 89, 140, 186, 190, 192, 218, 221, 225, 227, 284, 287, 290, 295 molecular weight, 33, 36, 60, 66, 92, 107, 126, 163, 173, 208 molecules, 3, 21, 22, 23, 30, 33, 35, 39, 40, 54, 93, 98, 116, 119, 122, 124, 126, 130, 137, 138, 141, 142, 143, 144, 149, 161, 163, 164, 165, 171, 176, 177, 178, 179, 210, 213, 216, 225, 238, 240, 245, 249, 252, 257, 280, 285, 291, 298 monoclonal antibody, 199 monohydrogen, 97, 98, 99, 100, 101 monolayer, 296, 297, 298 monomers, 126 morphology, 6 mosaic, 11, 13, 14 motif, 134 MRI, 230 mRNA, 179, 299 mRNAs, 299 mtDNA, 10, 11, 12, 13, 14, 15, 16 multicellular organisms, 8, 267 mutation, 2, 12, 13, 113 mutation rate, 2, 12, 13 mutations, 10, 12, 111, 260, 267 myelin, 157 myelodysplasia, 92 myocardial infarction, 176, 267 myoglobin, 33

N NAD, 98, 206, 231, 245, 257, 258, 260 NADH, 16, 98, 150, 158, 238, 258, 260, 263 nanometer, 286 nanometers, 28

natural selection, 4 nerve, 176 neurodegeneration, 43, 60 neurodegenerative diseases, 150, 155 neurodegenerative disorders, 147 neurological disease, 146 neuronal apoptosis, 157 neurotoxicity, 90, 103 neurotransmission, 46, 169 neutral, 12, 47, 54, 65, 97, 98, 100, 101, 105, 127, 165, 191, 210, 219, 221, 232, 237, 238, 239, 246 neutrophils, 2, 15, 40, 43, 94, 176, 177, 284 next generation, 11 NH2, 71 nicotinamide, 206, 245 nitrates, 63, 68, 162 nitric oxide, 2, 20, 21, 24, 38, 43, 44, 45, 46, 48, 57, 60, 62, 65, 66, 67, 68, 69, 70, 71, 72, 73, 74, 91, 93, 104, 105, 109, 132, 161, 162, 163, 169, 176, 177, 181, 182, 183, 184, 225, 226, 231, 260, 261, 262, 273, 281, 283, 299 nitric oxide synthase, 24, 46, 65, 93, 104, 163, 169, 176, 177, 183, 226 nitrite, 3, 39, 43, 44, 45, 48, 49, 50, 57, 58, 61, 62, 67, 68, 69, 70, 71, 95, 105, 122, 162, 172, 173, 182, 226, 227, 229, 230, 231, 232, 262 nitrogen, 1, 20, 24, 26, 28, 37, 39, 43, 44, 45, 46, 47, 49, 50, 51, 52, 53, 55, 58, 59, 62, 65, 66, 67, 70, 72, 73, 74, 82, 91, 93, 105, 106, 109, 121, 122, 124, 132, 135, 161, 162, 163, 173, 180, 181, 182, 207, 213, 215, 225, 227, 228, 232, 247, 248, 260, 262, 289 nitrogen compounds, 46 nitrogen dioxide, 24, 28, 43, 45, 49, 50, 52, 58, 62, 65, 70, 72, 73, 74, 93, 122, 161, 162, 180, 181, 182, 213, 225, 227, 228, 232, 248, 262 nitrosamines, 65, 73 nitroso compounds, 204 nitrous oxide, 45, 211, 225 nitroxide, 204, 224, 242, 243 NK cells, 134 NMR, 80 Nobel Prize, 3 NRP, 137, 142 nuclear receptors, 159 nuclei, 139, 153, 215 nucleic acid, 31, 32, 35, 36, 49, 93, 95, 139 nucleophiles, 37, 163, 174 nucleotides, 31, 40, 80, 95, 106 nucleus, 9, 11, 125 nutrient, 7, 19 nutrients, 8, 19

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O obesity, 164 oceans, 5, 10 old age, 14 oleic acid, 137, 139, 142, 164, 165, 175, 178, 179, 180, 183, 184 olive oil, 182 oncogenes, 151, 267 oocyte, 13, 14 optimization, 196 organelles, 11, 16, 129, 295 organic compounds, 30, 93, 96, 246 organic matter, 4, 208, 244 organic peroxides, 97, 98 organism, 8, 94, 97, 176, 272 organs, 102, 125, 153, 266, 298 osteoarthritis, 63 overproduction, 56, 60, 162 ox, 3, 79, 85, 90, 121, 220 oxidation products, 36, 62, 65, 114, 118, 141, 144, 145, 146, 151, 152, 170, 187, 197, 223, 231, 235, 240, 250 oxidative damage, 15, 34, 68, 99, 101, 113, 123, 130, 139, 146, 150, 161, 173, 199, 260, 263 oxidative stress, 1, 3, 4, 6, 8, 11, 12, 14, 23, 24, 38, 47, 60, 61, 80, 87, 90, 92, 93, 96, 99, 101, 102, 103, 110, 131, 134, 138, 139, 143, 145, 149, 150, 151, 152, 154, 156, 157, 158, 161, 162, 170, 185, 190, 193, 197, 198, 262, 263, 264, 272, 273, 278, 281, 285, 299, 300, 301, 302 oxidizability, 153 oxygen consumption, 14, 15, 149, 171, 249, 270 oxyhemoglobin, 54, 56, 69, 92, 95, 102, 104, 227, 229 ozone, 30, 32, 41, 227 ozone reaction, 41 ozonolysis, 41

P parallel, 7, 77, 86, 186, 292 parasite, 10 parasites, 9 partial differential equations, 270 partition, 47, 165, 182 pathogenesis, 143, 176 pathogens, 23 pathology, 67, 131

313

pathophysiological, 2, 20, 24, 49, 70, 143, 147, 150, 152, 153, 222 pathophysiology, 23, 38, 50, 66, 93, 104, 162, 182 pathways, 3, 20, 21, 25, 52, 60, 61, 62, 77, 82, 84, 114, 117, 150, 151, 152, 161, 162, 166, 167, 169, 170, 171, 186, 190, 197, 204, 208, 220, 236, 250, 251, 253, 255, 256, 257, 272, 284, 287, 290, 292, 300 peptide, 71, 81, 84, 85, 89, 90, 93, 103, 123, 127, 129, 131, 132, 191, 198 peptides, 16, 34, 40, 71, 75, 76, 80, 81, 84, 85, 86, 88, 89, 90, 131, 133 perfusion, 294 permeability, 139, 140, 145, 149, 155, 157, 176, 208, 224 peroxidation, 35, 39, 63, 72, 138, 139, 140, 141, 142, 143, 145, 149, 150, 151, 152, 153, 154, 155, 156, 157, 158, 169, 170, 193, 249, 263 peroxide, 33, 50, 51, 65, 102, 107, 110, 153, 274, 298, 299 peroxynitrite, 24, 26, 27, 28, 29, 30, 31, 35, 38, 39, 43, 44, 45, 46, 48, 49, 51, 54, 56, 58, 59, 60, 61, 62, 63, 65, 67, 68, 69, 70, 71, 72, 73, 74, 91, 92, 103, 104, 105, 106, 109, 121, 122, 132, 162, 166, 167, 169, 170, 171, 172, 181, 182, 224, 232, 233, 237, 260, 262, 263 personal communication, 11 phagocyte, 23, 104, 300 phagocytosis, 33, 34, 35, 40, 129, 277, 278 pharmacology, 38, 66, 104 phenol, 235 phenolic compounds, 205, 233, 251 phenotype, 176, 177 phenoxyl radicals, 32 phenylalanine, 61, 62, 72, 105, 116, 240 Philadelphia, 39 PhOH, 235, 251 phosphate, 3, 5, 97, 101, 273, 291, 294 phosphatidylcholine, 138, 157 phosphatidylethanolamine, 138, 144, 157 phosphatidylserine, 138, 144, 154 phospholipids, 72, 138, 139, 140, 141, 142, 143, 144, 145, 147, 148, 149, 151, 152, 154, 157, 164, 165, 176 phosphorylation, 1, 3, 10, 12, 60, 71, 95, 105, 122, 177, 181, 226 photoemission, 153 photolysis, 68, 77, 89, 202, 210, 211, 226, 234, 237, 248, 262 photons, 227, 301 photosynthesis, 3, 6, 8, 15, 19 phylogenetic tree, 4 physical interaction, 209, 216

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314 physical properties, 47, 69, 144, 149 physicochemical characteristics, 208 physicochemical properties, 138, 142, 173, 259 Physiological, 40, 66, 103, 138, 294 physiology, 7, 38, 66 Planck constant, 214 plants, 6, 8, 9, 12, 20, 125, 128, 138, 139, 154, 284 plaque, 41 plasma membrane, 281 plasma proteins, 173 plasmid, 289 plasticity, 43, 157 platelet aggregation, 43, 46 platelets, 105, 144, 151, 156, 176, 177 platinum, 93 playing, 176 polar, 144, 173, 174 polarity, 5, 144 polymer, 63 polymerization, 22 polymerization process, 22 polymerization processes, 22 polymorphism, 154 polymorphisms, 12 polypeptide, 30, 122, 123 polypeptides, 124, 125, 127, 131, 156 polyphenols, 69, 153, 208, 251 polyphosphates, 5 polyploid, 11 polyploidy, 11, 16 polystyrene, 297 polyunsaturated fat, 39, 80, 89, 137, 139, 145, 146, 153, 154, 155, 156, 164, 180 polyunsaturated fatty acids, 39, 80, 89, 137, 139, 145, 146, 153, 154, 155, 156, 164, 180 POOH, 206 population, 13, 14 porphyrins, 35, 224, 236 potassium, 106, 219, 227, 289 PPAR ligands, 151, 159 preparation, 144, 193, 195, 246, 248, 261 principles, 39, 134, 213, 235, 281 probability, 12, 53, 130, 189, 256 probe, 111, 201, 202, 203, 204, 205, 206, 208, 209, 213, 223, 226, 229, 230, 233, 235, 238, 240, 242, 243, 244, 250, 251, 252, 253, 255, 257, 258, 259, 262, 263, 300 producers, 150 pro-inflammatory, 161, 176, 177, 180 project, 195 prokaryotes, 6, 7, 9, 10, 11, 125 prokaryotic cell, 6

Index proliferation, 2, 143, 150, 151, 159, 165, 180, 285, 298 proline, 116, 117, 267 promoter, 105 propagation, 32, 48, 132, 139, 142, 147, 154, 170, 171, 172 protection, 73, 152, 155, 182 protective role, 99 protein components, 131 protein constituent, 163 protein kinase C, 150 protein kinases, 208, 285 protein oxidation, 89, 114, 123, 124, 129, 130, 131 protein structure, 55, 60, 76, 113, 124, 125, 132, 281 proteinase, 62, 72, 133, 158 proteolysis, 124, 133 proteolytic enzyme, 2 protons, 3, 4, 6, 82 pumps, 218, 294 purification, 134 pyrimidine, 186, 188, 189, 193 pyrophosphate, 5

Q quantification, 70, 130, 141, 144, 146, 179, 180, 201, 202, 204, 205, 213, 218, 222, 235, 243, 255 quantum mechanics, 21 quinones, 35

R race, 88 radiation, 23, 30, 31, 39, 77, 80, 131, 157, 186, 191, 192, 198, 199, 210, 211, 213, 214, 215, 239, 261, 267 Radiation, 88, 89, 212, 213 radical formation, 72, 73, 88, 90, 105, 109, 115, 203, 208, 250, 251, 267 radical reactions, 26, 29, 39, 90, 108, 110, 132, 139, 162, 197, 202 radiotherapy, 30, 213 radius, 28, 165 reactant, 122, 210, 212 reactants, 79, 83, 98, 206, 212, 225, 290 reaction mechanism, 141, 201 reaction rate, 47, 54, 106, 114, 116, 121, 123 reactive oxygen, 1, 15, 20, 23, 38, 60, 67, 70, 71, 82, 91, 138, 139, 153, 157, 158, 162, 198, 260, 283, 284, 298, 300, 301 reactive sites, 171

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Index reactivity, 7, 20, 26, 27, 29, 32, 33, 34, 35, 36, 47, 51, 54, 56, 58, 71, 89, 91, 92, 100, 101, 107, 109, 138, 142, 165, 172, 178, 208, 210, 212, 232, 235, 240, 242, 254, 255, 257, 258, 259, 262, 263, 272, 301 reagents, 26, 96, 101, 109, 165, 239, 259, 263, 292, 294 real time, 202, 234, 286, 293 reality, 13, 101, 205, 242 reasoning, 37 receptors, 113, 129, 145, 151, 159, 285 recognition, 107, 127, 133 recombination, 50, 59, 205, 235, 247, 259 recommendations, 38, 102, 265 recycling, 8 red blood cells, 54, 57, 100, 175, 179 red wine, 69 reducing sugars, 118 regression, 295 regression analysis, 295 rejection, 132, 179 relatives, 9 relaxation, 24, 69, 176, 217, 252 relevance, 2, 27, 52, 53, 75, 87, 89, 90, 154, 161, 265 reliability, 130 relief, 7 repair, 24, 68, 86, 89, 90, 108, 120, 124, 125, 130, 192, 197, 198, 207 repression, 151 requirements, 98, 138, 178, 204, 207, 209 researchers, 14, 151, 280 residues, 31, 34, 62, 68, 72, 76, 82, 89, 90, 95, 114, 116, 118, 119, 120, 122, 123, 124, 125, 132, 133, 144, 149, 158, 162, 163, 165, 172, 174, 178, 198, 235, 267, 285 resistance, 2, 30 resolution, 1, 133, 176, 178, 180, 288 respiration, 2, 5, 6, 7, 8, 9, 10, 11, 14, 16, 21, 22, 23, 57, 66, 91, 150, 158, 267, 298 respiratory rate, 2 response, 8, 10, 30, 46, 73, 131, 146, 150, 151, 169, 173, 176, 177, 178, 179, 184, 226, 227, 267, 271, 279, 285, 288 restrictions, 185 reticulum, 9, 125, 128, 134, 157 retina, 144, 145, 154 retinol, 157 rheumatoid arthritis, 265 ribonucleotide reductase, 80 rings, 126, 127 risk, 12, 70, 195 RNA, 30 ROOH, 27, 31, 35, 36, 245, 248, 250, 251

roots, 7 routes, 54, 60, 162, 166, 169, 170, 171 Royal Society, 102

S salt concentration, 270 salts, 96, 106 saturated fat, 141, 144 saturated fatty acids, 141 saturation, 175, 183 scattering, 203 scavengers, 100, 101, 107, 123, 133, 207, 208, 209, 219, 236, 238, 239, 240, 241, 242, 244, 250, 252, 255 scope, 5, 10 sea level, 266 secretion, 177, 180, 207 selectivity, 89, 90, 208, 210, 218, 240, 244, 258 selenium, 156 self-fulfilling prophesy, 260 senescence, 2 senses, 1 sensing, 273 sensitivity, 14, 139, 143, 185, 195, 203, 213, 216, 218, 231, 283, 286, 287, 288, 291, 293 sensitization, 186 sensors, 30, 151, 231, 288 sepsis, 56, 179 serine, 116, 191 serotonin, 156 serum, 62, 71, 92, 98, 103, 200, 297 serum albumin, 62, 71 sex, 9, 12 sexual development, 2 sexual reproduction, 13 shape, 144 shelter, 176 shock, 125 showing, 195 sickle cell, 92, 103 sickle cell anemia, 92 side chain, 76, 117, 118, 124, 146, 149 signal transduction, 21, 25, 55, 70, 113, 126, 143, 150, 151, 174, 181, 281, 285, 299 signaling circuits, 25 signaling pathway, 20, 24, 25, 153, 165, 179, 273, 296 signalling, 2, 39, 158, 298 signals, 1, 2, 14, 38, 151, 176, 242, 279, 288 skin, 30, 35 skin cancer, 30 smooth muscle, 57, 69, 176, 178, 180

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316

Index

smooth muscle cells, 57, 178 SO42-, 19 sodium, 220, 225, 229, 277, 288, 301, 302 sodium hydroxide, 225, 229 software, 294, 297 solid state, 76 solubility, 54, 172, 208, 219, 237, 267 solution, 4, 10, 39, 41, 67, 79, 85, 88, 89, 90, 92, 93, 96, 97, 98, 101, 103, 104, 106, 109, 132, 174, 187, 188, 189, 210, 211, 212, 216, 218, 219, 225, 226, 227, 229, 231, 232, 238, 239, 246, 248, 249, 250, 258, 262, 263, 287, 290, 291, 292, 293, 294, 297, 302 solvent molecules, 210 solvents, 219 speciation, 13, 14, 17, 92, 101, 103 spectrophotometric method, 202 spectrophotometry, 108, 175, 202, 215, 216, 222, 233, 234, 240 spectroscopic techniques, 241 spectroscopy, 57, 85, 173, 202, 213, 216, 230, 231, 260, 261 spin, 7, 21, 22, 38, 47, 67, 108, 109, 150, 158, 173, 204, 213, 214, 215, 221, 222, 229, 230, 237, 238, 241, 242, 243, 245, 250, 260, 261, 263, 264 stability, 88, 125, 173, 180, 197, 222, 225, 234, 243, 249, 273, 284 stabilization, 80, 267, 278 stable radicals, 22 starvation, 129 state, 1, 7, 12, 19, 20, 21, 33, 35, 45, 55, 58, 76, 79, 81, 88, 98, 99, 109, 126, 133, 150, 172, 176, 178, 189, 202, 203, 204, 206, 210, 211, 212, 213, 215, 216, 217, 219, 221, 223, 224, 226, 227, 234, 235, 239, 252, 256, 265, 268, 269, 270, 275, 276, 277, 278, 279, 280, 285, 295, 296 steroids, 164 sterols, 39 stimulus, 8, 177 stoichiometry, 4, 8, 141, 171, 225, 234, 268, 276, 289 stomach, 49, 68, 69 storage, 60, 91, 92, 103, 129, 164, 219, 225, 285 stress, 1, 2, 8, 14, 15, 26, 66, 88, 101, 110, 125, 130, 131, 139, 143, 147, 149, 150, 153, 154, 158, 159, 162, 176, 183, 260, 273, 278, 288, 299 stroke, 267 structural modifications, 149, 182 structure, 5, 6, 7, 9, 21, 46, 49, 51, 63, 67, 72, 75, 76, 83, 84, 85, 86, 89, 90, 96, 99, 110, 125, 126,

127, 128, 132, 138, 143, 144, 157, 159, 163, 164, 175, 208, 209, 215, 222, 225 substitutes, 162 substitution, 12, 76, 77, 120, 169, 171 substitution reaction, 120 substrate, 2, 3, 4, 10, 33, 94, 100, 102, 126, 127, 128, 129, 133, 134, 141, 152, 166, 212, 213, 236, 245, 246, 247, 248, 258, 270, 284, 294 substrates, 40, 106, 109, 124, 127, 129, 133, 134, 147, 221, 236, 251, 286, 287, 292 subtraction, 227 succession, 11 suicide, 16 sulfate, 19, 63, 72, 89, 106 sulfur, 5, 6, 26, 29, 37, 55, 57, 58, 75, 79, 84, 88, 119, 123, 163, 224, 240 Sun, 154, 262 supplementation, 180 suppression, 176, 179, 236 surface area, 10 survival, 13, 151 susceptibility, 2, 12, 14, 119 suspensions, 298 Switzerland, 91 symmetry, 55 synthesis, 3, 4, 6, 13, 45, 52, 70, 80, 104, 124, 145, 161, 162, 169, 171, 176, 179, 181, 184, 261, 272, 279, 284, 285 systemic change, 267

T T cell, 71 tanks, 225, 268 target, 27, 28, 31, 54, 81, 83, 100, 153, 175, 178, 189, 210, 211, 212, 239, 261, 278, 279, 285 techniques, 101, 153, 195, 199, 202, 207, 208, 210, 211, 212, 213, 221, 223, 230, 232, 235, 237, 267, 287, 292 technology, 146 TEG, 105 temperature, 231, 248, 270, 292, 293, 294 tempo, 12 tension, 2, 10, 155, 168, 171 tensions, 2, 57, 59 testing, 144 testis, 153, 155 tetrahydrofuran, 145, 155 textiles, 272 thalassemia, 92, 103 thallium, 104 therapeutics, 38, 182, 261 therapy, 35, 100, 281

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Index thermal decomposition, 219, 247 thermodynamics, 21 threonine, 116, 117 thrombin, 177 thromboxanes, 177 thymine, 186, 187, 198 thymus, 200, 266 thyroid, 2, 272, 284 time frame, 9, 92 tissue, 23, 24, 28, 33, 37, 41, 44, 47, 53, 54, 77, 79, 83, 95, 98, 100, 133, 134, 138, 139, 144, 153, 169, 171, 176, 179, 251, 266, 273, 277, 279, 280, 296, 301 tocopherols, 251 toxic effect, 21, 22 toxicity, 2, 10, 23, 36, 39, 47, 93, 97, 101, 103, 108, 243, 272, 285 toxin, 272 TPA, 41 trade, 6, 14 trade-off, 14 trafficking, 174, 175, 178 training, 2 traits, 9 transcription, 1, 10, 30, 60, 126, 138, 151, 176, 178, 183, 267, 273, 279, 281, 285 transcription factors, 1, 138, 151, 176, 178, 273, 285 transducer, 183 transduction, 2, 182, 183, 184, 285 transference, 58 transferrin, 91, 92, 103, 285 transformation, 141, 263 transformations, 93 transfusion, 103 transition metal, 5, 21, 22, 23, 29, 30, 31, 32, 33, 36, 43, 44, 47, 51, 58, 61, 65, 92, 94, 96, 122, 142, 162, 272, 284, 294 transition metal ions, 21, 22, 23, 29, 32, 33, 36, 94, 96, 142, 284 translation, 179 translocation, 7, 127 transplant, 179 transport, 6, 7, 24, 29, 52, 57, 91, 113 treatment, 123, 178, 249, 251 triggers, 21 triglycerides, 139, 153 trypsin, 127 tryptophan, 61, 62, 63, 71, 72, 123, 162 tumor, 30, 35, 111, 260, 267, 285 tumor cells, 30, 285 tumor development, 30 tumors, 267 turnover, 31, 35, 36, 41, 92, 128, 156, 270, 287

tyrosine, 50, 51, 53, 58, 60, 61, 62, 63, 66, 67, 68, 69, 70, 71, 72, 95, 105, 106, 116, 120, 122, 132, 162, 163, 172, 181, 233, 235, 238, 240, 254, 258, 273 Tyrosine, 60, 62, 70, 71, 72, 95, 122

U United, 4 United Nations, 4 uric acid, 31, 65, 70, 221 urine, 177, 179, 183 Uruguay, 43, 66, 161, 180 USA, 16, 75, 107, 109, 110, 155, 156, 158, 198, 201, 261, 299 UV light, 239, 272 UV radiation, 197

V vacuole, 11 valence, 47 validation, 277 valine, 116 vanadium, 231 variables, 251 variations, 196 vascular system, 178 vascular wall, 176, 177 vasculature, 54, 69, 162, 178, 180 vasoconstriction, 145 vasodilation, 25, 57, 93, 161, 169 vasodilator, 177, 181 vasomotor, 181 VCAM, 178 vegetables, 53 vesicle, 129 vessels, 54, 155, 268, 269, 280 viral infection, 8 viruses, 8 vitamin E, 101, 155, 170, 251 vitamins, 23, 164

W waste, 6, 19, 129, 151 water, 2, 5, 6, 19, 30, 44, 47, 49, 53, 54, 84, 86, 91, 93, 98, 99, 100, 102, 104, 120, 164, 186, 188, 191, 210, 218, 228, 231, 237, 238, 242, 247, 263, 267, 270, 272, 274, 284, 286, 294, 297 wavelengths, 217, 288 wells, 272

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western blot, 289 Western blot, 279 white blood cells, 273, 277, 284 wires, 3, 7 Wisconsin, 201 workers, 22, 96, 192 wound healing, 283, 288

X

Y yeast, 12, 38, 59, 125, 133 yield, 20, 35, 44, 48, 49, 52, 54, 56, 57, 58, 59, 60, 61, 62, 63, 65, 76, 78, 86, 95, 105, 110, 114, 116, 118, 120, 122, 142, 162, 171, 192, 193, 202, 203, 204, 205, 206, 209, 210, 211, 213, 217, 218, 219, 221, 223, 234, 235, 237, 239, 244, 246, 247, 249, 251, 253, 255, 256, 258, 259, 289, 290

X-irradiation, 38 X-ray diffraction, 157

Z

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zinc, 55, 59, 72, 73, 93, 96, 103, 107

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